We thank Shlomi Krispin for the neural tube image. In particular, we acknowledge Yuval Cinnamon for initiating the GFP transfection technique. This work was supported by grants from the Israel Science Foundation, Association Francaise contre les Myopathies, and Deutche Forschungsgemeinshaft, Collaborative Research Centre 488 to C.K.

1. Preparation of Micropipettes
We pull our pipettes on a standard Narishige pp-83 pipette puller with a heat setting of 13.5. The precise setting must be calibrated, aiming at a pipette tip diameter of between 8 and 10 microns. We use filament-containing borosilicate glass tubes with an outer diameter of 1.0 mm and inner diameter of 0.75-0.78 mm. The pipettes are consequently stored in a plastic dish with the tips protected from contact with the edges of the dish.
2. DNA Preparation
The plasmid of interest is transfected into DH5&alpha; E. coli using standard procedures. Overnight cultures from single colonies are used to purify the plasmid using maxi-prep kits (any commercial kit should do). We use a plasmid concentration of 1.0 &mu;g/ &mu;L for non- clonal injections. Higher concentrations increase clogging of pipette tips. For clonal applications, plasmid concentrations are much lower and must be adjusted (see below). For pCAGG-GFP, the optimal concentration producing clonal results was found to be about 0.05-0.1 &mu;g/ &mu;L. One to 2 &mu;L of plasmid is typically used per experiment, and a minimal amount of fast-green powder (adhering to the tip of a dry, sharp glass pipette) is added to the plasmid before the experiment begins. The plasmid is then centrifuged in a tabletop centrifuge for 2 minutes at maximal speed. This procedure seems to reduce pipette-clogging problems.
3. Injection of Embryos
3.1 Preparation of embryos
We use quail as well as chicken eggs for injections. The eggs are incubated lying on their long axis until the desired developmental stage. The eggs are swabbed with 70% ethanol and allowed to dry. The pointed end of the egg is pierced with surgical scissors (this is necessary only in chicken eggs) and 1 mL (in quails) or 2 mL (chick) of albumen is drawn from the egg with a 10 mL syringe (19G needle). The holes are then sealed with hot paraffin.
Prior to actual injection, a window is opened at the top of the egg shell using surgical scissors. Placing sticky tape over the shell and cutting through it can help reduce debris from the shell. PBS buffer containing antibiotics is pipetted immediately into the egg. To access the embryo, the vitelline and/or amniotic membranes are cut and diverted using 0.14 mm insect pins mounted onto a needle holder. Non-toxic ink (Pelican #17) is injected underneath the blastoderm to help visualize the embryo, using a fire-pulled capillary mounted on an aspirator.
3.2 Loading and mounting of micropipettes
The puller micropipettes can be loaded with the plasmid by fire-pulled capillaries of adequate diameter, mounted onto an aspirator. A minimal amount of plasmid is drawn using the capillary and loaded into the pipette from its opening opposite to the sharpened tip, and approaching the tip from the inside as much as possible. The pipette is consequently loaded onto a manual air pressure injector mounted on a micromanipulator. Minimal pressure is applied, if necessary, in order to allow the plasmid solution to reach the tip of the pipette.
3.3 Plasmid injection
The pipette is manipulated into the liquid medium of the egg while applying some air pressure to prevent clogging of the pipette tip. The relevant tissue domain is pierced. The change in pressure as the tip enters the tissue is often enough to release plasmid with visible fast green. If this is not the case, apply slight pressure to the syringe until the plasmid/fast green are ejected. This process can be repeated along the axis of the embryo, for example in adjacent somites or along the extent of the neural tube. Following injections, PBS may be added. The window is sealed using sticky tape (parafilm is not desirable especially for quail eggs-it increases embryo death rates, probably by blocking exchange of oxygen through the shell). The treated embryos are returned to an incubator. Avoid ventilation and place small containers with water in the incubator as they help maintain moisture and increase yields. 
4. Adjusting Injection Parameters and Monitoring Success Rates
Asessing and calibrating various injection parameters is critical for applications aiming at single-cell analysis. Not every injection event will lead to successful labeling of a single cell. Plasmid concentration which is too high can lead to non-clonal labeling, while very low concentrations result in high failure rates. The concentration of the injected plasmid must be optimized for clonal injections. We recommend dilution of the entire purified plasmid stock rather than diluting individual preps to a pre-determined concentration. The latter might introduce some variance of success rates, probably due to variations in purity and other parameters such as the proportion of supercoiled plasmid.
4.1 Assessing success rates by whole mount observation
Initial assessment of injected embryos can be performed by whole-mount observation under a fluorescent binocular. Six to eight hours post-injection are sufficient to assess success rates with a pCAGG-enhanced GFP construct. This is provided the target site is superficial enough to be readily apparent (e.g. the dorsal somite, dorsal neural tube, ectoderm etc, see Figure 1A-C.). This step is particularly helpful for evaluating experimental series exhibiting either too many labeled cells or alternatively, lacking any fluorescent cells. If further incubation is desired, antibiotics-containing PBS solution should be added to the egg after viewing and the sealing tape should be replaced.
4.2 Assessing clonality by serial-section analysis
The best means for assessing the rates of successful injections is by serially-sectioning injected embryos and using immunohistochemistry to visualize the labeled cells. Under these conditions, incubation times can be as short as 4 hours post-injection. The embryos are fixed and processed for paraffin sectioning (or cryosectioning). Glass-mounted sections are then de-paraffinized and immuno-stained for reporter visualization. Amplification methods such as biotin-streptavidin might be desirable. Finally, the serial sections are scanned to detect labeled cells in injected segments. If upon initial trials multiple rather than single cells are obtained, the plasmid stock must be diluted and the process repeated until adequate results are achieved. Two to 10-fold dilutions of the plasmid stock solution may be necessary in cases of success rates being too high.
For each plasmid to be transfected, a thorough calibration that ensures clonality should be repeated.
5. Representative Results
We typically carry out a single injection of pCAGG-GFP per somite, while targeting 6-12 segments per embryo 1 2. Under this setting, a clear signal can be detected by whole mount observation 8 hours post injection, in at least one segment of one embryo out of ten. Under these conditions, labeling was further confirmed to be clonal (Figure 1B,C). When using high DNA concentrations, most embryos may exhibit at least one successful injection, raising the possibility that multiple cells have been transfected.
Serial section analysis revealed that clonal transfections were attained when the success rate was about 10%. This means that in order to achieve clonality, the technique is per definition, highly inefficient. On the other hand, under these conditions, more than 93% of positive labeling events were found to be clonal 1.
<img src="/files/ftp_upload/2133/2133fig1.jpg" alt="Figure 1" /> Figure 1. Clonal injections of pCAGGS-GFP into neural and skeletal progenitors. (A-C) Transverse sections showing labeling of the dorsal neural tube (NT) (A), ventrolateral domain of the dermomyotome (DM) (B), or central DM sheet (C), 6hr after microinjection of pCAGGS-GFP to a single progenitor. (D) Two days following clonal transfection of the central DM, single progenitors generated derivatives in both dermis (D) as well as muscle (M) (see ref.1 for details).

