We thank Filip Wymeersch and Anestis Tsakiridis for comments on the manuscript, staff in the SCRM animal unit for help with animal maintenance and Prof. Stuart Forbes for immunohistochemistry reagents. This work was supported by MRC grant Mr/K011200/1 and the China Scholarship Council

All animal experiments were carried out in accordance with UK Home Office Regulations as specified in the Animals (Scientific Procedures) Act (1986) under Project Licence number 60/4435. To collect embryos at specific developmental stages, timed matings were set up O/N. Noon on the day of finding a vaginal plug was designated embryonic day (E) 0.5.

1. Dissecting the E7.5 or E8.5 Postimplantation Embryos for Ex Vivo Culture
<ol>
	<li>Sacrifice the pregnant female mice by cervical dislocation.</li>
	<li>Isolate the uterus using scissors, holding it with forceps, and place in a 30 mm dish filled with M2 medium.</li>
	<li>Carefully tear apart the myometrium using two pairs of fine forceps.</li>
	<li>Peel away the decidua, being careful not to puncture the extraembryonic cavities.</li>
	<li>Remove the Reichert&#8217;s membrane by pinching it with forceps and slowly separating it from the embryo.</li>
	<li>Check the embryos under a dissecting stereomicroscope to ensure that the yolk sac, amnion and ectoplacental cone are intact.</li>
	<li>Transfer the embryos to a clean dish of M2 using a pipette and place on a 30 mm plastic Petri dish lid on ice (the &#8216;ice platform&#8217;) to partially chill embryos.</li>
	<li>If required, store embryos in M2 on an ice platform for up to 1.5 hr, e.g., during media preparation or manipulation of small batches of ~3-4 embryos at RT. 
	Note: Recovery and dissection of rodent embryos has been described in detail previously7,8,16.</li>
</ol>
2. Prepare the Embryo Culture Medium
<ol>
	<li>Freshly thaw either commercially-available rat serum (see Glanville-Jones et al.13 for specifications), or rat serum prepared in-house according to Copp and Cockroft16 that was heat inactivated for 30 min&#160;at 56 &#730;C and frozen in 1 ml aliquots at -80 &#730;C 
	Note: Commercially-available rat serum is acceptable for culture periods of 24-36 hr, although serum prepared in-house is, in our experience, superior for culture periods up to 48 hr.</li>
	<li>Freshly prepare an excess (e.g., 10 ml) of defined supplements consisting of Glasgow Minimum Essential Medium (GMEM), 1% Non-Essential Amino Acids (NEAA), 2 mM L-glutamine and 1 mM sodium pyruvate.</li>
	<li>Calculate the volume of culture medium that is needed according to the number of embryos of each stage that are going to be cultured (see section 3). Mix the rat serum with the defined supplements to make up 50% culture medium (1:1 (v/v) rat serum:defined supplements) and/or 75% culture medium (3:1 (v/v) rat serum: defined supplements).</li>
	<li>Pass the embryo culture medium through a 0.45 &#181;m filter and add 10,000 I.U/ml penicillin and 10 &#181;g/ml streptomycin. 
	Note: Freshly thawed L-glutamine and sodium pyruvate solution is crucial for embryo development during ex vivo culture.</li>
</ol>
3. Ex Vivo Culture Conditions for Different Embryonic Stages
<ol>
	<li>For E7.5 embryos: static culture using 4-well plates in an incubator supplied with 5% CO2 in air at 37 &#730;C for 24 hr. Culture up to two embryos per well in 1 ml of 50% culture medium.</li>
	<li>For E8.5 embryos: use a roller culture apparatus8 rotating at 35 revs/min incorporating continuous gassing with 5% CO2 in air at 37 &#730;C for 24 hr (1 ml of 50% culture medium per embryo).</li>
	<li>For E9.5 embryos: use a roller culture apparatus rotating at 35 revs/min incorporating gases supplied with 5% CO2, 40% O2 55% N2 at 37 &#730;C for 24 hr (1 ml of 75% culture medium per embryo). 
	Note: Culture embryos within 3 hr after euthanizing the mice as prolonged periods in M2 adversely affect development. See Copp and Cockroft16 for the method of ex vivo embryo culture.</li>
</ol>
4. Grafting Cultured Cells into E7.5 or E8.5 Mouse Embryos
<ol>
	<li>Physically scrape EpiSCs, which ubiquitously express GFP, from a 6-well culture plate using a 20-200 &#181;l pipette tip and place them in a 30 mm dish containing the embryos 
	Note: To insert cell clumps in the embryos, cells should be physically scrapped rather than trypsinized.</li>
	<li>Attach a hand-pulled grafting capillary to the aspirator tube to make a mouth pipette.</li>
	<li>Gently suck the mouth pipette to draw one or more cell clumps of size &#62;20 cells into the grafting capillary.</li>
	<li>Gently blow out the cells, to partially disperse large clumps.</li>
	<li>Select one cell clump containing ~10-20 cells and suck into the grafting capillary again, keeping it close to the opening of the capillary. Be careful not to move the cell clump in and out of the capillary repeatedly, to avoid break-up into smaller pieces.</li>
	<li>Hold the embryo loosely in place with a pair of forceps and insert the grafting capillary into the region of interest to create an opening.</li>
	<li>Gently expel the clump out of the grafting capillary, leaving the short string of 10-20 cells lodged in the embryo.</li>
