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 border="0" cellpadding="0" cellspacing="0" style="width:793px;" width="794">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Glass pasteur pipettes</td>
			<td style="width:200px;">VWR</td>
			<td style="width:211px;">14673-043</td>
			<td style="width:167px;"> </td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Inverted microscope Axiovert 40C</td>
			<td style="width:200px;">Zeiss</td>
			<td style="width:211px;">4109431007990000</td>
			<td>Injection microscope</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Microloader tips</td>
			<td style="width:200px;">Eppendorf</td>
			<td style="width:211px;">5242 956.003</td>
			<td style="width:167px;">Load injection solution</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Nylon filter mesh 80 &#956;m</td>
			<td style="width:200px;">Amazon.com</td>
			<td style="width:211px;">03-80-37</td>
			<td style="width:167px;">Filter eggs to get rid of debris</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">P20 or P200 Aerosol Barrier Pipette Tips</td>
			<td style="width:200px;">Fisher Scientific</td>
			<td style="width:211px;">02707432 or 02707430</td>
			<td style="width:167px;">Part of a mouth pipette</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Parafilm</td>
			<td style="width:200px;">Fisher Scientific</td>
			<td style="width:211px;">13 374 12</td>
			<td style="width:167px;">Part of a mouth pipette</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Polyethylene tubing</td>
			<td style="width:200px;">Intramedic</td>
			<td style="width:211px;">PE-160</td>
			<td style="width:167px;">Part of a mouth pipette</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Protamine sulfate</td>
			<td style="width:200px;">MP Biomedicals, LLC</td>
			<td style="width:211px;">194729</td>
			<td style="width:167px;">Attach dejellied eggs to injection dishes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sea urchins S. purpuratus</td>
			<td>Pt. Loma Marine Invertebrate Lab</td>
			<td style="width:211px;">N/A</td>
			<td style="width:167px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Sea water</td>
			<td style="width:200px;">any pet store</td>
			<td style="width:211px;">Instant Ocean</td>
			<td style="width:167px;"> </td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Sterile 60 mm x 15 mm Polystyrene Petri Dish</td>
			<td style="width:200px;">Fisher Scientific</td>
			<td style="width:211px;">0875713A</td>
			<td style="width:167px;">Injection dishes</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:216px;">Three-Axis Coarse Positioning Micromanipulator MMN-1</td>
			<td style="width:200px;">Narishige</td>
			<td style="width:211px;">9124</td>
			<td>Manipulate injection needle</td>
		</tr>
		<tr height="84">
			<td height="84" style="height:84px;width:216px;">Three-Axis Joystick Type Oil Hydraulic Fine Micromanipulator MMO-202ND</td>
			<td style="width:200px;">Narishige</td>
			<td style="width:211px;">9212</td>
			<td>Manipulate injection needle</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Transfer pipettes</td>
			<td style="width:200px;">Fisher Scientific</td>
			<td style="width:211px;">13-711-9AM</td>
			<td style="width:167px;"> </td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:216px;">Vertical needle puller </td>
			<td style="width:200px;">Narishige</td>
			<td style="width:211px;">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...

Watch this Scientific Journal Video about High Throughput Microinjections of Sea Urchin Zygotes at JoVE.com

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

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Biology

14.4K 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

High-throughput Yeast Plasmid Overexpression Screen

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, including those with direct relevance to human disease. Because of its short generation time and well-characterized genome, a major experimental advantage of the yeast model system is the ability to perform genetic screens to identify genes and pathways that are involved in a given process. Over the last thirty years such genetic screens have been used to elucidate the cell cycle, secretory pathway, and many more highly conserved aspects of eukaryotic cell biology 1-5. In the last few years, several genomewide libraries of yeast strains and plasmids have been generated 6-10. These collections now allow for the systematic interrogation of gene function using gain- and loss-of-function approaches 11-16. Here we provide a detailed protocol for the use of a high-throughput yeast transformation protocol with a liquid handling robot to perform a plasmid overexpression screen, using an arrayed library of 5,500 yeast plasmids. We have been using these screens to identify genetic modifiers of toxicity associated with the accumulation of aggregation-prone human neurodegenerative disease proteins. The methods presented here are readily adaptable to the study of other cellular phenotypes of interest.

