This work was supported by NIH R01 HG002995 "TILLING the Zebrafish Genome: a Reverse Genetics Approach" to CBM.

Part A: Sperm Freezing Solutions
1. Ginsberg Fish Ringers (Make fresh every 3 days ;Ginsburg, 1963)
NOTE: ORDER OF ADDITION IS IMPORTANT TO PREVENT PRECIPITATION
In 450 ml sterile ddH2O dissolve:
<ul>
<li>NaCl 3.25 g</li>
<li>KCl 0.125 g</li>
<li>CaCl2&bull;2 H2O 0.175 g 
 Then add:</li>
<li>NaHCO3 0.10 g</li>
</ul>
Bring final volume to 500 ml with sterile ddH2O and store at 4&deg;C.
2. Freezing Medium (Make fresh daily)
<ol style="list-style-type:lower-roman">
<li>WITHOUT Methanol
<ul><li>Ginsburg Fish Ringers 10 ml (room temp.)</li>
<li>Powdered Skim Milk 1.5 gm</li>
</ul></li>
<li>WITH Methanol
 NOTE: ORDER OF ADDITION IS IMPORTANT TO PREVENT PRECIPITATION 
<ul>
<li>Ginsburg Fish Ringers 9 ml (room temp.)</li>
<li>Methanol 1 ml</li>
<li>Powdered Skim Milk 1.5 g
</li></ul>
 After assembling freezing media, mix well for 20 min. on orbital shaker or rocker. Before using, aliquot into 1 ml eppendorf tubes, avoiding surface bubbles. 
</li></ol>
Part B: Materials needed for sperm freezing
Materials needed for sperm freezing
<ol>
 <li>10 &mu;l disposable pipettes (Fisherbrand cat# 22-358697) </li>
 <li>250ml beakers for fish water containing MESAB/Tricaine </li>
 <li>watch glasses (Pyrex cat# 9985-75) </li>
 <li>P20 pipetman (or equivalent) and tips </li>
 <li>Sponge fish holder (to hold male while squeezing) </li>
 <li>Dissecting microscope with above stage lighting </li>
 <li>Flat forceps (e.g. Millipore type) </li>
 <li>2.0ml cryogenic vials (Corning cat# 430488) </li>
 <li>10X10 cryoboxes (Nalgene cat#03-337-7AA) </li>
 <li>Styrofoam container filled with ~ 15 cm of finely crushed dry ice1 </li>
 <li>15 ml conical tubes (Falcon # 352099) </li>
 <li>Dewar flask containing liquid nitrogen </li>
 <li>Long tongs (e.g. CMS Fisher Health Care cat# 10-316B) </li>
 <li>Cryogloves </li>
 <li>Recovery tank for males </li>
 <li>One eppendorf tube filled with freezing media without methanol </li>
 <li>One eppendorf tube filled with freezing media with methanol</li>
 <li>Cryofreezer (e.g. Taylor-Warton K-series)</li>
</ol>
1 A simple method for making finely crushed dry ice is to wrap it in a towel and pulverize it with a hammer. A more efficient method is to use a dry ice grinder, e.g. Clawson model RE-2.
Part C: Sperm Freezing Method
<ol>
<li>MARK CAPILLARY TUBES: Prior to beginning, mark the 10&mu;l capillary tubes with a lab pen at the level of 3.33 &mu;l (i.e. 16.7 mm from bottom; see Fig. 1.1).</li>
<li>ANESTHETIZE MALE: Place male(s) in beaker containing MESAB/tricaine diluted in fish water (recipe in "The Zebrafish Book")</li>
<li>DRY FISH: Once anesthetized, lift fish out of anesthetic using a plastic spoon and gently blot fish dry on a paper towel, paying special attention to drying the ventral side. Water activates sperm so it is important to thoroughly dry around the cloaca. Position fish on sponge holder, ventral side up (Fig. 1.2). If region around anal fin is still moist, gently dry with a kimwipe. Take care to not squeeze sperm prematurely!</li>
<li>Place marked capillary tube in rubber mouth pipette adapter that is included with capillary tubes. The adaptor is a rubber hose that has a mouthpiece on one end and a capillary holder on the other. Alternatively, the capillary tube can be connected to a 50&mu;l Hamilton syringe via a short piece of tygon tubing of the appropriate diameter.</li>
<li>Place male under the scope and expose the urogenital opening by carefully spreading the pelvic fins apart using the end of the capillary tube.</li>