	<li>Repeat the grafting procedure for the desired number of embryos. Use batch sizes of 3-4 embryos for convenience.</li>
	<li>Leaving embryos in the same dish of M2 medium, image the grafted embryos using a fluorescence compound dissecting microscope with camera, keeping imaging time to a minimum to avoid exposure of the embryos to excessive light and heat. 
	Note: Determine imaging times empirically as these will depend on the specifications of the camera and microscope, as well as the nature and fluorescence intensity of the fluorophore.</li>
	<li>Transfer the embryo in a pastette with a minimal volume of M2 medium to pre-equilibrated culture medium (see section 3) immediately after imaging. 
	Note: Carefully examine the morphology of the embryos after grafting. Only culture the intact embryos.</li>
</ol>
5. Handmade Electroporation Materials and Apparatus Setup (Prepare the Following in Advance of Electroporation Experiments):
<ol>
	<li>For DNA injection pipettes: pull DNA injection pipettes using a horizontal micropipette puller. Injection pipettes should have a fine tip with an opening less than 10 &#181;m to avoid tissue damage when injecting DNA into the embryonic cavities.</li>
	<li>For glass capillary electroporation: use a microforge to cut the opening of DNA injection pipettes to an internal diameter of either 20 or 30 &#181;m. To avoid cellular damage when the capillary is in contact with the embryo for electroporation, the tip of the glass capillary must be cut cleanly and not contain sharp, broken edges.</li>
	<li>For handmade capillary electrode (anode) (Figure 1): insert a 0.2 mm diameter platinum wire into an electroporation glass capillary with a fixed opening of 20 or 30 &#181;m diameter to focus the electric current and deliver the plasmid DNA to a small region of interest in the embryo.</li>
	<li>For handmade L-shaped electrode (cathode) (Figure 1): bend a 0.2 mm diameter platinum wire creating an &#8220;L&#8221; shape, with the horizontal part of the &#8220;L&#8221; around 1 mm in length.</li>
	<li>Attach each platinum electrode to a thin insulated wire and insert it into a microinjection needle holder covered by insulation tape.</li>
	<li>Mount the needle holders on standard micromanipulation instrument holders.</li>
	<li>Connect the circuit as shown in Figure 1: connect the capillary electrode to the anode of the power supply; connect the L-shaped electrode to the anode of the multimeter; connect the cathode of the mulimeter to the cathode of the power supply.</li>
</ol>
6. Electroporation E7.5 or E8.5 Mouse Embryos
<ol>
	<li>Fill the electroporation glass capillary with PBS to within 1-2 mm of the top and insert the straight platinum electrode (anode) into the glass capillary until it reaches the bottom of the capillary.</li>
	<li>Anchor the L-shaped electrode (cathode) on the surface of a 30 mm petri dish filled with PBS.</li>
	<li>Transfer the embryo from M2 medium into the PBS-filled electroporation dish.</li>
	<li>Insert the injection needle from lateral epiblast into the amniotic cavity of the embryo. Using a pneumatic pico pump, inject DNA solution (pCAG-Cre:GFP or pCAG-GFP 1-1.5 &#181;g/ml with 0.01% green food coloring dye) into the cavity until it is completely full. For E7.5-E8.5 embryos, less than 5 &#181;l DNA solution is required for one embryo. Using the green dye as an indicator, be careful not to burst the embryo.</li>
	<li>Carefully position the embryo between the electrodes and move the capillary electrode to the precise location where the DNA is to be delivered. 
	Note: The orientation of the embryo depends on the region where the DNA is to be electroporated.</li>
	<li>Electroporate the embryo using 200 volts (V) in 6 pulses, each of 50 ms duration with a 1 sec interval between each pulse.</li>
	<li>Transfer the embryo to pre-equilibrated culture medium immediately after electroporation. If desired, repeat the process for the next embryo. 
	Note: Add the culture medium into a sterile container and put it in the culture incubator to pre-equilibrate the medium.</li>
	<li>To detect electroporated cells 2 hr after culture, transfer the embryos to a clean 30 mm Petri dish of M2 medium using a pastette. Image the embryos as in step 5.9 using a fluorescence compound dissecting microscope.</li>
	<li>Transfer the embryos back to culture using a pastette immediately after imaging.</li>
	<li>To detect dead cells caused by electroporation, stain the embryos with a far-red cell membrane-impermeable nuclear dye (1:200 in embryo culture medium) at 37 &#730;C for 10 min&#160;2 hr after electroporation (Optional)</li>
	<li>To count the electroporated cells, fix the embryos in 4% paraformaldehyde (PFA) for 2-4 hr at 4 &#730;C, stain the nucleus with an ultraviolet or far-red fluorescent nuclear counterstain and image the embryos using a confocal microscope (Optional). 
	Note: Embryonic growth is adversely affected if left too long in PBS. Therefore, ensure that the time taken to electroporate each embryo is minimized (&#60;5 min per embryo).</li>
</ol>