Here we describe a plasmid overexpression screen in Saccharomyces cerevisiae, using an arrayed plasmid library and a high-throughput yeast transformation protocol with a liquid handling robot.

The budding yeast, Saccharomyces cerevisiae, is a powerful model system for defining fundamental mechanisms of many important cellular processes, ...

High-throughput Saccharification Assay for Lignocellulosic Materials

Polysaccharides that make up plant lignocellulosic biomass can be broken down to produce a range of sugars that subsequently can be 
used in establishing a biorefinery. These raw materials would constitute a new industrial platform, which is both sustainable and carbon neutral, to replace 
the current dependency on fossil fuel. The recalcitrance to deconstruction observed in lignocellulosic materials is produced by several intrinsic properties 
of plant cell walls. Crystalline cellulose is embedded in matrix polysaccharides such as xylans and arabinoxylans, and the whole structure is encased by the 
phenolic polymer lignin, that is also difficult to digest 1. In order to improve the digestibility of plant materials we need to discover the main bottlenecks 
for the saccharification of cell walls and also screen mutant and breeding populations to evaluate the variability in saccharification 2. These tasks require 
a high throughput approach and here we present an analytical platform that can perform saccharification analysis in a 96-well plate format. This platform has 
been developed to allow the screening of lignocellulose digestibility of large populations from varied plant species. We have scaled down the reaction volumes 
for gentle pretreatment, partial enzymatic hydrolysis and sugar determination, to allow large numbers to be assessed rapidly in an automated system.
This automated platform works with milligram amounts of biomass, performing ball milling under controlled conditions to reduce the plant 
materials to a standardised particle size in a reproducible manner. Once the samples are ground, the automated formatting robot dispenses specified and recorded 
amounts of material into the corresponding wells of 96 deep well plate (Figure 1). Normally, we dispense the same material into 4 wells to have 4 replicates for 
analysis. Once the plates are filled with the plant material in the desired layout, they are manually moved to a liquid handling station (Figure 2). In this 
station the samples are subjected to a mild pretreatment with either dilute acid or alkaline and incubated at temperatures of up to 90&deg;C. The pretreatment solution 
is subsequently removed and the samples are rinsed with buffer to return them to a suitable pH for hydrolysis. The samples are then incubated with an enzyme
mixture for a variable length of time at 50&deg;C. An aliquot is taken from the hydrolyzate and the reducing sugars are automatically determined by the MBTH 
colorimetric method.

A simple, rapid method for determining the saccharification potential of large numbers of plant biomass samples is described. The automated platform for this analysis involves the preparation of the plant biomass for analysis in 96 well plates and the subsequent performance of pretreatment, hydrolysis and quantification of the sugars released.

Polysaccharides that make up plant lignocellulosic biomass can be broken down to produce a range of sugars that subsequently can be 
used in establishing ...