<li>COLLECT SPERM: Expel sperm by gently stroking the sides of the fish with smooth forceps (e.g. Millipore), or by gently squeezing the sides of the fish between your index finger and thumb. Collect sperm in capillary tube as it is expelled using gentle suction (Fig. 1.2). Avoid feces that may be expelled with sperm.</li>
<li>NORMALIZE SPERM VOLUME TO 3.3 &mu;l: If the volume of sperm reaches, or exceeds pen mark on capillary tube (i.e. 16.7 mm or greater), then proceed to step 8. If sperm volume does not reach target volume, then normalize volume to pen mark using Freeze Medium WITHOUT Methanol (Fig. 1.3). Minimum amount of sperm that is acceptable varies with quality of sperm. Good sperm is white and opaque. Poor sperm looks watery. In general, we accept as minimums: 1 &mu;l good sperm (or 1/3 target) and 2 &mu;l poor sperm (or 2/3 target). Volume is now normalized to 3.3 &mu;l.</li>
<li>ADD CRYOPROTECTANT TO SPERM : Suck up Freezing Medium WITH Methanol to the orange mark (approx. total volume is now 20 &mu;l; Fig. 1.4). Expel sperm and cryoprotectant mixture onto clean area of a watch glass, paying special attention to not introduce bubbles. Gently mix by pipetting up and down ~ 4 times (avoid introducing bubbles).</li>
<li>PLACE 10&mu;l SPERM INTO EACH OF 2 CRYOVIALS: Pipette 10 &mu;l of the sperm:cryoprotectant solution into the bottom of two separate cryovials that have been labeled with relevant information (Fig. 1.5). Moving quickly, cap vials and drop them into the bottom of room temperature 15 ml Falcon tubes- one cryovial/tube. Cap Falcon tube and insert the pair of cryovial-containing Falcon tubes into crushed dry ice.</li>
<li>FREEZE SPERM FOR 20 MIN. ON DRY ICE: Dry ice should be in fine powder form, not pellets. The tubes should be inserted into the dry ice deep enough that only their caps show (Fig. 1.6). In order to keep track of tubes in dry ice, give each pair of Falcon tubes a number from 1-20 (written on the caps) and record the time they go into the dry ice.</li>
<li>PLACE CRYOVIALS INTO LIQUID NITROGEN: After the sperm has been frozen on dry ice for 20 min., transfer them to a liquid nitrogen- containing dewar flask. Sperm is stored here until time to place into liquid nitrogen freezer. When placing in the freezer boxes, place freezer box in a bath of liquid nitrogen so that samples do not heat up. Use long metal tongs to recover vials from dewar flask and handle with cryogloves. Alternatively, cryovials can be transferred directly from dry ice into the freezer box if the freezer box is kept immersed in liquid nitrogen in a Styrofoam container.</li>
<li>Store cryovials long term in a cryogenic liquid nitrogen freezer. To maintain viability of sperm, it is important that vials are stored immersed in liquid nitrogen. In our experience, vials stored in the vapor phase lose viability over time. Speed is important! The time between adding the cryoprotectant and placing sperm vials in dry ice (i.e. steps 6-10) should be no more than 30 sec. </li></ol>
Part D: In Vitro Fertilization Method Using Cryopreserved Sperm
We find that this works best with two people. One person squeezes eggs from females while the other person thaws the sperm.
<ol><li>Set a water bath to 33&deg;C.</li>
<li>Remove cryovial from liquid nitrogen freezer and transfer to liquid nitrogen-containing dewar flask until ready to thaw.</li>
<li>Squeeze eggs from females into 35mm plastic petri dish. If possible, try to obtain 3 clutches of eggs and combine into one dish. If you cannot get three good clutches within 1 min of your first clutch, then proceed to step 4 (one good clutch is sufficient). Definition of good clutch: &gt;150 uniformly nice looking eggs (i.e. yellowish, with no white debris indicative of degradation). Keep dish covered while sperm is thawed.</li>
<li>Remove vial from liquid nitrogen and remove cap. Quickly immerse vial ~1/2 way into 33&deg;C water bath for 8-10 sec.</li>