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J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+cell+lines+from+mouse+epiblast+share+defining+features+with+human+embryonic+stem+cells.">New cell lines from mouse epiblast share defining features with human embryonic stem cells.</a> Nature. 448, 196-199 (2007).</li><li>Huang, Y., Osorno, R., Tsakiridis, A., Wilson, V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+Vivo+differentiation+potential+of+epiblast+stem+cells+revealed+by+chimeric+embryo+formation.">In Vivo differentiation potential of epiblast stem cells revealed by chimeric embryo formation.</a> Cell rep. 2, 1571-1578 (2012).</li><li>Sadik, M. 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The authors have no conflicts of financial or other interest to declare.

Grafting
The critical step for cell grafting experiments is the insertion of a coherent string of cells ideally in a single action, to avoid breakup of the clump. This technique requires some practice in controlling the mouth pipette. If donor cells incorporate well in the host, their derivatives will disperse in the embryo. To further determine whether the dispersed donor derived-cells differentiate appropriately in the host, immunostaining can be performed on the embryo sections. If donor cells are not compatible with the host environment, they either cannot be detected (as they are expelled from the embryo) or form unincorporated clumps in the embryos after culture. If both dispersed cells and cell clumps were observed, this may indicate that too many cells were grafted and excessive donor cells which cannot interact with surrounding host cells resulted in clump formation. In this case, additional grafts containing a smaller number of cells can be performed.
The major limitation of the cell grafting technique is that it is not possible to determine the full in vivo potential of cells since mouse ex vivo culture over periods longer than 48 hr has not been achieved. However, if combined with ultrasound-guided cell injection, it may be possible to transfer cultured cells to the embryos in utero. To summarize, cell grafting experiments have been widely used in our group and have given us valuable clues about the in vivo potential of various cell types4,21,22. It is a technique of general utility to assess the in vivo potential of in vitro cultured cells in early postimplantation embryos.
Electroporation
Although in this study we have only shown that it is efficient to use the capillary electroporation technique to target the epiblast, it is also possible to intentionally target other germ layers such as endoderm cells. The critical step for the capillary electroporation technique is to minimize the time taken to electroporate each embryo (&#60; 5 min per embryo) since PBS is very suboptimal for early mouse embryos. Our data above has shown that, in most areas in the embryos, electroporation does not affect the embryo growth. However, electroporation in the node caused abnormal development and led to the premature death of the embryo. This is likely due to damage or death of the cells that form important signalling centres23. Hence, this region would have to be avoided with this technique. A further caveat is that, as mentioned in the results section, whilst epiblast or primitive streak cells were targeted, some endoderm cells were also electroporated. This may be because DNA reaches to the endoderm through gaps under the epiblast epithelium. Endoderm is composed of epithelial cells and in our experience these cells have a higher propensity to take up DNA. Therefore, when applying this technique for fate mapping, it is important to assess which cells initially take up DNA.
It should also be noted that although pCAG-GFP and pCAG-Cre:GFP plasmids can be efficiently delivered using the electroporation parameters shown in this study, the efficiency of other DNA constructs may vary and need individual optimisation. Alterations in DNA concentration, electroporation voltage or the number of pulses can be made if plasmids are found to be difficult to transfect.
To summarize, our optimized capillary electroporation system can efficiently and reproducibly deliver GFP or Cre:GFP plasmids into a very few cells in the embryo with limited cell death. Since this method does not require expensive or highly specialized equipment, it can be of great use for cell tracking studies or in testing the effect of ectopic expression or conditional deletion of genes in early embryos, if electroporation is performed in embryos carrying floxed conditional mutant alleles. Therefore, this electroporation technique provides a useful functional tool for understanding on a cell-by-cell basis the roles of cell-intrinsic factors in the context of localized wildtype embryonic environments.

