We are grateful to Nancy Papalopulu and Boyan Bonev for assistance with Xenopus electroporation. We also thank Marc Dionne for critical reading, Jeremy Green and John Wallingford for helpful discussions and members of the Liu lab for their support. This work was funded by grants from the BBSRC (BB/E013872/1) and the Wellcome Trust (081880/Z/06/Z) to KJL.

Part 1A. Equipment
Microscope: upright stereo-dissecting scope with low power objective
<ul>
 <li>Voltage/Pulse Generator: BTX ECM 830 Square Wave Electroporation System</li>
 <li>Pipette puller: P-87 Micropipette Puller (Sutter Instrument Company, CA)</li>
 <li>Manipulator: coarse, or combined coarse and fine depending on preparation.</li>
 <li>Micropipette holder: Fine Science Tools</li>
 <li>Electrode: homemade</li>
 <li>Electroporation chamber: homemade</li>
</ul>
L-shaped electrodes:
<ol>
 <li>Cut 8 cm of high-purity 0.4 mm tungsten wire (Goodfellow) and affix at midpoint a 1ml syringe using putty (we use Blu-Tack). Leave 4 cm tungsten wire exposed from the tip of the syringe and bend tip into L-shape, 1 cm from end (Fig. 1A).</li>
 <li>Trim tip so that the end measures 0.5 mm in length. This tip is the electrode terminus.</li>
 <li>Run excess tungsten wire parallel to syringe and use it to connect the electrode pulse generator.</li>
 <li>Repeat process making a pair of electrodes.</li>
 <li>Attach electrodes to square wave pulse generator via DC cables.</li>
</ol>
Electroporation chamber
<ol>
 <li>Line bottom of 90 mm dish with &sim;5 mm non-toxic plasticine.</li>
 <li>Fill dish with electroporation media.</li>
 <li>Using No. 5 watchmaker's forceps carve a T-shaped well (Fig. 2). The long well should measure &sim;2 mm x 2 mm x 10 mm and the short &sim;2 mm x 2 mm x 5 mm. The electroporation dish can be washed and reused.</li>
</ol>
Part 1B. Reagents
DNA or charged macromolecules
<ul>
 <li>Micropipettes: 1 mm wide 4" long borosilicate glass capillaries (WPI, TW100-F)</li>
 <li>Culture media: Normal Amphibian Media (NAM)</li>
 <li>&gt;10 X stock: 1100 mM NaCl, 20 mM KCL, 10 mM Ca(NO3)2&bull;4H2O, 1 mM EDTA.
 <ul>
 <li>Autoclave and store at 4 &deg;C.</li>
 <li>1 X NAM: Dilute from 10x stock, buffer with 0.1 mM NaHCo3 and 0.2 mM Na3PO4.</li>
 </ul>
 </li>
 <li>Xenopus laevis tadpoles, stage 28</li>
</ul>
DNA preparation:
<ol>
 <li>Prepare expression plasmids using standard protocols.</li>
 <li>Resuspend DNA to a final concentration of 1 &mu;g/&mu;l in nuclease free H20.</li>
</ol>
* We have had success with vectors containing a strong CMV promoter, such as pCS2+ [16]. For lineage analysis, we usually include DNA encoding green fluorescent protein (pCS2+GFP) at a final concentration of 0.1 &mu;g/&mu;l. [DNA concentrations between 0.1-3 &mu;g/&mu;l were also tested. We found that concentrations below 0.8 &mu;g/&mu;l inefficiently labelled cells, whereas DNA concentrations greater than 2 &mu;g/&mu;l did not improve electroporation efficiency.]
Morpholino oligonucleotide preparation:
(Note: MOs need to be fluoresceinated (3'-carboxyfluorescein modified) or otherwise charged.)
<ol>
 <li>Resuspend morpholino oligonucleotides (MOs) (Genetools, <a href="http://www.genetools.com" target="_blank">www.genetools.com</a>) at a concentration of 2 mM in nuclease free H20.</li>
 <li>Heat aliquot of stock solution at 65&deg;C for 5 minutes.</li>
 <li>Dilute to final concentration of 0.5 mM in nuclease free water.</li>
</ol>
* 0.1-1mM MO solutions were tested. 0.5 mM MO solutions were sufficient for electroporation of many mesenchymal cells.
Micropipettes
<ol>
 <li>Prepare micropipettes from borosilicate glass capillaries (1 mm wide, 4" long, WPI no. TW100-F). Use needle puller to prepare micropipettes with an 8-12 mm long taper and fine tip.</li>
 <li>Crush tip &sim;2 mm from tip using forceps, creating a jagged break.</li>
</ol>
Media
<ol>
 <li>Incubation media: Prepare fresh 3/4 Normal Amphibian Media (NAM) from 1x stock. Add 0.025 mg/ml gentamycin.</li>
 <li>Electroporation media: as above, with 0.1% Benzocaine (Sigma, 06950).</li>
</ol>
2. Electroporation
Micropipette setup
<ol>
 <li>Fill micropipette with &sim;1 &mu;l injection solution.</li>
 <li>Secure micropipette in micromanipulator and attach to microinjector (Picospritzer II).</li>
 <li>Angle micropipette at 50&deg; from the tabletop.</li>
 <li>Set injection pressure at 20 PSI.</li>
 <li>Calibrate micropipette to inject 30 nl per pulse.</li>
</ol>
Tadpole preparation
<ol>
 <li>Anesthetize stage 28 Xenopus larvae by incubating in electroporation media for 5 minutes.</li>
 <li>Transfer anaesthetised tadpole into electroporation chamber filled with electroporation media. Position embryo within the long well so that the head rests in the T-junction with dorsal side down and ventral side exposed. The head should be slightly elevated compared to the tail.</li>
 <li>Using forceps, gently secure tadpole in well with surrounding plasticine. (Note: If the tadpole is not secured, it may twitch and contact electrode during electroporation. In this case discard the tadpole as facial tissues will be severely damaged.)</li>
</ol>
Electroporation
<ol>
 <li>Insert micropipette tip immediately posterior to the cement gland and into facial mesenchyme.</li>
 <li>Inject 30 nl solution into mesenchyme.</li>
 <li>Retract micropipette.</li>
 <li>Quickly align electrode tips parallel to the head of the embryo (Fig. 3).</li>
 <li>Apply 8 50 ms, 20 V square pulses.</li>
 <li>Retract electrodes.</li>
 <li>Using forceps carefully release tadpole from well and transfer to 3/4 NAM, 0.025 mg/ml gentamycin.</li>
 <li>Tadpoles can be incubated in 3/4 NAM, 0.025 mg/ml overnight, or longer.</li>
 <li>Screen embryos for efficient electroporation by fluorescence microscopy after 24 hours.</li>
</ol>
3. Representative Results:
The use of fluorescent molecules allows easy screening of electroporated embryos. Figure 4 shows a typical batch of MO electroporated tadpoles &sim;12, 48 and 96 hours after electroporation, incubated at 14.5&deg;C. Using fluorescence microscopy, MOs can be visualised immediately after electroporation and persist for several days after electroporation. In our experience, fluorescence is weakly evident at stage 46 (&sim;5 days later). In the cartilages, fluorescence decreases dramatically after the onset of differentiation (&sim;st 42); however, MO fluorescence persists more strongly in other cell types such as the pharyngeal endoderm. Fluorescence microscopy shows that oligonucleotides are incorporated into several craniofacial tissues including cartilage. Oligonucleotide fluorescence can often be visualised in tissue on either side of the head. This is likely due to rapid diffusion of the injection solution throughout the loose craniofacial mesenchyme prior to electroporation.
 