High-throughput Purification of Affinity-tagged Recombinant Proteins

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further biochemical analyses to obtain detailed insights into structure/function relationships. Advances in oligonucleotide- and gene synthesis technology make large-scale mutagenesis strategies increasingly feasible, including the substitution of target residues by all 19 other amino acids. Gain- or loss-of-function phenotypes then allow systematic conclusions to be drawn, such as the contribution of particular residues to catalytic activity, protein stability and/or protein-protein interaction specificity.
In order to attribute the different phenotypes to the nature of the mutation - rather than to fluctuating experimental conditions - it is vital to purify and analyse the proteins in a controlled and reproducible manner. High-throughput strategies and the automation of manual protocols on robotic liquid-handling platforms have created opportunities to perform such complex molecular biological procedures with little human intervention and minimal error rates1-5.
Here, we present a general method for the purification of His-tagged recombinant proteins in a high-throughput manner. In a recent study, we applied this method to a detailed structure-function investigation of TFIIB, a component of the basal transcription machinery. TFIIB is indispensable for promoter-directed transcription in vitro and is essential for the recruitment of RNA polymerase into a preinitiation complex6-8. TFIIB contains a flexible linker domain that penetrates the active site cleft of RNA polymerase9-11. This linker domain confers two biochemically quantifiable activities on TFIIB, namely (i) the stimulation of the catalytic activity during the 'abortive' stage of transcript initiation, and (ii) an additional contribution to the specific recruitment of RNA polymerase into the preinitiation complex4,5,12 . We exploited the high-throughput purification method to generate single, double and triple substitution and deletions mutations within the TFIIB linker and to subsequently analyse them in functional assays for their stimulation effect on the catalytic activity of RNA polymerase4. Altogether, we generated, purified and analysed 381 mutants - a task which would have been time-consuming and laborious to perform manually. We produced and assayed the proteins in multiplicates which allowed us to appreciate any experimental variations and gave us a clear idea of the reproducibility of our results.
This method serves as a generic protocol for the purification of His-tagged proteins and has been successfully used to purify other recombinant proteins. It is currently optimised for the purification of 24 proteins but can be adapted to purify up to 96 proteins.

We describe a method for the affinity-tagged purification of recombinant proteins using liquid-handling robotics. This method is generally applicable to the small-scale purification of soluble His-tagged proteins in a high-throughput format.

X-ray crystallography is the method of choice for obtaining a detailed view of the structure of proteins. Such studies need to be complemented by further ...

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 ...

High-throughput Titration of Luciferase-expressing Recombinant Viruses

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with viruses that do not form plaques. As an alternative to plaque assays, we have developed a high-throughput titration method that allows for the simultaneous titration of a high volume of samples in a single day. This approach involves infection of the samples with a Firefly luciferase tagged virus, transfer of the infected samples onto an appropriate permissive cell line, subsequent addition of luciferin, reading of plates in order to obtain luminescence readings, and finally the conversion from luminescence to viral titers. The assessment of cytotoxicity using a metabolic viability dye can be easily incorporated in the workflow in parallel and provide valuable information in the context of a drug screen. This technique provides a reliable, high-throughput method to determine viral titers as an alternative to a standard plaque assay.

This article presents a high-throughput luciferase expression-based method of titrating various RNA and DNA viruses using automated and manual liquid handlers.

Standard plaque assays to determine infectious viral titers can be time consuming, are not amenable to a high volume of samples, and cannot be done with ...

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 ...

Recovery of Adult Zebrafish Hearts for High-throughput Applications

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and purification of RNA, DNA, and proteins. All of these applications demand the rapid recovery of significant numbers of zebrafish hearts to avoid gene regulatory, metabolic, and other changes that begin after death. Adult zebrafish hearts are also required for studying heart structure for a variety of mutants and for studying heart regeneration. However, the traditional zebrafish heart dissection is slow and difficult and requires specialized tools, making large-scale dissection of adult zebrafish hearts tedious. Traditional methods also harbor the risk of damaging the heart during the dissection. Here, we describe a method for dissection of adult zebrafish hearts that is fast, reproducible, and preserves heart architecture. Furthermore, this method does not require specialized tools, is painless for the zebrafish, can be performed on fresh or fixed specimens, and can be performed on zebrafish as young as one month old. The approach described expands the use of adult zebrafish for cardiovascular research.

Use of zebrafish for cardiovascular research is expanding towards research on adult hearts. For these applications, quick and simple isolation of cardiac tissues is key to avoid post-mortem changes and to obtain an adequate number of samples. Here, we describe a fast and reproducible method for dissecting adult zebrafish hearts.

Use of the zebrafish model system for studying development, regeneration, and disease is expanding toward use of adult hearts for cell dissociation and ...