<li>Quickly add 70 &mu;l room temperature Hanks to vial and mix by pipetting up and down. Immediately add to eggs and mix gently with pipette tip.</li>
<li>Without delay, activate sperm and eggs by adding 750 &mu;l fish water. Swirl to mix. Incubate 5 min at room temperature.</li>
<li>After 5 min, fill dish with fish water and incubate dish at 28&deg;C. After 2-3 hrs, count and transfer fertile embryos to 100mm dishes (50/dish). Count infertile eggs so that fertilization frequency can be determined.</li>
<li>Take good care of the larvae! For first 5 days, change water daily and remove dead embryos. Put only larvae that have swim bladders into nursery on day 5.</li>
</ol>
Part E: Representative Results
We used this crypreservation methodology to make a library of 8600 frozen sperm samples for TILLING between the years 2001 and 2003. We have thawed 106 of these samples and determined their fertility. Fertility is the ratio of viable embryos produced in an in vitro fertilization divided by the total number of eggs that were fertilized with the frozen sperm. Fresh (non cryporeserved) sperm typically has &gt;95% fertility; cryopreserved sperm has much lower fertility: an average of 29% (n=106 vials). The vial-to-vial variability in fertility is large, ranging from less than 1% to as high as 62%. This variability is represented by the large error bars representing standard deviation in Fig. 2. However in 106 sperm samples thawed to date we have only once experienced 0% fertility in both sperm samples and failed to recover the corresponding TILLING mutation.
By thawing sperm vials from our library over the course of several years (2001-2008) we have found that the average fertility of sperm samples frozen by this method remains stable over time (Fig. 2). The average fertility of frozen sperm samples thawed in 2001 was 23.5% (based on 11 sperm samples) and in 2008 was 25% (based on 21 sperm samples).
A number of other investigators are now using this sperm freezing protocol in their labs. Some have reported good success with fertility rates and sample-to-sample variability similar to what we have observed. Others have reportedly failed to make this protocol work in their hands. We believe that likely sources of problems are:
<ol>
<li>Failure to use the exact cryovials or conical tubes we have suggested in the protocol, leading to a different rate of cooling when in dry ice;</li>
<li>Failure to use powdered dry ice, leading to a sub-optimal cooling rate;</li>
<li>Using male fish with poor fertility: you should get at least 1 &mu;l of sperm from a single male. You can also test male fertility by doing an in vitro fertilization with fresh sperm, however this can be misleading because it takes very little fresh sperm to get near 100% fertility.</li>
<li>Using sub-standard quality eggs in the in vitro fertilization. Using anything but perfect clutches always leads to near 0% fertility, even from sperm samples that actually have good fertility (based on the fertility of the duplicate vial).</li>
</ol>
<img src="/files/ftp_upload/1395/1395fig1.jpg" alt="Figure 1" /> Figure 1. Schematic of sperm freezing methodology.
<img src="/files/ftp_upload/1395/1395fig2.jpg" alt="Figure 2" /> Figure 2. Cryopreserved sperm fertility remains stable over time. Percent fertility is the number of viable embryos produced in an in vitro fertilization divided by the total number of eggs that were fertilized with the frozen sperm. "n" refers to the number of cryopreserved sperm vials whose fertility was tested in the corresponding year. Error bars represent standard deviation. Since each sperm vial is from a different male, vial-to-vial variability is observed.