Cell culture has been widely used in developmental biology studies. Mouse embryonic stem cells (ESCs) and epiblast stem cells (EpiSCs) can differentiate into all three germ layers in vitro and are a useful model for cell differentiation in early mammalian embryogenesis. The derivation of these cell lines has opened up an opportunity for in vitro manipulation and detailed investigation of the localized signaling events and transcriptional networks operating during early embryonic patterning. However, it remains important to determine the in vivo relevance of any manipulations performed in culture. The in vivo potential of preimplantation embryo-derived mouse ESCs has been assessed by introducing them back into preimplantation embryos (morulae or blastocysts)1. However, EpiSCs that represent the epiblast cells in postimplantation embryos cannot integrate efficiently in preimplantation embryos2,3. Our previous findings have shown that EpiSCs can efficiently generate chimeras and contribute to all germ layers, when grafted into postimplantation embryos4. Thus, the best way to evaluate the in vitro cultured cells is to introduce them to their corresponding environment in vivo.
Electroporation is a widely used method to deliver exogenous molecules into targeted cells in both in vivo and in vitro experiments. Electrical energy can generate a large number of pores in the cell membrane, which allows exogenous deoxyribonucleic acid (DNA) or ribonucleic acid (RNA) to enter the cells. One of the biggest challenges for this technique is to combine optimal cell viability with high electrotransfection efficiency 5,6. For electroporation of nucleic acids in embryonic tissues, gold plated electrodes have most commonly been used, allowing targeting of cells in a broad spatial range7-9. To achieve a more localized gene transfer, a needle-shaped electrode has been utilized to achieve a focal electric field10,11. Using this method, the authors showed that after the electroporation, around 30-60 cells had taken up the DNA construct11. Nevertheless, it seems that accurately adjusting the number of electroporated cells remains difficult with a fixed-width electrode. The capillary electroporation technique has been used to deliver plasmids to single cells12-14. However, this technique has not been applied for electroporating plasmids to embryos ex vivo. More recently, a microdevice has been reported to locally electroporate a few distal visceral endoderm cells (less than 4 cells) in early postimplantation mouse embryos15. However, it is still unknown whether this device can efficiently target ectoderm and mesoderm ex vivo.
In this study we describe two novel methods to assess cellular and gene function in early post-implantation embryos. We first demonstrate how to graft in vitro cultured cells into defined sites in early mouse embryos to assess their in vivo potential. The integration of the grafted cells and their descendants, all labelled by a genetic tag (e.g., a green fluorescent protein (GFP), can be further examined by immunostaining of tissue specific proteins4. Secondly, we describe an improved method to precisely deliver DNA to localized sites in the embryo via electroporation. Rather than using a needle-shaped electrode, we inserted a thin wire inside a fine-tipped glass capillary, and demonstrate that this modification can deliver DNA to a small number of cells with high efficiency and limited cell death. Moreover, we show that by using glass capillaries with different opening sizes, we can control the number of electroporated cells. Therefore, we believe this method can be of great use to study early embryonic patterning involving small numbers of cells.