<ol>
	<li>Ben-Yair, R., Kalcheim, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lineage+analysis+of+the+avian+dermomyotome+sheet+reveals+the+existence+of+single+cells+with+both+dermal+and+muscle+progenitor+fates.">Lineage analysis of the avian dermomyotome sheet reveals the existence of single cells with both dermal and muscle progenitor fates.</a> Development. 132, 689-701 (2005).</li><li>Ben-Yair, R., Kalcheim, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Notch+and+bone+morphogenetic+protein+differentially+act+on+dermomyotome+cells+to+generate+endothelium,+smooth,+and+striated+muscle.">Notch and bone morphogenetic protein differentially act on dermomyotome cells to generate endothelium, smooth, and striated muscle.</a> J Cell Biol. 180, 607-618 (2008).</li><li>Cinnamon, Y., Kahane, N., Kalcheim, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterization+of+the+early+development+of+specific+hypaxial+muscles+from+the+ventrolateral+myotome.+.">Characterization of the early development of specific hypaxial muscles from the ventrolateral myotome. .</a> Development. 126, 4305-4315 (1999).</li><li>Stark, D. A., Kulesa, P. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Photoactivatable+green+fluorescent+protein+as+a+single-cell+marker+in+living+embryos.">Photoactivatable green fluorescent protein as a single-cell marker in living embryos.</a> Dev Dyn. 233, 983-992 (2005).</li><li>Haas, K., Jensen, K., Sin, W. C., Foa, L., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Targeted+electroporation+in+Xenopus+tadpoles+in+vivo--from+single+cells+to+the+entire+brain.">Targeted electroporation in Xenopus tadpoles in vivo--from single cells to the entire brain.</a> Differentiation. 70, 148-154 (2002).</li><li>Haas, K., Sin, W. C., Javaherian, A., Li, Z., Cline, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single-cell+electroporation+for+gene+transfer+in+vivo.">Single-cell electroporation for gene transfer in vivo.</a> Neuron. 29, 583-591 (2001).</li><li>Hewapathirane, D. S., Haas, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+electroporation+in+vivo+within+the+intact+developing+brain.">Single cell electroporation in vivo within the intact developing brain.</a> J Vis Exp. , (2008).</li><li>Nagai, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+variant+of+yellow+fluorescent+protein+with+fast+and+efficient+maturation+for+cell-biological+applications.">A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications.</a> Nat Biotechnol. 20, 87-90 (2002).</li><li>Sato, Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+integration+and+conditional+expression+of+electroporated+transgenes+in+chicken+embryos.">Stable integration and conditional expression of electroporated transgenes in chicken embryos.</a> Dev Biol. 305, 616-624 (2007).</li></ol>