<ol>
	<li>Ginsburg, A. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sperm-egg+association+and+its+relationship+to+activation+of+the+egg+in+salmonid+fishes.">Sperm-egg association and its relationship to activation of the egg in salmonid fishes.</a> J. Embryol. Exp. Morphol. 11, 13-33 (1963).</li><li>Harvey, B., Kelley, R. N., Ashwood-Smith, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cryopreservation+of+zebra+fish+spermatozoa+using+methanol.">Cryopreservation of zebra fish spermatozoa using methanol.</a> Can. J. Zoology. 60, 1867-1870 (1982).</li></ol>

A number of other investigators are now using this sperm freezing protocol in their labs. Some have reported similar fertility and the same sample-to-sample variability that we have observed. Others have reportedly failed to make this protocol work in their hands. We believe that likely sources of problems are: 
<ol>
<li>Failure to use the exact cryovials or conical tubes we have suggested in the protocol, leading to a different rate of cooling when in dry ice;</li>
<li>Failure to use powdered dry ice, leading to a sub-optimal cooling rate;</li>
<li>Using male fish with poor fertility: you should get at least 1 &mu;l of sperm from a single male. You can also test male fertility by doing an in vitro fertilization with fresh sperm, however this can be misleading because it takes very little fresh sperm to get near 100% fertility. Larger, more robustly colored males typically give more higher quality sperm.</li>
<li>Using sub-standard quality eggs in the in vitro fertilization. Using anything but perfect clutches always leads to near 0% fertility, even from sperm samples that actually have good fertility (based on the fertility of the duplicate vial). </li>
</ol>

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>10 &mu;l disposable pipettes</td>
<td>&nbsp;</td>
<td>Fisher</td>
<td>22-358697</td>
<td>&nbsp;</td>
<tr>
<td>Watch glasses</td>
<td>&nbsp;</td>
<td>Pyrex</td>
<td>9985-75</td>
<td>&nbsp;</td>
<tr>
<td>2 ml cryogenic vials</td>
<td>&nbsp;</td>
<td>Corning</td>
<td>430488</td>
<td>&nbsp;</td>
<tr>
<td>10 x 10 cryoboxes</td>
<td>&nbsp;</td>
<td>Nalgene</td>
<td>03-337-7AA</td>
<td>&nbsp;</td>
<tr>
<td>15 ml conical tubes</td>
<td>&nbsp;</td>
<td>Falcon</td>
<td>352099</td>
<td>&nbsp;</td>
<tr>
<td>Tongs</td>
<td>&nbsp;</td>
<td>Fisher Health Care</td>
<td>10-316B</td>
<td>&nbsp;</td>
<tr>
<td>Cryofreezer</td>
<td>&nbsp;</td>
<td>Taylor-Warton</td>
<td>K-series</td>
<td>For example</td>
</tr>		</tbody>
		</table>
		</div>

a high-throughput method for zebrafish sperm cryopreservation and in vitro fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to the original protocol that both streamline the procedure and increase sample uniformity.  First, we normalize all sperm volumes using freezing media that does not contain the cryoprotectant. Second, cryopreserved sperm are stored in cryovials instead of capillary tubes.  The rates of sperm freezing and thawing (&delta;&deg;C/time) are probably the two most critical variables to control in this procedure.  For this reason, do not substitute different tubes for those specified.  Working in teams of 2 it is possible to freeze the sperm of 100 males per team in ~2 hrs.  Sperm cryopreserved using this protocol has an average of 25% fertility (measured as the number of viable embryos generated in an in vitro fertilization divided by the total number of eggs fertilized) and this percent fertility is stable over many years.

Watch this Scientific Journal Video about A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization at JoVE.com

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a high-throughput sperm cryopreservation protocol for zebrafish. Sperm cryopreserved using this protocol has an average of 25% fertility in subsequent vitro fertilization and is stable over many years.

A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

This is a method for zebrafish sperm cryopreservation that is an adaptation of the Harvey method (Harvey et al., 1982).  We have introduced two changes to ...

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30.9K vistas.  University of California, Davis. Este es un protocolo de esperma de alto rendimiento para la criopreservación de pez cebra. Espermatozoides criopreservados utilizando este protocolo tiene un promedio de 25% de la fertilidad en la fertilización in vitro posterior y es estable durante muchos años.

