This work is supported by research grants from the National Institutes of Health (HL068915; HL088206), a New Researcher Award from the Thrasher Research Fund, and contributions to the Cardiac Conduction Fund at Children's Hospital Boston.

Part 1: Assemble construct casting molds
<ol>
	<li>Use a razor blade to halve silicone tubing (VWR) and cut it into 3 cm long pieces.</li>
	<li>Place a drop of implant-grade RTV silicone adhesive (Rhodia) on the inside of each end of the tubing.</li>
	<li>Quickly place a small piece (1 cm x 1 cm) of polyester mesh (McMaster-Carr) on the silicone adhesive drop and align it with the end of the tubing. This will provide a slightly raised and flat surface for construct attachment. Repeat for the other end.</li>
	<li>Allow the mold to dry at room temperature for 3 days. It is helpful to make 20 to 30 molds at a time, as these are essentially disposable.</li>
	<li>Once the adhesive is completely dry, store these molds in a beaker filled with 70% ethanol until they are needed. This step will help sterilize the molds and the finished product is shown in Figure 1.</li>
</ol>
Part 2: Preparation for myoblast cell isolation
<ol>
	<li>Dilute 1 mg Laminin (Sigma) in 250 mL 0.22 &#956;m filter-sterilized phosphate-buffered saline (PBS) containing 1% Penicillin/Streptomycin (Invitrogen) and 1% Fungizone (Invitrogen).</li>
	<li>Coat 150 mm tissue culture plates (BD Falcon) with 4 mL of the diluted Laminin solution from step 1.1 and incubate at 37&#186;C for at least 4 hours before plating the primary skeletal myoblasts.</li>
	<li>Make the myoblast medium by mixing Ham&#8217;s F-10 Nutrient Mixture (Sigma) with 20% FBS (Atlanta Biologicals), 5 ng/L basic Fibroblast Growth Factor (Promega), 1% Penicillin/Streptomycin (Invitrogen), and 1% Fungizone (Invitrogen).</li>
	<li>Prepare the skeletal muscle digestion solution by dissolving 1 g of ~295 U/mg Collagenase 2 (Worthington) in 100 mL 2.4 U/mL Dispase-2 (Roche). Add 0.037 g CaCl2 and mix well. Filter sterilize the digestion buffer by using a 0.2 &#181;m filter disk fitted onto a 10 mL syringe and store aliquots at -20&#176;C. Place the amount needed for tissue digestion at 37&#176;C along with the myoblast media.</li>
</ol>
Part 3: Myoblast isolation from skeletal muscle
<ol>
	<li>Using forceps and small scissors, remove paraspinal muscles from dead neonatal Lewis rats by slicing down the back along the spinal cord, peeling back the skin and fascia layers and then excising the muscles.</li>
	<li>In a tissue culture hood, place the excised muscles in a 50 mL conical tube (BD Falcon) filled with 40 mL of 1X Hanks Balanced Salt Solution (HBSS) (Invitrogen) on ice. Allow the muscle strips to settle to the bottom of the tube and carefully remove most of the liquid and blood remnants by vacuum suction with a sterile Pasteur pipette in a culture hood (Baker).</li>
	<li>Pour the harvested muscle pieces onto a 150 mm culture dish and remove as much of the remaining liquid as possible with suction. Using 2 single-edged razor blades mince the tissue to a thick paste. Pour pre-warmed digestion solution on the paste and transfer the mixture to a fresh 50 mL tube using a 10 mL pipette. During the enzymatic digestion, the tissue is occasionally triturated (without bubbling) with a 10 mL pipette fitted onto a cordless Pipette-Aid (Drummond). Again, allow the tissue to settle and remove the excess liquid.</li>
	<li>Place the tube on a rocking platform and digest at 37&#186;C for about 30 min.</li>
	<li>Once the muscle is liquefied, pass the solution through a 70 &#956;m cell strainer (BD Falcon) and centrifuge at room temperature for 10 min at 600 x g (Beckman GP). With a Pasteur pipette and suction, remove as much of the liquid as possible and resuspend the pellet with 10 mL of warm myoblast medium.</li>
	<li>Pre-plate the cells on an uncoated 150 mm tissue culture plate for 15 minutes to help remove fibroblasts.</li>
	<li>Transfer the cells that have not yet attached to the laminin-coated cultureware and dilute so that there is one plate for every 1 to 2 rat pups. Place the plate in a humidified incubator set at 37 &#176;C with 5% CO2.</li>
</ol>
Part 4: Casting engineered tissue constructs (5 mL)
<ol>
	<li>After 1 to 2 days, remove 10 of the 150 mm myoblast tissue culture plates from the incubator, rinse them twice with 37&#176;C 1X PBS, and add 4 mL of warm 0.05% Trypsin-EDTA to each plate. Return the plates to the incubator for 5 min.</li>
	<li>Harvest the detached myoblasts from the plates in the culture hood and collect the liquid in a 50 mL tube (BD Falcon). Use 10 mL of myoblast medium to rinse all of the plates and combine this with the cells in the centrifuge tube. Pellet the cells at room temperature for 10 min at 600 x g (Beckman GP) and remove excess liquid as described in step 3.5.</li>
	<li>Resuspend the myoblast cells in 1.6 mL of myoblast medium and place on ice.</li>
	<li>Using forceps, carefully remove 4 construct molds from step 1.5 and rinse them twice with 1X PBS. Vacuum suction off all traces of PBS with a Pasteur pipette before placing each mold in a 6-well tissue culture plate (BD Falcon).</li>
	<li>Gather the other reagents needed for construct preparation (10X Ham&#8217;s F10, antibiotics, type 1 collagen [3.9 mg/mL][6] and Matrigel&#8482;) on ice and keep myoblast media at 37&#186;C. Combine 500 &#956;L 10X Ham&#8217;s F10, 100 &#181;L Penicillin/Streptomycin, 100 &#181;L Fungizone, 1.6 mL collagen, and 750 &#181;L Matrigel&#8482; in a 15 mL conical tube. Mix the solution for 10 sec using a Mini-Vortexer (VWR) set to 10 and place on ice.</li>
	<li>Next, add 80 &#181;L of NaHCO3 and the myoblast suspension from step 3.3 to the mixture from step 3.5. Vortex the entire mixture for 10 sec and return the tube to the ice bucket for a few min to clear the solution of bubbles.</li>
	<li>Carefully cast the mixture into the molds before it begins to solidify using a 10 mL pipette. Each mold will hold approximately 1 mL of the viscous liquid. Try to avoid introducing air bubbles during this procedure.</li>
	<li>Carefully transfer the plate containing the cast constructs to the incubator. After ~30 min, remove the solidified constructs from the incubator and carefully add myoblast medium to each well to cover the tissue construct. Return the plate to the incubator (Figure 2).</li>
</ol>
Part 5: Representative results
When this protocol is properly executed, the myoblast-containing tissue construct is ready for in vivo implantation (i.e. when removed from the mold) or for further in vitro analyses after 2 days.
<img alt="" src="/files/ftp_upload/1137/Figure1.jpg" />
Figure 1. Examples of completed construct molds (see Step 1.5).
<img alt="" height="321" src="/files/ftp_upload/1137/Figure2.jpg" width="400" />
Figure 2. A solidified myoblast-containing tissue construct[1].
<img alt="" src="/files/ftp_upload/1137/Figure3.jpg" />
Figure 3. H&#38;E stained sections of a myoblast-containing tissue construct. &#34;A&#34; depicts a longitudinal section and &#34;B&#34; shows a cross-section.