No conflicts of interest declared.

The method described above is a modification of a previously described technique for delivering lipophilic dyes to small cell subsets or single progenitors 3. It has three main advantages over the standard electroporation technique. First, it provides a means for labeling with confidence discrete sites in target tissues, allowing much greater spatial resolution than electroporation (see for example Figure 1B,C). Second, it permits labeling of single cells, thus allowing to trace their lineage in-vivo ...

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single cell transfection in chick embryos

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a thorough analysis of the fates of single cells under normal and experimental conditions. To this end, transfection methods that target single progenitors are a prerequisite. We describe here a technically straightforward method for transfecting single cells in chicken tissues in-ovo, allowing reliable lineage tracing as well as genetic manipulation. Specific tissue domains are targeted within the somite or neural tube, and DNA is injected directly into the epithelium of interest, resulting in sporadic transfection of single cells. Using reporters, clonal populations may consequently be traced for up to three days, and behavior of genetically manipulated clonal populations can be compared with that of controls. This method takes advantage of the accessibility of the chick embryo along with emerging tools for genetic manipulation. We compare and discuss its advantages over the widely-used electroporation method, and possible applications and use in additional in-vivo models are also suggested. We advocate the use of this method as a significant addition and complement for existing lineage tracing and genetic interference tools.

Watch this Scientific Journal Video about Single Cell Transfection in Chick Embryos at JoVE.com

Single Cell Transfection in Chick Embryos

Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations.

A central theme in developmental biology is the diversification of lineages and the elucidation of underlying molecular mechanisms. This entails a ...

single-cell-transfection-in-chick-embryos

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11.1K Views.  Hadassah Medical School - Hebrew University. Using fine tip micropipettes we inject plasmid DNA into subdomains of chicken somites or neural tubes. The concentration of the plasmid is adjusted to generate single transfected cells. We then allow the cells to develop into clonal populations. 

Video: Single Cell Transfection in Chick Embryos

Dissection of 6.5 dpc Mouse Embryos

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell interaction and signaling mechanisms that guide embryonic patterning. However, dissection of the mouse embryo from the decidua shortly after implantation can be a challenging procedure, and detailed step-by-step documentation of this process is lacking.

Here we demonstrate how post-implantation (6.5 dpc) embryos are isolated by first dissecting the uterus of a pregnant mouse (detection of the vaginal plug was designated day 0.5 poist coitum) and subsequently dissecting the embryo from maternal decidua. The dissection of Reichert's membrane is described as well as the removal of the ectoplacental cone.

Isolation of postimplantation-stage embryos allows one to study gene patterning and analyze cell-lineage decision making processes during embryonic development, but proper dissection of the early embryo can be challenging. This protocol describes a method for isolating early primitive-streak-stage embryos (~6.5 days post coitum [dpc]).

Analysis of gene expression patterns during early stages of mammalian embryonic development can provide important clues about gene function, cell-cell ...

In Ovo Electroporations of HH Stage 10 Chicken Embryos

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of molecular biological techniques, the chick has become a useful system in which to study gene regulation and function.  By electroporating DNA or RNA constructs into the developing chicken embryo, genes can be expressed or knocked down in order to analyze in vivo gene function.  Similarly, reporter constructs can be used for fate mapping or to examine putative gene regulatory elements.  Compared to similar experiments in mouse, chick electroporation has the advantages of being quick, easy and inexpensive.  This video demonstrates first how to make a window in the eggshell to manipulate the embryo.  Next, the embryo is visualized with a dilute solution of India ink injected below the embryo.  A glass needle and pipette are used to inject DNA and Fast Green dye into the developing neural tube, then platinum electrodes are placed parallel to the embryo and short electrical pulses are administered with a pulse generator.  Finally, the egg is sealed with tape and placed back into an incubator for further development.  Additionally, the video shows proper egg storage and handling and discusses possible causes of embryo loss following electroporation.