Video: A High-Throughput Method For Zebrafish Sperm Cryopreservation and In Vitro Fertilization

A high-throughput method to globally study the organelle morphology in S. cerevisiae

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help achieve an comprehensive understanding of molecular pathways. In Saccharomyces cerevisiae, with the development of the non-essential gene deletion array, we can globally study the morphology of different organelles like the endoplasmic reticulum (ER) and the mitochondria using GFP (or variant)-markers in different gene backgrounds. However, incorporating GFP markers in each single mutant individually is a labor-intensive process. Here, we describe a procedure that is routinely used in our laboratory. By using a robotic system to handle high-density yeast arrays and drug selection techniques, we can significantly shorten the time required and improve reproducibility. In brief, we cross a GFP-tagged mitochondrial marker (Apc1-GFP) to a high-density array of 4,672 nonessential gene deletion mutants by robotic replica pinning.  Through diploid selection, sporulation, germination and dual marker selection, we recover both alleles. As a result, each haploid single mutant contains Apc1-GFP incorporated at its genomic locus. Now, we can study the morphology of mitochondria in all non-essential mutant background. Using this high-throughput approach, we can conveniently study and delineate the pathways and genes involved in the inheritance and the formation of organelles in a genome-wide setting.

GFP-fusion proteins are widely used to visualize organelles by confocal microscopy. However, screening for mutations that affect the morphology of organelles generally requires individual mutagenesis and is time consuming. Here, we demonstrate a method to simultaneously incorporate organelle-GFP markers in almost 5,000 non-essential genes in yeast.

High-throughput methods to examine protein localization or organelle morphology is an effective tool for studying protein interactions and can help ...

A Rapid High-throughput Method for Mapping Ribonucleoproteins (RNPs) on Human pre-mRNA

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding location often influences function of a splicing factor.  However, as with any sampling strategy the chance of identifying an RNA bound to a splicing factor is proportional to its cellular abundance.  We have developed a novel in vitro approach for surveying binding specificity on otherwise transient pre-mRNA. This approach utilizes a specifically designed oligonucleotide pool that tiles across introns, exons, splice junctions, or other pre-mRNA.  The pool is subjected to some kind of molecular selection. Here, we demonstrate the method by separating the oligonucleotide into a bound and unbound fraction and utilize a two color array strategy to record the enrichment of each oligonucleotide in the bound fraction. The array data generates high-resolution maps with the ability to identify sequence-specific and structural determinates of ribonucleoprotein (RNP) binding on pre-mRNA.  A unique advantage to this method is its ability to avoid the sampling bias towards mRNA associated with current IP and SELEX techniques, as the pool is specifically designed and synthesized from pre-mRNA sequence. The flexibility of the oligonucleotide pool is another advantage since the experimenter chooses which regions to study and tile across, tailoring the pool to their individual needs.  Using this technique, one can assay the effects of polymorphisms or mutations on binding on a large scale or clone the library into a functional splicing reporter and identify oligonucleotides that are enriched in the included fraction.  This novel in vitro high-resolution mapping scheme provides a unique way to study RNP interactions with transient pre-mRNA species, whose low abundance makes them difficult to study with current in vivo techniques.  



A Rapid High-throughput Method for Mapping Ribonucleoproteins RNPs on Human pre-mRNA

Due to the transient nature of pre-mRNA, it can be difficult to isolate and study in vivo. Here, we present a novel in vitro approach to investigate RNA-protein interactions using a synthetic oligo pool that tiles across selected regions of pre-mRNA.

Sequencing RNAs that co-immunoprecipitate (co-IP) with RNA binding proteins has increased our understanding of splicing by demonstrating that binding ...