 
 
<img alt="Figure 1" src="/files/ftp_upload/3381/3381fig1.jpg" /> 
Figure 1 Homemade electrodes. L-shaped tungsten wire is attached to a 1 ml syringe using non-toxic clay or putty. (A) The electrode terminus measures 5 mm. (B) Attach a pair of electrodes, such that the termini run parallel. Electrodes are attached to pulse generator by DC cables.
<img alt="Figure 2" src="/files/ftp_upload/3381/3381fig2.jpg" /> 
Figure 2 Electroporation chamber. (A) 90 mm dish lined with plasticine is filled with media and a T-shaped chamber carved with No 5 watchmaker's forceps. (B) The long side measures 2 mm X 2 mm X10 mm whilst the short measures 2 mm X 2 mm X 5 mm. The head of the embryo rests in the T-junction, ventral side up.
<img alt="Figure 3" src="/files/ftp_upload/3381/3381fig3.jpg" /> 
Figure 3 Schematic illustrating electroporation procedure. St. 28 tadpole is placed in electroporation chamber, ventral side up. Micropipette is inserted into facial mesenchyme underlying cement gland. Inject. Micropipette is removed and L-shaped electrodes are aligned parallel flanking the head. Apply eight 50 ms, 20 V square pulses. Retract electrodes. Grow tadpoles to desired stages. Visualise MOs or GFP expression using fluorescence microscopy.
<img alt="Figure 4" src="/files/ftp_upload/3381/3381fig4.jpg" /> 
Figure 4 Representative tadpoles 12 (A), 48 (B), and 96 (C) hours post electroporation (stages 30, 34 and 44 respectively). (A"-B") Fluorescent MO can be visualised within craniofacial mesenchyme at stages 30 and 34. Fluorescence can be detected in cartilages at stage 44 (arrowhead, C'-C"). The gut is highly autofluorescent.