Chick in ovo electroporation is a technique which allows genetic manipulation of the avian embryo. Common applications of this technique include functional analysis of genes and putative enhancer elements. This video demonstrates neural tube electroporation in HH 10 chick embryos. Injection technique and proper egg handling are discussed.

Large size and external development of the chicken embryo have long made it a valuable tool in the study of developmental biology.  With the advent of ...

Method for Culture of Early Chick Embryos ex vivo (New Culture)

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  the formation and patterning of brain, neural tube, somite and heart primordia. Applications of chick embryo culture include electroporation of DNA or RNA constructs in order to analyze gene function, grafts of growth factor coated beads such as FGFs and BMPs , as well as whole mount in situ hybridization and immunohistochemistry. This video demonstrates the different steps in chick embryo culture; First, the embryo is explanted in saline. Then, the embryo is centered on a glass ring. The membranes surrounding the embryo are lifted along the walls of the ring. The ring is then placed in a culture dish containing a pool of albumine. The culture dish is sealed and placed in a humid chamber, where the embryo is cultured for up to 24 hrs. Finally, the embryo is removed from the ring, fixed and processed for further applications. A troubleshooting guide is also presented.

Method for Culture of Early Chick Embryos ex vivo New Culture

This video demonstrates New culture, a method by which chick embryos are cultured outside the egg for up to 24 hr. This method enables one to study early development (primitive streak to 14 som.), a period corresponding to E7-9 in mouse. Applications of this technique include electroporation, in situ hybridization and immunohistochemistry.

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ...

Double Whole Mount in situ Hybridization of Early Chick Embryos

The chick embryo is a valuable tool in the study of early embryonic development. Its transparency, accessibility and ease of manipulation, make it an ideal tool for studying  gene expression in brain, neural tube, somite and heart primordia formation. This video demonstrates the different steps in 2-color whole mount in situ hybridization; First, the embryo is dissected from the egg and fixed in paraformaldehyde. Second, the embryo is processed for prehybridization. The embryo is then hybridized with two different probes, one coupled to DIG, and one coupled to FITC.  Following overnight hybridization, the embryo is incubated with DIG coupled antibody. Color reaction for DIG substrate is performed, and the region of interest appears blue. The embryo is then incubated with FITC coupled antibody. The embryo is processed for color reaction with FITC, and the region of interest appears red. Finally, the embryo is fixed and processed for phtograph and sectioning. A troubleshooting guide is also presented.

This video demonstrates 2-color whole mount in situ hybridization, a method by which the spatial and temporal expression pattern of 2 different genes can be visualized in young chick embryos. This method was originally introduced by David Wilkinson, Domingos Henrique, Phil Ingham and David Ish -Horowicz.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment are open and allow cations to flow steadily inwards across the membrane, depolarizing the cell. Light exposure triggers the closure of the CNG channels, blocks the inward cation current flow, and thus results in cell hyperpolarization. Based on the polarity of photoreceptors, a suction recording method was developed in 1970s that, unlike the classic patch-clamp technique, does not require penetrating the plasma membrane 1. Drawing the outer segment into a tightly-fitting glass pipette filled with extracellular solution allows recording the current changes in individual cells upon test-flash exposure. However, this well-established "outer-segment-in (OS-in)" suction recording is not suitable for mouse cone recordings, because of the low percentage of cones in the mouse retina (3%) and the difficulties in identifying the cone outer segments. Recently, an inner-segment-in (IS-in) recording configuration was developed to draw the inner segment/nuclear region of the photoreceptor into the recording pipette 2,3. In this video, we will show how to record from individual mouse cone photoresponses using single-cell suction electrode.

We will show how to record flash responses from single mouse cones using a suction electrode.

Rod and cone photoreceptors in the retina are responsible for light detection. In darkness, cyclic nucleotide-gated (CNG) channels in the outer segment ...

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic development, cellular signaling and adult physiology. In this respect, the use of multi-photon microscopy enables quantitative photoactivation of a given light responsive agent in deep tissues at a single cell resolution. As zebrafish embryos are optically transparent, their development can be monitored in vivo. These traits make the zebrafish a perfect model organism for controlling the activity of a variety of chemical agents and proteins by focused light. Here we describe the use of two-photon microscopy to induce the activation of chemically caged fluorescein, which in turn allows us to follow cell's destiny in live zebrafish embryos. We use embryos expressing a live genetic landmark (GFP) to locate and precisely target any cells of interest. This procedure can be similarly used for precise light induced activation of proteins, hormones, small molecules and other caged compounds.