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping project is to collect a tissue sample from each individual plant. The traditional approach to this task is to sample plants one-at-a-time. If one wishes to genotype hundreds or thousands of individuals, however, using this strategy results in a significant bottleneck in the genotyping pipeline. The Ice-Cap method that we describe here provides a high-throughput solution to this challenge by allowing one scientist to collect tissue from several thousand seedlings in a single day 1,2. This level of throughput is made possible by the fact that tissue is harvested from plants 96-at-a-time, rather than one-at-a-time.
The Ice-Cap method provides an integrated platform for performing seedling growth, tissue harvest, and DNA extraction. The basis for Ice-Cap is the growth of seedlings in a stacked pair of 96-well plates. The wells of the upper plate contain plugs of agar growth media on which individual seedlings germinate. The roots grow down through the agar media, exit the upper plate through a hole, and pass into a lower plate containing water. To harvest tissue for DNA extraction, the water in the lower plate containing root tissue is rapidly frozen while the seedlings in the upper plate remain at room temperature. The upper plate is then peeled away from the lower plate, yielding one plate with 96 root tissue samples frozen in ice and one plate with 96 viable seedlings. The technique is named "Ice-Cap" because it uses ice to capture the root tissue. The 96-well plate containing the seedlings can then wrapped in foil and transferred to low temperature. This process suspends further growth of the seedlings, but does not affect their viability. Once genotype analysis has been completed, seedlings with the desired genotype can be transferred from the 96-well plate to soil for further propagation. We have demonstrated the utility of the Ice-Cap method using Arabidopsis thaliana, tomato, and rice seedlings. We expect that the method should also be applicable to other species of plants with seeds small enough to fit into the wells of 96-well plates.

Ice-Cap: A Method for Growing Arabidopsis and Tomato Plants in 96-well Plates for High-Throughput Genotyping

The Ice-Cap method allows one to grow plants in 96-well plates and non-destructively harvest root tissue from each seedling. DNA extracted from this root tissue can be used for genotyping reactions. We have found that Ice-Cap works well for Arabidopsis thaliana, tomato, and rice seedlings.

It is becoming common for plant scientists to develop projects that require the genotyping of large numbers of plants. The first step in any genotyping ...

High-throughput Crystallization of Membrane Proteins Using the Lipidic Bicelle Method

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane bilayer that surrounds all cells and organelles. Alterations in the function of MPs result in many human diseases and disorders; thus, an intricate understanding of their structures remains a critical objective for biological research. However, structure determination of MPs remains a significant challenge often stemming from their hydrophobicity. 
MPs have substantial hydrophobic regions embedded within the bilayer. Detergents are frequently used to solubilize these proteins from the bilayer generating a protein-detergent micelle that can then be manipulated in a similar manner as soluble proteins. Traditionally, crystallization trials proceed using a protein-detergent mixture, but they often resist crystallization or produce crystals of poor quality. These problems arise due to the detergent&prime;s inability to adequately mimic the bilayer resulting in poor stability and heterogeneity. In addition, the detergent shields the hydrophobic surface of the MP reducing the surface area available for crystal contacts. To circumvent these drawbacks MPs can be crystallized in lipidic media, which more closely simulates their endogenous environment, and has recently become a de novo technique for MP crystallization.
Lipidic cubic phase (LCP) is a three-dimensional lipid bilayer penetrated by an interconnected system of aqueous channels1. Although monoolein is the lipid of choice, related lipids such as monopalmitolein and monovaccenin have also been used to make LCP2. MPs are incorporated into the LCP where they diffuse in three dimensions and feed crystal nuclei. A great advantage of the LCP is that the protein remains in a more native environment, but the method has a number of technical disadvantages including high viscosity (requiring specialized apparatuses) and difficulties in crystal visualization and manipulation3,4. Because of these technical difficulties, we utilized another lipidic medium for crystallization-bicelles5,6 (Figure 1). Bicelles are lipid/amphiphile mixtures formed by blending a phosphatidylcholine lipid (DMPC) with an amphiphile (CHAPSO) or a short-chain lipid (DHPC). Within each bicelle disc, the lipid molecules generate a bilayer while the amphiphile molecules line the apolar edges providing beneficial properties of both bilayers and detergents. Importantly, below their transition temperature, protein-bicelle mixtures have a reduced viscosity and are manipulated in a similar manner as detergent-solubilized MPs, making bicelles compatible with crystallization robots. 
Bicelles have been successfully used to crystallize several membrane proteins5,7-11 (Table 1). This growing collection of proteins demonstrates the versatility of bicelles for crystallizing both alpha helical and beta sheet MPs from prokaryotic and eukaryotic sources. Because of these successes and the simplicity of high-throughput implementation, bicelles should be part of every membrane protein crystallographer&prime;s arsenal. In this video, we describe the bicelle methodology and provide a step-by-step protocol for setting up high-throughput crystallization trials of purified MPs using standard robotics.