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The authors have no conflict of interests.

In this video, we have demonstrated the feasibility of electroporation-mediated gene delivery into the facial mesenchyme of Xenopus tadpoles. Using this approach, we can bypass early developmental effects of manipulating gene function allowing us to target specific tissues at later time points. Our studies show that heterogenous populations of craniofacial mesenchymal cells can be affected, allowing us to examine lineage of electroporated cells as well as cell autonomous requirements for proteins of interest. Combined with live imaging, we can use this approach to study gene function, over time, during craniofacial development. This novel method highlights the tractability of Xenopus for the study of organogenesis. We anticipate that this method can be broadly adapted to study morphogenesis and differentiation of other tissues as well.

A correction was made to <a href="http://http://www.jove.com/video/3381">Electroporation of Craniofacial Mesenchyme</a>. There was an incorrect unit used for the voltage.

Erratum: Electroporation of Craniofacial Mesenchyme

electroporation of craniofacial mesenchyme

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. For example, electroporation has been used with great success to study neural and retinal development in Xenopus, chicken and mouse 1-10. However, it is important to note that in all of these studies, investigators were not targeting soft tissues. Because we are interested in craniofacial development, we adapted a method to target facial mesenchyme.
When we searched the literature, we found, to our surprise, very few reports of successful gene transfer into cartilaginous tissue. The majority of these studies were gene therapy studies, such as siRNA or protein delivery into chondrogenic cell lines, or, animal models of arthritis 11-13. In other systems, such as chicken or mouse, electroporation of facial mesenchyme has been challenging (personal communications, Dept of Craniofacial Development, KCL). We hypothesized that electroporation into procartilaginous and cartilaginous tissues in Xenopus might work better. In our studies, we show that gene transfer into the facial cartilages occurs efficiently at early stages (28), when the facial primordium is still comprised of soft tissue prior to cartilage differentiation.
Xenopus is a very accessible vertebrate system for analysis of craniofacial development. Craniofacial structures are more readily visible in Xenopus than in any other vertebrate model, primarily because Xenopus embryos are fertilized externally, allowing analyses of the earliest stages, and facilitating live imaging at single cell resolution, as well as reuse of the mothers 14. Among vertebrate models developing externally, Xenopus is more useful for craniofacial analysis than zebrafish, as Xenopus larvae are larger and easier to dissect, and the developing facial region is more accessible to imaging than the equivalent region in fish. In addition, Xenopus is evolutionarily closer to humans than zebrafish (&tilde;100 million years closer) 15. Finally, at these stages, Xenopus tadpoles are transparent, and concurrent expression of fluorescent proteins or molecules will allow easy visualization of the developing cartilages. We anticipate that this approach will allow us to rapidly and efficiently test candidate molecules in an in vivo model system.

Watch this Scientific Journal Video about Electroporation of Craniofacial Mesenchyme at JoVE.com

Electroporation of Craniofacial Mesenchyme

Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Electroporation is an efficient method of delivering DNA and other charged macromolecules into tissues at precise time points and in precise locations. ...

electroporation-of-craniofacial-mesenchyme

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11.7K Views.  King's College London. Craniofacial cartilages develop in close contact with other tissues and are difficult to manipulate in live animals. We are using electroporation to deliver molecular tools during growth of the craniofacial skeleton while bypassing early embryonic effects. This approach will allow us to efficiently test candidate molecules in vivo.

Video: Electroporation of Craniofacial Mesenchyme

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High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly caused by the thick and waxy cell wall, but is compounded by the fact that most molecular techniques have been developed for distantly-related species such as Escherichia coli and Bacillus subtilis. In spite of these obstacles, mycobacterial plasmids have been identified and DNA transformation of many mycobacterial species have now been described. The most successful method for introducing DNA into mycobacteria is electroporation. Many parameters contribute to successful transformation; these include the species/strain, the nature of the transforming DNA, the selectable marker used, the growth medium, and the  conditions for the electroporation pulse. Optimized methods for the transformation of both slow- and fast-grower are detailed here. Transformation efficiencies for different mycobacterial species and with various selectable markers are reported.