High-throughput Screening for Chemical Modulators of Post-transcriptionally Regulated Genes

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays focus on transcriptional regulation, being essentially aimed at the identification of promoter-targeting molecules. As a result, post-transcriptional mechanisms are largely uncovered by gene expression targeted drug development. Here we describe a cell-based assay aimed at investigating the role of the 3&#39; untranslated region (3&#8217; UTR) in the modulation of the fate of its mRNA, and at identifying compounds able to modify it. The assay is based on the use of a luciferase reporter construct containing the 3&#8217; UTR of a gene of interest stably integrated into a disease-relevant cell line. The protocol is divided into two parts, with the initial focus on the primary screening aimed at the identification of molecules affecting luciferase activity after 24 hr of treatment. The second part of the protocol describes the counter-screening necessary to discriminate compounds modulating luciferase activity specifically through the 3&#8217; UTR. In addition to the detailed protocol and representative results, we provide important considerations about the assay development and the validation of the hit(s) on the endogenous target. The described cell-based reporter gene assay will allow scientists to identify molecules modulating protein levels via post-transcriptional mechanisms dependent on a 3&#8217; UTR.

Here we describe a cell-based reporter gene assay as a valuable tool to screen chemical libraries for compounds modulating post-transcriptional control mechanisms exerted through 3&#8217; UTR.

Both transcriptional and post-transcriptional regulation have a profound impact on genes expression. However, commonly adopted cell-based screening assays ...

Identification of Kinase-substrate Pairs Using High Throughput Screening

We have developed a screening platform to identify dedicated human protein kinases for phosphorylated substrates which can be used to elucidate novel signal transduction pathways. Our approach features the use of a library of purified GST-tagged human protein kinases and a recombinant protein substrate of interest. We have used this technology to identify MAP/microtubule affinity-regulating kinase 2 (MARK2) as the kinase for a glucose-regulated site on CREB-Regulated Transcriptional Coactivator 2 (CRTC2), a protein required for beta cell proliferation, as well as the Axl family of tyrosine kinases as regulators of cell metastasis by phosphorylation of the adaptor protein ELMO. We describe this technology and discuss how it can help to establish a comprehensive map of how cells respond to environmental stimuli.

Protein phosphorylation is a central feature of how cells interpret and respond to information in their extracellular milieu. Here, we present a high throughput screening protocol using kinases purified from mammalian cells to rapidly identify kinases that phosphorylate a substrate(s) of interest.


We have developed a screening platform to identify dedicated human protein kinases for phosphorylated substrates which can be used to elucidate novel ...

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of energy homeostasis and obesity. The small size of zebrafish allows for assays on embryos to be conducted in a 96- or 384-well plate format, Morpholino and CRISPR based technologies promote ease of genetic manipulation, and drug treatment by bath application is viable. Moreover, zebrafish are ideal for forward genetic screens allowing for novel gene discovery. Given the relative novelty of zebrafish as a model for obesity, it is necessary to develop tools that fully exploit these benefits. Herein, we describe a method to measure energy expenditure in thousands of embryonic zebrafish simultaneously. We have developed a whole animal microplate platform in which we use 96-well plates to isolate individual fish and we assess cumulative NADH2 production using the commercially available cell culture viability reagent alamarBlue. In poikilotherms the relationship between NADH2 production and energy expenditure is tightly linked. This energy expenditure assay creates the potential to rapidly screen pharmacological or genetic manipulations that directly alter energy expenditure or alter the response to an applied drug (e.g. insulin sensitizers).

High Throughput Danio Rerio Energy Expenditure Assay

Zebrafish are an important model organism for the study of energy homeostasis. By utilizing a NADH2 sensitive redox indicator, alamar Blue, we have developed an assay that measures the metabolic rate of zebrafish larvae in a 96 well plate format and can be applied to drug or gene discovery.

Zebrafish are an important model organism with inherent advantages that have the potential to make zebrafish a widely applied model for the study of ...