<table border="0" cellpadding="0" cellspacing="0" width="1172">
	<tbody>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Forceps</td>
			<td style="width:275px;">Dumostar</td>
			<td style="width:275px;">T5390</td>
			<td style="width:275px;"></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Dissecting stereomicroscope&#160;</td>
			<td style="width:275px;">&#160;Zeiss</td>
			<td style="width:275px;">Stemi 2000-C</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Stereomicroscope system with fluorescence&#160;</td>
			<td style="width:275px;">Nikon</td>
			<td style="width:275px;">AZ100</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Inverted microscope with a digital camera</td>
			<td style="width:275px;">Olympus</td>
			<td style="width:275px;">Olympus BX61</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Inverted confocal microscope&#160;</td>
			<td style="width:275px;">Leica Microsystems</td>
			<td style="width:275px;">Leica TCS SP8</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Low melting point agarose&#160;</td>
			<td style="width:275px;">Life Technologies</td>
			<td style="width:275px;">16520-050</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:347px;">Pasteur pipettes</td>
			<td style="width:275px;">Fisher Scientific</td>
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methods for precisely localized transfer of cells or dna into early postimplantation mouse embryos

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, cell culture has been widely used in developmental biology studies. However, it is important to determine whether in vitro cultured cells truly represent in vivo cell types. Grafting cells into embryos, followed by an assessment of their contribution during development is a useful method to determine the potential of in vitro cultured cells. In this study, we describe a method for grafting cells into a defined site of early postimplantation mouse embryos, followed by ex vivo culture. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing precise localization and adjustment of the number of cells receiving exogenous DNA with both high transfection efficiency and low cell death. These techniques, which do not require any specialized equipment, render experimental manipulations of the gastrulation and early organogenesis-stage mouse embryo possible, allowing analysis of commitment in cultured cell subpopulations and the effect of genetic manipulations in situ on cell differentiation.