Bicelles are lipid/amphiphile mixtures that maintain membrane proteins (MPs) within a lipid bilayer but have unique phase behavior that facilitates high-throughput screening by crystallization robots. This technique has successfully produced a number of high-resolution structures from both prokaryotic and eukaryotic sources. This video describes protocols for generating the lipidic bicelle mixture, incorporating MPs into the bicelle mixture, setting up crystallizations trials (manually as well as robotically) and harvesting crystals from the medium.

Membrane proteins (MPs) play a critical role in many physiological processes such as pumping specific molecules across the otherwise impermeable membrane ...

Mouse Sperm Cryopreservation and Recovery using the I&middot;Cryo Kit

Thousands of new genetically modified (GM) strains of mice have been created since the advent of transgenesis and knockout technologies. Many of these valuable animals exist only as live animals, with no backup plan in case of emergency. Cryopreservation of embryos can provide this backup, but is costly, can be a lengthy procedure, and generally requires a large number of animals for success. Since the discovery that mouse sperm can be successfully cryopreserved with a basic cryoprotective agent (CPA) consisting of 18&#37; raffinose and 3&#37; skim milk, sperm cryopreservation has become an acceptable and cost-effective procedure for archiving, distributing and recovery of these valuable strains.
 Here we demonstrate a newly developed I&bull;Cryo kit for mouse sperm cryopreservation. Sperm from five commonly-used strains of inbred mice were frozen using this kit and then recovered. Higher protection ratios of sperm motility (&gt; 60&#37;) and rapid progressive motility (&gt; 45&#37;) compared to the control (basic CPA) were seen for sperm frozen with this kit in 5 inbred mouse strains. Two cell stage embryo development after IVF with the recovered sperm was improved consistently in all 5 mouse strains examined. Over a 1.5 year period, 49 GM mouse lines were archived by sperm cryopreservation with the I&bull;Cryo kit and later recovered by IVF.

Mouse Sperm Cryopreservation and Recovery using the I&amp;middot;Cryo Kit

Here we demonstrate the newly developed I&#x2022;Cryo kit for mouse sperm cryopreservation. Two-cell stage embryo development with frozen-thawed sperm was improved consistently in 5 mouse strains with the use of this kit. Over a 1.5 year period, 49 genetically modified mouse lines were archived by sperm cryopreservation with the I&#x2022;Cryo kit and later successfully recovered by IVF.

Thousands of new genetically modified (GM) strains of mice have been created since the advent of transgenesis and knockout technologies.  Many of these ...

A High-throughput Method for Measurement of Glomerular Filtration Rate in Conscious Mice

The measurement of glomerular filtration rate (GFR) is the gold standard in kidney function assessment. Currently, investigators determine GFR by measuring the level of the endogenous biomarker creatinine or exogenously applied radioactive labeled inulin (3H or 14C). Creatinine has the substantial drawback that proximal tubular secretion accounts for ~50% of total renal creatinine excretion and therefore creatinine is not a reliable GFR marker. Depending on the experiment performed, inulin clearance can be determined by an intravenous single bolus injection or continuous infusion (intravenous or osmotic minipump). Both approaches require the collection of plasma or plasma and urine, respectively. Other drawbacks of radioactive labeled inulin include usage of isotopes, time consuming surgical preparation of the animals, and the requirement of a terminal experiment. Here we describe a method which uses a single bolus injection of fluorescein isothiocyanate-(FITC) labeled inulin and the measurement of its fluorescence in 1-2 &mu;l of diluted plasma. By applying a two-compartment model, with 8 blood collections per mouse, it is possible to measure GFR in up to 24 mice per day using a special work-flow protocol. This method only requires brief isoflurane anesthesia with all the blood samples being collected in a non-restrained and awake mouse. Another advantage is that it is possible to follow mice over a period of several months and treatments (i.e. doing paired experiments with dietary changes or drug applications). We hope that this technique of measuring GFR is useful to other investigators studying mouse kidney function and will replace less accurate methods of estimating kidney function, such as plasma creatinine and blood urea nitrogen.