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High efficiency transformation is a major limitation in the study of mycobacteria. The genus Mycobacterium can be difficult to transform; this is mainly ...

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant variants) in order to investigate their endogenous distribution and functional relevance. An interesting approach that has been implemented to fulfill this objective in fully differentiated cells is the in vivo transfection of plasmids by various methods into specific tissues such as liver1, skeletal muscle2,3, and even the brain4. We present here a detailed description of the steps that must be followed in order to efficiently transfect genetic material into fibers of the flexor digitorum brevis (FDB) and interosseus (IO) muscles of adult mice using an in vivo electroporation approach. The experimental parameters have been optimized so as to maximize the number of muscle fibers transfected while minimizing tissue damages that may impair the quality and quantity of the proteins expressed in individual fibers. We have verified that the implementation of the methodology described in this paper results in a high yield of soluble proteins, i.e. EGFP and ECFP3, calpain, FKBP12, &beta;2a-DHPR, etc. ; structural proteins, i.e. minidystrophin and &alpha;-actinin; and membrane proteins, i.e. &alpha;1s-DHPR, RyR1, cardiac Na/Ca2+ exchanger , NaV1.4 Na channel, SERCA1, etc., when applied to FDB, IO and other muscles of mice and rats. The efficient expression of some of these proteins has been verified with biochemical3 and functional evidence5. However, by far the most common confirmatory approach used by us are standard fluorescent microscopy and 2-photon laser scanning microscopy (TPLSM), which permit to identify not only the overall expression, but also the detailed intracellular localization, of fluorescently tagged protein constructs. The method could be equally used to transfect plasmids encoding for the expression of proteins of physiological relevance (as shown here), or for interference RNA (siRNA) aiming to suppress the expression of normally expressed proteins (not tested by us yet). It should be noted that the transfection of FDB and IO muscle fibers is particularly relevant for the investigation of mammalian muscle physiology since fibers enzymatically dissociated from these muscles are currently one of the most suitable models to investigate basic mechanisms of excitability and excitation-contraction coupling under current or voltage clamp conditions2,6-8.

DNA Transfection of Mammalian Skeletal Muscles using In Vivo Electroporation

We describe detailed procedures for the efficient transfection of plasmid DNA into the fibers of foot muscles of live mice using electroporation and the subsequent visualization of protein expression using fluorescence microscopy.

 A growing interest in cell biology is to express transgenically modified forms of essential proteins (e.g. fluorescently tagged constructs and/or mutant ...

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue interactions that regulate patterning and morphogenesis of vertebrates (e.g., 1, 2, 3, 4). Here, we illustrate a method that exploits this accessibility to test the signaling and patterning properties of ectodermal tissues during facial development. In these experiments, we create quail-chick 5 or mouse-chick 6 chimeras by transplanting the surface cephalic ectoderm that covers the upper jaw from quail or mouse onto either the same region or an ectopic region of chick embryos. The use of quail as donor tissue for transplantation into chicks was developed to take advantage of a nucleolar marker present in quail but not chick cells, thus allowing investigators to distinguish host and donor tissues 7. Similarly, a repetitive element is present in the mouse genome and is expressed ubiquitously, which allows us to distinguish host and donor tissues in mouse-chick chimeras 8. The use of mouse ectoderm as donor tissue will greatly extend our understanding of these tissue interactions, because this will allow us to test the signaling properties of ectoderm derived from various mutant embryos.

This article describes a tissue transplantation technique that was designed to test the signaling and patterning properties of surface cephalic ectoderm during craniofacial development.

The accessibility of avian embryos has helped experimental embryologists understand the fates of cells during development and the role of tissue ...

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the neural tube in a process known as neurulation. During neurulation, morphogenesis of the mesenchyme that underlies the neural plate is believed to drive neural fold elevation. The cranial mesenchyme is comprised of the paraxial mesoderm and neural crest cells. The cells of the cranial mesenchyme form a pourous meshwork composed of stellate shaped cells and intermingling extracellular matrix (ECM) strands that support the neural folds. During neurulation, the cranial mesenchyme undergoes stereotypical rearrangements resulting in its expansion and these movements are believed to provide a driving force for neural fold elevation. However, the pathways and cellular behaviors that drive cranial mesenchyme morphogenesis remain poorly studied. Interactions between the ECM and the cells of the cranial mesenchyme underly these cell behaviors. Here we describe a simple ex vivo explant assay devised to characterize the behaviors of these cells. This assay is amendable to pharmacological manipulations to dissect the signaling pathways involved and live imaging analyses to further characterize the behavior of these cells. We present a representative experiment demonstrating the utility of this assay in characterizing the migratory properties of the cranial mesenchyme on a variety of ECM components.