Grafting
EpiSCs that ubiquitously express EGFP (r04-GFP, derived from E6.5 epiblast, and C2, derived in vitro from mESCs)4 were manually scraped from the culture dish and grafted into different sites of E7.5 embryos (Figure 2A). The embryos were cultured ex vivo and analysed after 24 hr. The distribution of donor cells was assessed by fluorescence microscopy. If donor cells incorporated, they proliferated and their derivatives dispersed within the host embryos (Figure 2B). It has been observed that grafts containing 10-16 cells incorporated efficiently in the host embryos (Figure 2A, and 2B), however, grafting more cells does not result in better chimaerism. Instead, grafted cells produced unincorporated clumps (Figure 2C and 2D).
Electroporation
To assess the efficiency of our electroporation system, we delivered GFP-expressing plasmids (pCAG-GFP and pCAG-Cre:GFP) to specific sites in the embryo. Consistent with a previous study11, GFP+ cells were detected in embryos 1-2 hr after electroporation (Figure 2E and 2G). When distal epiblast cells at the late primitive streak stage embryo were electroporated, labelled cells contributed to the neural ectoderm after 24 hr in culture (Figure 2E and 2F). This result corresponds well to known fate maps of epiblast cells in gastrulation stage embryos17. Similarly, when the GFP expression plasmid was electroporated in the primitive streak at E8.5 (2-5 somites), GFP+ cells contributed to the paraxial mesoderm (Figure 2G and 2H), consistent with known fate maps of the late primitive streak18. Moreover, we observed contribution to all three germ layers from electroporated cells (Figure 2I-K), suggesting that the electroporation procedure does not compromise cell behaviour in vivo. However, we also noticed that whilst epiblast (E7.5) or primitive streak cells (E8.5) were targeted, some endoderm cells were also electroporated (Figure 3C and Table 1).
One of the major advantages of using a capillary electrode is that the number of electroporated cells can be controlled, simply by changing the diameter of its opening. To determine the number of electroporated cells, embryos were fixed 2 hr after electroporation and imaged in wholemount on a confocal microscope. The number of GFP+ cells was manually counted in the confocal z-stacks. Table 1 shows that, for a given stage, increasing the opening size of the glass capillary from 20 to 30 &#181;m results in DNA uptake by more cells. When a single opening size was compared between stages (E7.5 versus E8.5), more cells were found to be electroporated at the latter stage. This effect may be due to a higher concentration of DNA present in the amniotic cavity at E8.5. Because the DNA solution was mixed with the green food dye, we can use the green color to assess the DNA concentration in the amniotic cavity. In the microscope, it is clear that, when compared with E8.5 embryos, the green color after DNA injection is much lighter in the cavity of E7.5 embryos. Although the same concentration of DNA solution was injected into E7.5 and E8.5 embryos, more DNA solution was in the amniotic cavity of E8.5 embryos in order to fill it completely because they are larger in the size. After withdrawing the injection needle, there is always some degree of leakage of DNA solution from the amniotic cavity, and since the puncture hole is larger in comparison to the size of the amniotic cavity in earlier embryos, it is likely that there was proportionately more leakage from E7.5 than E8.5 embryos, leading to a lower DNA concentration. The different number of transfected cells could also be due to the different diameters or induced transmembrane voltage (ITV) thresholds of cells at different stages.
A drawback of electroporation is the associated cell death. Similar to the traditional gold plated or needle-shaped electrodes, electroporation using a capillary electrode also causes the cell death. After electroporation the targeted region appeared darker in colour compared to neighbouring regions (Figure 3A and 3B), indicating that some degree of cell death must have occurred in this area. To further determine the number of dead cells caused by the electroporation procedure, embryos were stained with a fluorescent cell membrane-impermeable nuclear dye. The nuclei of dead cells were labelled with a membrane-impermeable far-red fluorescence dye. The staining confirmed that this capillary electroporation technique only results in a small number of dead cells near the electroporation site (Figure 3D and Table 1).
We noticed that although dead cells appear at the electroporation site, GFP+ cells and dead cells are also most exclusive from each other (Figure 3E and 3F). Moreover, when the caudal lateral epiblast at E8.5 was electroporated with pCAG-GFP and a glass capillary opening of 20&#181;m, a large number of GFP+ cells was detected after 48 hr in culture (Figure 3G and 3H). Taken together, these results suggest most GFP+ cells detected 2 hr after electroporation are still viable during the further culture.
We scored the number of GFP+ cells after 24 hr ex vivo culture. Six embryos were electroporated with pCAG-GFP at E7.5, using a capillary opening of 20&#181;m diameter. 107&#177;31 (mean &#177; s.d.) GFP+ cells/embryo were detected. Since at the start of culture (2 hr), 9 cells were electroporated on average per embryo (Table 1), this suggests that electroporated cells underwent 3-4 divisions within 2 hr. The average cell doubling time from E7.5 to E8.5 embryos is around 6-7 hr in all cells apart from those in the ventral node19,20. This suggests that the electroporation procedure does not hamper normal cell growth.