<ol>
	<li>Choi, Y. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cardiac+conduction+through+engineered+tissue.">Cardiac conduction through engineered tissue.</a> Am J Pathol. 169 (1), 72-85 (2006).</li><li>Rando, T. A., Blau, H. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+mouse+myoblast+purification,+characterization,+and+transplantation+for+cell-mediated+gene+therapy.">Primary mouse myoblast purification, characterization, and transplantation for cell-mediated gene therapy.</a> J Cell Biol. 125 (6), 1275-1287 (1994).</li><li>Blau, H. M., Webster, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+and+characterization+of+human+muscle+cells.">Isolation and characterization of human muscle cells.</a> Proc Natl Acad Sci U S A. 78 (9), 5623-5627 (1981).</li><li>Powell, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+human+bioartificial+muscles+expressing+a+foreign+recombinant+protein+for+gene+therapy.">Tissue-engineered human bioartificial muscles expressing a foreign recombinant protein for gene therapy.</a> Hum Gene Ther. 10 (4), 565-577 (1999).</li><li>Vandenburgh, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tissue-engineered+skeletal+muscle+organoids+for+reversible+gene+therapy.">Tissue-engineered skeletal muscle organoids for reversible gene therapy.</a> Hum Gene Ther. 7 (17), 2195-2200 (1996).</li><li>Gallop, P. M., Seifter, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+and+Properties+of+Soluble+Collagens.">Preparation and Properties of Soluble Collagens.</a> Methods in Enzymology. 6, 635-641 (1963).</li></ol>