Measurement of glomerular filtration rate (GFR) is the gold standard for kidney function assessment. Here we describe a high-throughput method which allows the determination of GFR in conscious mice by using a single bolus injection, determination of fluorescein isothiocyanate (FITC)-inulin in plasma and calculation of GFR by a two-phase exponential decay model.

The measurement of  glomerular filtration rate (GFR) is the gold standard in kidney function  assessment. Currently, investigators determine GFR by ...

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for forward genetic analysis of developmental processes due to their external fertilization, embryonic size, rapid ontogeny, and optical clarity &#8211; a constellation of traits that enable the direct observation of events ranging from gastrulation to organogenesis with a basic stereomicroscope. Further, zebrafish embryos can survive for several days in the haploid state. The production of haploid embryos in vitro is a powerful tool for mutational analysis, as it enables the identification of recessive mutant alleles present in first generation (F1) female carriers following mutagenesis in the parental (P) generation. This approach eliminates the necessity to raise multiple generations (F2, F3, etc.) which involves breeding of mutant families, thus saving the researcher time along with reducing the needs for zebrafish colony space, labor, and the husbandry costs. Although zebrafish have been used to conduct forward screens for the past several decades, there has been a steady expansion of transgenic and genome editing tools. These tools now offer a plethora of ways to create nuanced assays for next generation screens that can be used to further dissect the gene regulatory networks that drive vertebrate ontogeny. Here, we describe how to prepare haploid zebrafish embryos. This protocol can be implemented for novel future haploid screens, such as in enhancer and suppressor screens, to address the mechanisms of development for a broad number of processes and tissues that form during early embryonic stages.

Production of Haploid Zebrafish Embryos by In Vitro Fertilization

The zebrafish is a powerful model system for developmental biology and human disease research due to their genetic similarity with higher vertebrates. This protocol describes a methodology to create haploid zebrafish embryos that can be utilized for forward screen strategies to identify recessive mutations in genes essential for early embryogenesis.

The zebrafish has become a mainstream vertebrate model that is relevant for many disciplines of scientific study. Zebrafish are especially well suited for ...

A Manual Small Molecule Screen Approaching High-throughput Using Zebrafish Embryos

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology due to their considerable degree of genetic conservation with humans. Chemical genetics entails testing the effect that small molecules have on a biological process and is becoming a popular translational research method to identify therapeutic compounds. Zebrafish are specifically appealing to use for chemical genetics because of their ability to produce large clutches of transparent embryos, which are externally fertilized. Furthermore, zebrafish embryos can be easily drug treated by the simple addition of a compound to the embryo media. Using whole-mount in situ hybridization (WISH), mRNA expression can be clearly visualized within zebrafish embryos. Together, using chemical genetics and WISH, the zebrafish becomes a potent whole organism context in which to determine the cellular and physiological effects of small molecules. Innovative advances have been made in technologies that utilize machine-based screening procedures, however for many labs such options are not accessible or remain cost-prohibitive. The protocol described here explains how to execute a manual high-throughput chemical genetic screen that requires basic resources and can be accomplished by a single individual or small team in an efficient period of time. Thus, this protocol provides a feasible strategy that can be implemented by research groups to perform chemical genetics in zebrafish, which can be useful for gaining fundamental insights into developmental processes, disease mechanisms, and to identify novel compounds and signaling pathways that have medically relevant applications.

The zebrafish is an excellent experimental organism to study vertebrate developmental processes and model human disease. Here, we describe a protocol on how to perform a manual high-throughput chemical screen in zebrafish embryos with a whole-mount in situ hybridization (WISH) read-out.

Zebrafish have become a widely used model organism to investigate the mechanisms that underlie developmental biology and to study human disease pathology ...

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 ...