<img alt="Figure 1" src="/files/ftp_upload/53295/53295fig1.jpg" /> 
Figure 1. Circuit diagram showing the electroporation setup. The embryo containing DNA solution in its amniotic cavity was positioned between the two electrodes. Current at the chosen parameters was provided by a square wave pulse&#160;generator (power supply). A multimeter was connected in series to detect the electric current passing the embryo. <a href="https://www.jove.com/files/ftp_upload/53295/53295fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53295/53295fig2.jpg" /> 
Figure 2. The distribution of grafted or electroporated cells in host embryos. (A-H) GFP fluorescence overlays (green) on brightfield images of wholemount embryos (grayscale) (A) 10-16 GFP+ EpiSCs were grafted into the distal region of a late-streak stage embryo. (C) A larger clump of GFP+ EpiSCs was grafted into the distal region of a mid-streak stage embryo. (B and D) The distribution of EpiSCs derived cells (green) in the host embryos (shown in A and C) after 24 hr in culture. (B) GFP+ cells dispersed in the host embryo, suggesting correct integration of the donor cells. (D) Grafting larger cell clumps resulted in unincorporated clump formation in the host embryo. (E-K) pCAG-Cre:GFP plasmid electroporated into specific areas of wildtype embryos. Electroporating the distal region of an early bud stage embryo (E) or the primitive streak of a 2-5 somite stage embryo (G) resulted in GFP+ cells in these regions 2 hr after the procedure. (F and H) The distribution of GFP+ cells in the host embryos after 24 hr in culture, showing that electroporated cells contribute to the neuroectoderm (black arrow) (F) and paraxial mesoderm (white arrow) (H). (I-K) DAB immunostaining for the GFP+ cells showing that the electroporated cells can give rise to the neuroectoderm (I), mesoderm (J) and endoderm (J and K) after 24 hr in culture. Scale bar (A-H) = 250 &#181;m; scale bar (I-K) = 100 &#181;m. Note: Figure 1A and 1B are reprinted from our previous publication4. <a href="https://www.jove.com/files/ftp_upload/53295/53295fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53295/53295fig3.jpg" /> 
Figure 3. Distribution of GFP+ cells and dead cells in the embryos after electroporation. (A-C) pCAG-Cre:GFP plasmid electroporated into the caudal lateral epiblast cells of an E8.5 (2-5 somite stage) embryo (capillary opening size: 20 &#181;m). (A) 2 hr after the procedure, the targeted region showed a dark color (white arrow) compared to other parts of the embryo. Inset shows an enlargement of the electroporated region. (B) Brightfield image (grayscale) overlaid with the green fluorescent channel showing the electroporated cells (green). (C) A confocal z-slice showing that two endoderm cells (green) also took up the plasmid when the caudal lateral epiblast cells were targeted. The cell nuclei are shown in red. (D-F) pCAG-Cre:GFP plasmid was electroporated in the caudal aspect of the node of an E8.5 embryo (capillary opening size: 30 &#181;m). The embryo was cultured for 2 hr. Electroporated cells are shown in green and dead cells in red. (D) The electroporated area contains both GFP+ cells as well as dead cells. The area in the white box was further analyzed in a confocal microscope. Manual counting of the z-stack showed that there were 33 GFP+ cells and 23 dead cells in this area. Only two cells were both positive for both fluorophores. (E and F) XYZ view of a confocal z-slice from the white boxed region in D showing GFP+ cells are separate from the dead cells. The nuclei are shown in blue. (G and H) pCAG-Cre:GFP plasmid was electroporated into a few cells in the caudal lateral epiblast of an E8.5 embryo (capillary opening size: 20 &#181;m) and imaged after two (G) and 48 (H) hours ex vivo culture Note: (H) The embryo were severed in two after culture. The head and heart regions were removed. Scale bar (A, B, D, G and H) = 250 &#181;m; scale bar (C, E and F) = 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/53295/53295fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Diameter of opening of the capillary tube&#160;</td>
			<td>Embryo stage</td>
			<td>Electroporation efficiency: no. embryos containing GFP+ cells after 2h&#160; / total no. of electroporated embryos (no. GFP+ embryos that developed normally after 24 or 48h culture)</td>
			<td>Average number of GFP+ cells per embryo &#177;s.d. (n= no. of examined embryos)</td>
			<td>No. of GFP+ endodermal cells per each embryo &#177;s.d. (n= no. of examined embryos)</td>
		</tr>
		<tr>
			<td>20&#956;m</td>
			<td>E7.5 (LS-LB)</td>
			<td>7 / 9 (7)</td>
			<td>9&#177;3 (n=4)</td>
			<td>4&#177;2 (n=4)</td>
		</tr>
		<tr>
			<td>30&#956;m</td>
			<td>E7.5 (LS-LB)</td>
			<td>13 / 15 (12)</td>
			<td>17&#177;2(n=4)</td>
			<td>6&#177;1 (n=4)</td>
		</tr>
		<tr>
			<td>20&#956;m</td>
			<td>E8.5 (2-5 somites)</td>
			<td>12 / 13 (10)</td>
			<td>21&#177;4 (n=4)</td>
			<td>11&#177;4 (n=4)</td>
		</tr>
		<tr>
			<td>30&#956;m</td>
			<td>E8.5 (2-5 somites)</td>
			<td>2 / 2 (2)</td>
			<td>33 and 26 (n=2)</td>
			<td>14 and 16 (n=2)</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;Electroporation efficiency of pCAG-Cre:GFP plasmid in mouse embryos. 
Abbreviation: LS, late primitive streak stage; LB: late bud stage. Embryos are staged according to Downs and Davies12