The cranial mesenchyme undergoes dramatic morphogenic movements that likely provides a driving force for elevation of the neural folds1,2. Here we describe a simple ex vivo explant assay to characterize the cellular behaviors of the cranial mesenchyme during neurulation. This assay has numerous applications including being amenable to pharmacological manipulations and live imaging analyses.

The central nervous system is derived from the neural plate that undergoes a series of complex morphogenetic movements resulting in formation of the ...

Genome-wide Gene Deletions in Streptococcus sanguinis by High Throughput PCR

Transposon mutagenesis and single-gene deletion are two methods applied in genome-wide gene knockout in bacteria 1,2. Although transposon mutagenesis is less time consuming, less costly, and does not require completed genome information, there are two weaknesses in this method: (1) the possibility of a disparate mutants in the mixed mutant library that counter-selects mutants with decreased competition; and (2) the possibility of partial gene inactivation whereby genes do not entirely lose their function following the insertion of a transposon. Single-gene deletion analysis may compensate for the drawbacks associated with transposon mutagenesis. To improve the efficiency of genome-wide single gene deletion, we attempt to establish a high-throughput technique for genome-wide single gene deletion using Streptococcus sanguinis as a model organism. Each gene deletion construct in S. sanguinis genome is designed to comprise 1-kb upstream of the targeted gene, the aphA-3 gene, encoding kanamycin resistance protein, and 1-kb downstream of the targeted gene. Three sets of primers F1/R1, F2/R2, and F3/R3, respectively, are designed and synthesized in a 96-well plate format for PCR-amplifications of those three components of each deletion construct. Primers R1 and F3 contain 25-bp sequences that are complementary to regions of the aphA-3 gene at their 5' end. A large scale PCR amplification of the aphA-3 gene is performed once for creating all single-gene deletion constructs. The promoter of aphA-3 gene is initially excluded to minimize the potential polar effect of kanamycin cassette. To create the gene deletion constructs, high-throughput PCR amplification and purification are performed in a 96-well plate format. A linear recombinant PCR amplicon for each gene deletion will be made up through four PCR reactions using high-fidelity DNA polymerase. The initial exponential growth phase of S. sanguinis cultured in Todd Hewitt broth supplemented with 2.5% inactivated horse serum is used to increase competence for the transformation of PCR-recombinant constructs. Under this condition, up to 20% of S. sanguinis cells can be transformed using ~50 ng of DNA. Based on this approach, 2,048 mutants with single-gene deletion were ultimately obtained from the 2,270 genes in S. sanguinis excluding four gene ORFs contained entirely within other ORFs in S. sanguinis SK36 and 218 potential essential genes. The technique on creating gene deletion constructs is high throughput and could be easy to use in genome-wide single gene deletions for any transformable bacteria.

Genome-wide Gene Deletions in Streptococcus sanguinis by High Throughput PCR

An efficient genome-wide single gene mutation method has been established using Streptococcus sanguinis as a model organism. This method has achieved via high throughput recombinant PCRs and transformations.

Transposon  mutagenesis and single-gene deletion are two methods applied in genome-wide  gene knockout in bacteria 1,2. Although transposon mutagenesis is ...

Orofacial clefts are the most frequent craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal facial development3. In human and mouse, initial growth and patterning of the face relies on several small buds of tissue, the facial prominences4,5. The face is derived from six main prominences: paired frontal nasal processes (FNP), maxillary prominences (MxP) and mandibular prominences (MdP). These prominences consist of swellings of mesenchyme that are encased in an overlying epithelium. Studies in multiple species have shown that signaling crosstalk between facial ectoderm and mesenchyme is critical for shaping the face6. Yet, mechanistic details concerning the genes involved in these signaling relays are lacking. One way to gain a comprehensive understanding of gene expression, transcription factor binding, and chromatin marks associated with the developing facial ectoderm and mesenchyme is to isolate and characterize the separated tissue compartments.
Here we present a method for separating facial ectoderm and mesenchyme at embryonic day (E) 10.5, a critical developmental stage in mouse facial formation that precedes fusion of the prominences. Our method is adapted from the approach we have previously used for dissecting facial prominences7. In this earlier study we had employed inbred C57BL/6 mice as this strain has become a standard for genetics, genomics and facial morphology8. Here, though, due to the more limited quantities of tissue available, we have utilized the outbred CD-1 strain that is cheaper to purchase, more robust for husbandry, and tending to produce more embryos (12-18) per litter than any inbred mouse strain8. Following embryo isolation, neutral protease Dispase II was used to treat the whole embryo. Then, the facial prominences were dissected out, and the facial ectoderm was separated from the mesenchyme. This method keeps both the facial ectoderm and mesenchyme intact. The samples obtained using this methodology can be used for techniques including protein detection, chromatin immunoprecipitation (ChIP) assay, microarray studies, and RNA-seq.