Multiphoton microscopy allows control of low energy photons with deep optical penetration and reduced phototoxicity. We describe the use of this technology for live cell labeling in zebrafish embryos. This protocol can be readily adapted for photo-induction of various light-responsive molecules.

Photoactivation of target compounds in a living organism has proven a valuable approach to investigate various biological processes such as embryonic ...

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

Single Cell Fate Mapping in Zebrafish

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, lymphatic, and blood vessel lineages arise in developing embryos requires fate mapping and lineage tracing of undifferentiated precursor cells. Recently, photoactivatable proteins which include: Eos1, 2, PAmCherry3, Kaede4-7, pKindling8, and KikGR9, 10 have received wide interest as cell tracing probes. The fluorescence spectrum of these photosensitive proteins can be easily converted with UV excitation, allowing a population of cells to be distinguished from adjacent ones. However, the photoefficiency of the activated protein may limit long-term cell tracking11. As an alternative to photoactivatable proteins, caged fluorescein-dextran has been widely used in embryo model systems7, 12-14. Traditionally, to uncage fluorescein-dextran, UV excitation from a fluorescence lamp house or a single photon UV laser has been used; however, such sources limit the spatial resolution of photoactivation. Here we report a protocol to fate map, lineage trace, and detect single labeled cells. Single cells in embryos injected with caged fluorescein-dextran are photoactivated with near-infrared laser pulses produced from a titanium sapphire femtosecond laser. This laser is customary in all two-photon confocal microscopes such as the LSM 510 META NLO microscope used in this paper. Since biological tissue is transparent to near-infrared irradiation15, the laser pulses can be focused deep within the embryo without uncaging cells above or below the selected focal plane. Therefore, non-linear two-photon absorption is induced only at the geometric focus to uncage fluorescein-dextran in a single cell. To detect the cell containing uncaged fluorescein-dextran, we describe a simple immunohistochemistry protocol16 to rapidly visualize the activated cell. The activation and detection protocol presented in this paper is versatile and can be applied to any model system. 
Note: The reagents used in this protocol can be found in the table appended at the end of the article.

A method is described to photoactivate single cells containing a caged fluorescent protein using two-photon absorption from a Ti:Sapphire femtosecond laser oscillator. To fate map the photoactivated cell, immunohistochemistry is used. This technique can be applied to any cell type.

The ability to differentially label single cells has important implications in developmental biology. For instance, determining how hematopoietic, ...

Facial Transplants in Xenopus laevis Embryos

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. To identify where signaling pathways and tissues act during patterning of the developing face, a 'face transplant' technique has been developed in embryos of the frog Xenopus laevis. A region of presumptive facial tissue (the "Extreme Anterior Domain" (EAD)) is removed from a donor embryo at tailbud stage, and transplanted to a host embryo of the same stage, from which the equivalent region has been removed. This can be used to generate a chimeric face where the host or donor tissue has a loss or gain of function in a gene, and/or includes a lineage label. After healing, the outcome of development is monitored, and indicates roles of the signaling pathway within the donor or surrounding host tissues. Xenopus is a valuable model for face development, as the facial region is large and readily accessible for micromanipulation. Many embryos can be assayed, over a short time period since development occurs rapidly. Findings in the frog are relevant to human development, since craniofacial processes appear conserved between Xenopus and mammals.

Facial Transplants in Xenopus laevis Embryos

A technique for transplanting "Extreme Anterior Domain" facial tissue between Xenopus laevis embryos has been developed. Tissue can be moved from one gene expression background into another, allowing the study of local requirements for craniofacial development and for signaling interactions between facial regions.

Craniofacial birth defects occur in 1 out of every 700 live births, but etiology is rarely known due to limited understanding of craniofacial development. ...

Single Cell Transfection in Chick Embryos

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Rozdziały w tym wideo

Dissection of 6.5 dpc Mouse Embryos

In Ovo Electroporations of HH Stage 10 Chicken Embryos

Method for Culture of Early Chick Embryos ex vivo (New Culture)

Double Whole Mount in situ Hybridization of Early Chick Embryos

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

Single-cell Suction Recordings from Mouse Cone Photoreceptors

Two-Photon-Based Photoactivation in Live Zebrafish Embryos

Imaging Cell Shape Change in Living Drosophila Embryos

Single Cell Fate Mapping in Zebrafish

Facial Transplants in Xenopus laevis Embryos