Experiments on animals were performed in accordance with the guidelines and regulations set forth by the Institutional Animal Care and Use Committee at Children's Hospital Boston.

The molds in which the tissue construct will be cast can be made in any shape and size; however, there needs to be at least two points of attachment. Otherwise, the matrix and cells form a spherical structure and the cells die. In the present protocol, we describe the use of a polyester mesh for this purpose, yet we have also successfully used stainless steel mesh. Obviously, larger molds will require more cells and a larger volume of the other ingredients. When making the molds, it is important to minimize the amount of...

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		<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>Silicone tubing</td>
<td>&nbsp;</td>
<td>VWR</td>
<td>60985-724</td>
<td>&nbsp;</td>
<tr>
<td>Silicone adhesive</td>
<td>&nbsp;</td>
<td>Rhodia Silicones</td>
<td>MED ADH 4300 RTV</td>
<td>&nbsp;</td>
<tr>
<td>Polyester Mesh</td>
<td>&nbsp;</td>
<td>McMaster-Carr</td>
<td>93185T17</td>
<td>&nbsp;</td>
<tr>
<td>Laminin</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>L2020</td>
<td>&nbsp;</td>
<tr>
<td>Nutrient Mixture F-10 HAM</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>N6908</td>
<td>&nbsp;</td>
<tr>
<td>Fetal Bovine Serum</td>
<td>&nbsp;</td>
<td>Atlanta Biologicals</td>
<td>S11550</td>
<td>&nbsp;</td>
<tr>
<td>Penicillin/Streptomycin</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15140</td>
<td>&nbsp;</td>
<tr>
<td>Fungizone</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>15290-018</td>
<td>&nbsp;</td>
<tr>
<td>Dispase-2</td>
<td>&nbsp;</td>
<td>Roche</td>
<td>10295825001</td>
<td>&nbsp;</td>
<tr>
<td>Collagenase 2</td>
<td>&nbsp;</td>
<td>Worthington</td>
<td>46H8863</td>
<td>&nbsp;</td>
<tr>
<td>Basic Fibroblast Growth Factor</td>
<td>&nbsp;</td>
<td>Promega</td>
<td>G5071</td>
<td>&nbsp;</td>
<tr>
<td>150 mm tissue culture dishes</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>353025</td>
<td>&nbsp;</td>
<tr>
<td>0.05% (1X) Trypsin-EDTA</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>25300</td>
<td>&nbsp;</td>
<tr>
<td>1X Hanks Balanced Salt Solution</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>14170-112</td>
<td>&nbsp;</td>
<tr>
<td>7.5% NaHCO3</td>
<td>&nbsp;</td>
<td>Gibco</td>
<td>25080-094</td>
<td>&nbsp;</td>
<tr>
<td>70 &mu;m cell strainer</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>352350</td>
<td>&nbsp;</td>
<tr>
<td>6-well plates</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>353046</td>
<td>&nbsp;</td>
<tr>
<td>50 mL Conical Vial</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>352098</td>
<td>&nbsp;</td>
<tr>
<td>15 mL Conical Vial</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>352099</td>
<td>&nbsp;</td>
<tr>
<td>0.2 &mu;m filter</td>
<td>&nbsp;</td>
<td>Nalgene</td>
<td>194-2520</td>
<td>&nbsp;</td>		</tbody>
		</table>
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fabrication of myogenic engineered tissue constructs