A protocol for separation of embryo facial ectoderm and mesenchyme is described. We use Dispase II to treat whole embryos first, dissect whole facial prominences out, and then separate the facial ectoderm and mesenchyme.

 Orofacial clefts are the most frequent  craniofacial defects, which affect 1.5 in 1,000 newborns worldwide1,2. Orofacial clefting is caused by abnormal ...

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, convergence and extension of facial prominences, and maturation of the craniofacial skeleton. To study the contribution of the cranial neural crest to specific regions of the zebrafish palate a sox10: kaede transgenic zebrafish line was generated. Sox10 provides lineage restriction of the kaede reporter protein to the neural crest, thereby making the cell labeling a more precise process than traditional dye or reporter mRNA injection. Kaede is a photo-convertible protein that turns from green to red after photo activation and makes it possible to follow cells precisely. The sox10: kaede transgenic line was used to perform lineage analysis to delineate CNC cell populations that give rise to maxillary versus mandibular elements and illustrate homology of facial prominences to amniotes. This protocol describes the steps to generate a live time-lapse video of a sox10: kaede zebrafish embryo. Development of the ethmoid plate will serve as a practical example. This protocol can be applied to making a time-lapse confocal recording of any kaede or similar photoconvertible reporter protein in transgenic zebrafish. Furthermore, it can be used to capture not only normal, but also abnormal development of craniofacial structures in the zebrafish mutants.

Visualization of experimental data has become a key element in presenting results to the scientific community. Generation of live time-lapse recording of growing embryos contributes to better presentation and understanding of complex developmental processes. This protocol is a step-by-step guide to cell labeling via photoconversion of kaede protein in zebrafish.

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, ...

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

Alternative Cultures for Human Pluripotent Stem Cell Production, Maintenance, and Genetic Analysis

Human pluripotent stem cells (hPSCs) hold great promise for regenerative medicine and biopharmaceutical applications. Currently, optimal culture and efficient expansion of large amounts of clinical-grade hPSCs are critical issues in hPSC-based therapies. Conventionally, hPSCs are propagated as colonies on both feeder and feeder-free culture systems. However, these methods have several major limitations, including low cell yields and generation of heterogeneously differentiated cells. To improve current hPSC culture methods, we have recently developed a new method, which is based on non-colony type monolayer (NCM) culture of dissociated single cells. Here, we present detailed NCM protocols based on the Rho-associated kinase (ROCK) inhibitor Y-27632. We also provide new information regarding NCM culture with different small molecules such as Y-39983 (ROCK I inhibitor), phenylbenzodioxane (ROCK II inhibitor), and thiazovivin (a novel ROCK inhibitor). We further extend our basic protocol to cultivate hPSCs on defined extracellular proteins such as the laminin isoform 521 (LN-521) without the use of ROCK inhibitors. Moreover, based on NCM, we have demonstrated efficient transfection or transduction of plasmid DNAs, lentiviral particles, and oligonucleotide-based microRNAs into hPSCs in order to genetically modify these cells for molecular analyses and drug discovery. The NCM-based methods overcome the major shortcomings of colony-type culture, and thus may be suitable for producing large amounts of homogeneous hPSCs for future clinical therapies, stem cell research, and drug discovery.

Here, we present human pluripotent stem cell (hPSC) culture protocols, based on non-colony type monolayer (NCM) growth of dissociated single cells. This new method, utilizing Rho-associated kinase inhibitors or the laminin isoform 521 (LN-521), is suitable for producing large amounts of homogeneous hPSCs, genetic manipulation, and drug discovery.

Human pluripotent stem cells (hPSCs) hold great promise for regenerative medicine and biopharmaceutical applications. Currently, optimal culture and ...