Despite the fact that electronic pacemakers are life-saving medical devices, their long-term performance in pediatric patients can be problematic owing to the restrictions imposed by a child's small size and their inevitable growth. Consequently, there is a genuine need for innovative therapies designed specifically for pediatric patients with cardiac rhythm disorders. We propose that a conductive biological alternative consisting of a collagen-based matrix containing autologously-derived cells could better adapt to growth, reduce the need for recurrent surgeries, and greatly improve the quality of life for these patients. In the present study, we describe a procedure for incorporating primary skeletal myoblast cell cultures within a hydrogel matrix to fashion a surgically-implantable tissue construct that will serve as an electrical conduit between the upper and lower chambers of the heart. Ultimately, we anticipate using this type of engineered tissue to restore atrioventricular electrical conduction in children with complete heart block. In view of that, we isolate myoblasts from the skeletal muscles of neonatal Lewis rats and plate them onto laminin-coated tissue culture dishes using a modified version of established protocols[2, 3]. After one to two days, cultured cells are collected and mixed with antibiotics, type 1 collagen, Matrigel&trade;, and NaHCO3. The result is a viscous, uniform solution that can be cast into a mold of nearly any shape and size[1, 4, 5]. For our tissue constructs, we employ type 1 collagen isolated from fetal lamb skin using standard procedures[6]. Once the tissue has solidified at 37&deg;C, culture media is carefully added to the plate until the construct is submerged. The engineered tissue is then allowed to further condense through dehydration for 2 more days, at which point it is ready for in vitro assessment or surgical-implantation.

Watch this Scientific Journal Video about Fabrication of Myogenic Engineered Tissue Constructs at JoVE.com

Fabrication of Myogenic Engineered Tissue Constructs

Here, we demonstrate fabrication of collagen-based, tissue constructs containing skeletal myoblasts. These 3-D engineered constructs may be used to replace or repair tissues in vivo. For our purposes, we have designed these as an atrioventricular electrical conduit for the repair of complete heart block[1].

Despite the fact that electronic pacemakers are life-saving medical devices, their long-term performance in pediatric patients can be problematic owing to ...

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10.4K Views.  Children's Hospital Boston and Harvard Medical School. Here, we demonstrate fabrication of collagen-based, tissue constructs containing skeletal myoblasts. These 3-D engineered constructs may be used to replace or repair tissues in vivo. For our purposes, we have designed these as an atrioventricular electrical conduit for the repair of complete heart block[1].

Video: Fabrication of Myogenic Engineered Tissue Constructs

PDMS Device Fabrication and Surface Modification

Fabrication of the Thermoplastic Microfluidic Channels

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using polymer/nanoparticle composite microscale lysis and solid phase extraction columns. The recovered samples are concentrated, small volume samples that are PCRable, without any additional cleanup. Here, we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then, we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns. We demonstrate grafting and polymerization of a carbon nanotube/polymer composite column for bacterial cell lysis. We then show the lysis process followed by solid phase extraction of nucleic acids from the sample on chip using a silica/polymer composite column. The attached protocols contain detailed instructions on how to make both lysis and solid phase extraction columns.

Here we demonstrate how to fabricate thermoplastic microfluidic chips using hot embossing and heat sealing. Then we demonstrate how to use in situ light directed surface grafting and polymerization through the sealed chip to form the composite solid phase columns.

In our lab, we have successfully isolated nucleic acids directly from microliter and submicroliter volumes of human blood, urine and stool using ...

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Fabrication of Amperometric Electrodes

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we describe the techniques needed to generate low noise (&lt;0.5 pA) electrodes.  The techniques have been modified from published descriptions by previous researchers (1,2).  Electrodes are made by preparing carbon fibers and threading them individually into each capillary tube by using a vacuum with a filter to aspirate the fiber.  Next, the capillary tube with fiber is pulled by an electrode puller, creating two halves, each with a fine-pointed tip.  The electrodes are dipped in hot, liquid epoxy mixed with hardener to create an epoxy-glass seal.  Lastly, the electrodes are placed in an oven to cure the epoxy.  Careful handling of the electrodes is critical to ensure that they are made consistently and without damage.  This protocol shows how to fabricate and cut amperometric electrodes for recording from single cells.

This protocol describes how to generate carbon fiber electrodes. The electrodes are subsequently used to detect catecholamine release from vesicles with carbon fiber amperometry.

Carbon fiber electrodes are crucial for the detection of catecholamine release from vesicles in single cells for amperometry measurements.  Here, we ...

Implantation of Engineered Tissue in the Rat Heart

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in common laboratory rats or mice and these quite frequently serve as an intermediate step between bench-top experiments and large animal testing or human trials. Given that rodents provide an established, cost-effective, and physiologically-relevant model system in which to test novel combinations of scaffolding materials and cells, they are particularly well-suited for cardiovascular tissue engineering studies. Presently, we describe an open-heart surgical procedure to implant engineered tissue containing myogenic progenitor cells in the atrioventricular (AV) groove of a rat heart. These implants are intended to create an electrical conduit between the right atrium and right ventricle with the ultimate goal of providing an alternative treatment to conventional pacemaker implantation in pediatric patients with complete heart block[1]. The engineered tissue is implanted in the AV-groove by means of a thoracotomy. For our purposes, Lewis rats are anesthetized and invasively ventilated to maintain positive airway pressure during the sterile surgical procedure. The approach to the heart is performed by a right thoracotomy through an antero-lateral incision at the 5th intercostal space. The tissue construct is fixed in the AV groove using a single 7-0 Prolene suture and positioned between the right ventricle and atrium at the ventral portion of the heart. The epicardium is partially removed to allow direct contact between the recipient myocardial cells and those contained in the engineered tissue. Following implantation, the chest wall is closed in layers, any pneumothorax is evacuated, and the animal is extubated and treated with analgesic.

Here, we describe a cardiac surgical procedure to implant engineered tissue in the atrioventricular (AV)-groove of an adult Lewis rat.

Rodent surgery is often an important component in assessing the utility of engineered tissues. A wide variety of surgical procedures can be performed in ...

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 ...

Recombineering Homologous Recombination Constructs in Drosophila

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a genetic model organism for human-disease related research. Recent years have seen technical advancements like homologous recombination and recombineering. However, generating unequivocal null mutations or tagging endogenous proteins remains a substantial effort for most genes. Here, we describe and demonstrate techniques for using recombineering-based cloning methods to generate vectors that can be used to target and manipulate endogenous loci in vivo. Specifically, we have established a combination of three technologies: (1) BAC transgenesis/recombineering, (2) ends-out homologous recombination and (3) Gateway technology to provide a robust, efficient and flexible method for manipulating endogenous genomic loci. In this protocol, we provide step-by-step details about how to (1) design individual vectors, (2) how to clone large fragments of genomic DNA into the homologous recombination vector using gap repair, and (3) how to replace or tag genes of interest within these vectors using a second round of recombineering. Finally, we will also provide a protocol for how to mobilize these cassettes in vivo to generate a knockout, or a tagged gene via knock-in. These methods can easily be adopted for multiple targets in parallel and provide a means for manipulating the Drosophila genome in a timely and efficient manner.

Recombineering Homologous Recombination Constructs in Drosophila

Homologous recombination techniques greatly advance Drosophila genetics by enabling the creation of molecularly precise mutations. The recent adoption of recombineering allows one to manipulate large pieces of DNA and transform them into Drosophila6. The methods presented here combine these techniques to rapidly generate large homologous recombination vectors.

The continued development of techniques for fast, large-scale manipulation of endogenous gene loci will broaden the use of Drosophila melanogaster as a ...

Preparation of Primary Myogenic Precursor Cell/Myoblast Cultures from Basal Vertebrate Lineages

Due to the inherent difficulty and time involved with studying the myogenic program in vivo, primary culture systems derived from the resident adult stem cells of skeletal muscle, the myogenic precursor cells (MPCs), have proven indispensible to our understanding of mammalian skeletal muscle development and growth. Particularly among the basal taxa of Vertebrata, however, data are limited describing the molecular mechanisms controlling the self-renewal, proliferation, and differentiation of MPCs. Of particular interest are potential mechanisms that underlie the ability of basal vertebrates to undergo considerable postlarval skeletal myofiber hyperplasia (i.e.&#160;teleost fish) and full regeneration following appendage loss (i.e.&#160;urodele amphibians). Additionally, the use of cultured myoblasts could aid in the understanding of regeneration and the recapitulation of the myogenic program and the differences between them. To this end, we describe in detail a robust and efficient protocol (and variations therein) for isolating and maintaining MPCs and their progeny, myoblasts and immature myotubes, in cell culture as a platform for understanding the evolution of the myogenic program, beginning with the more basal vertebrates. Capitalizing on the model organism status of the zebrafish (Danio rerio), we report on the application of this protocol to small fishes of the cyprinid clade Danioninae. In tandem, this protocol can be utilized to realize a broader comparative approach by isolating MPCs from the Mexican axolotl (Ambystomamexicanum) and even laboratory rodents. This protocol is now widely used in studying myogenesis in several fish species, including rainbow trout, salmon, and sea bream1-4.

In vitro culture systems have proven indispensible to our understanding of vertebrate myogenesis. However, much remains to be learned about nonmammalian skeletal muscle development and growth, particularly in basal taxa. An efficient and robust protocol for isolating the adult stem cells of this tissue, the myogenic precursor cells (MPCs), and maintaining their self-renewal, proliferation, and differentiation in a primary culture setting allows for the identification of conserved and divergent regulatory mechanisms throughout the vertebrate lineages.

Due to the inherent difficulty and time involved with studying the myogenic program in vivo, primary culture systems derived from the resident adult stem ...

Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

RNA-mediated knockdowns are widely used to control gene expression. This versatile family of techniques makes use of short RNA (sRNA) that can be synthesized with any sequence and designed to complement any gene targeted for silencing. Because sRNA constructs can be introduced to many cell types directly or using a variety of vectors, gene expression can be repressed in living cells without laborious genetic modification. The most common RNA knockdown technology, RNA interference (RNAi), makes use of the endogenous RNA-induced silencing complex (RISC) to mediate sequence recognition and cleavage of the target mRNA. Applications of this technique are therefore limited to RISC-expressing organisms, primarily eukaryotes. Recently, a new generation of RNA biotechnologists have developed alternative mechanisms for controlling gene expression through RNA, and so made possible RNA-mediated gene knockdowns in bacteria. Here we describe a method for silencing gene expression in E. coli that functionally resembles RNAi. In this system a synthetic phagemid is designed to express sRNA, which may designed to target any sequence. The expression construct is delivered to a population of E. coli cells with non-lytic M13 phage, after which it is able to stably replicate as a plasmid. Antisense recognition and silencing of the target mRNA is mediated by the Hfq protein, endogenous to E. coli. This protocol includes methods for designing the antisense sRNA, constructing the phagemid vector, packaging the phagemid into M13 bacteriophage, preparing a live cell population for infection, and performing the infection itself. The fluorescent protein mKate2 and the antibiotic resistance gene chloramphenicol acetyltransferase (CAT) are targeted to generate representative data and to quantify knockdown effectiveness.

Phage-mediated Delivery of Targeted sRNA Constructs to Knock Down Gene Expression in E. coli

We describe a method to knock down gene expression in a growing population of E. coli cells using sequence-targeted sRNA expression cassettes delivered by an M13 phagemid vector.

RNA-mediated knockdowns are widely used to control gene expression. This versatile family of techniques makes use of short RNA (sRNA) that can be ...

Dissection of the Auditory Bulla in Postnatal Mice: Isolation of the Middle Ear Bones and Histological Analysis

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called the auditory bulla houses the ossicles, whereas the auditory capsule encloses the inner ear, namely the cochlea and semicircular canals. Murine ossicles are essential for hearing and thus of great interest to researchers in the field of otolaryngology, but their metabolism, development, and evolution are highly relevant to other fields. Altered bone metabolism can affect hearing function in adult mice, and various gene-deficient mice show changes in morphogenesis of auditory ossicles in utero. Although murine auditory ossicles are tiny, their manipulation is feasible if one understands their anatomical orientation and 3D structure. Here, we describe how to dissect the auditory bulla and capsule of postnatal mice and then isolate individual ossicles by removing part of the bulla. We also discuss how to embed the bulla and capsule in different orientations to generate paraffin or frozen sections suitable for preparation of longitudinal, horizontal, or frontal sections of the malleus. Finally, we enumerate anatomical differences between mouse and human auditory ossicles. These methods would be useful in analyzing pathological, developmental and evolutionary aspects of auditory ossicles and the middle ear in mice.

We present a protocol to isolate the auditory bulla, capsule, and ossicles from postnatal mice for whole mount and histological analysis.

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called ...