Watch this Scientific Journal Video about Methods for Precisely Localized Transfer of Cells or DNA into Early Postimplantation Mouse Embryos at JoVE.com

Methods for Precisely Localized Transfer of Cells or DNA into Early Postimplantation Mouse Embryos

We demonstrate a method for grafting cultured cells into defined sites of early mouse embryos to determine their in vivo potential. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing the precise delivery of exogenous DNA into a few cells in the embryos.

Manipulation and culture of early mouse embryos is a powerful yet largely under-utilized technology enhancing the value of this model system. Conversely, ...

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Developmental Biology

10.3K vistas.  School of Biological Sciences, University of Edinburgh. We demonstrate a method for grafting cultured cells into defined sites of early mouse embryos to determine their in vivo potential. We also introduce an optimized electroporation method that uses glass capillaries of known diameter, allowing the precise delivery of exogenous DNA into a few cells in the embryos.

Video: Methods for Precisely Localized Transfer of Cells or DNA into Early Postimplantation Mouse Embryos

Loss- and Gain-of-function Approach to Investigate Early Cell Fate Determinants in Preimplantation Mouse Embryos

Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene modification in preimplantation embryos provides a means to study the underlying mechanisms of genes implicated in, for instance, cellular differentiation into the trophectoderm (TE) and the inner cell mass (ICM). Here, we describe a protocol that examines the role of neogenin as an authentic receptor for initial cell fate determination in preimplantation mouse embryos. First, we discuss the experimental manipulations that were used to produce gain and loss of neogenin function by microinjecting neogenin cDNA and shRNA; the effectiveness of this approach was confirmed by a strong correlation between the pair-wise expression levels of either red fluorescent protein (RFP) or green fluorescent protein (GFP) and the immunocytochemical quantification of neogenin expression. Secondly, overexpression of neogenin in preimplantation mouse embryos leads to normal ICM development while neogenin knockdown causes the ICM to develop abnormally, implying that neogenin could be a receptor that relays extracellular cues to drive blastomeres to early cell fates. Given the success of this detailed protocol in investigating the function of a novel embryonic developmental stage-specific receptor, we propose that it has the potential to aid in exploration and identification of other stage-specific genes during embryogenesis.

The goal of this protocol is to describe a loss- and gain-of function method that is applicable to identify neogenin as a stage-specific receptor that leads to trophectoderm and inner cell mass differentiation in preimplantation mouse embryos.

 Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene ...

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo development due to their similar developmental process. Although genetic factors are known to affect early embryonic development, the discovery of such factors has been a serious challenge. Microarray technology allows quantitative measurement and gene expression profiling of transcript levels on a genome-wide basis. One of the main decisions that have to be made when planning a microarray experiment is whether to use a one- or two-color approach. Two-color design increases technical replication, minimizes variability, improves sensitivity and accuracy as well as allows having loop designs, defining the common reference samples. Although microarray is a powerful biological tool, there are potential pitfalls that can attenuate its power. Hence, in this technical paper we demonstrate an optimized protocol for RNA extraction, amplification, labeling, hybridization of the labeled amplified RNA to the array, array scanning and data analysis using the two-color analysis strategy.

Transcriptome Profiling of In-Vivo Produced Bovine Pre-implantation Embryos Using Two-color Microarray Platform

Microarray technology allows quantitative measurement and gene expression profiling of transcripts on a genome-wide basis. Therefore, this protocol provides an optimized technical procedure in a two-color custom made bovine array using Day 7 bovine embryos to demonstrate the feasibility of using low amount of total RNA.

Early embryonic loss is a large contributor to infertility in cattle. Moreover, bovine becomes an interesting model to study human preimplantation embryo ...

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. The inner ear auditory sensory organ is critical for our ability to hear. It houses a highly specialized sensory epithelium that consists of mechano-transducing hair cells (HCs) and surrounding glial-like supporting cells (SCs). Despite structural differences in the auditory organs, molecular mechanisms regulating the development of the auditory organ are evolutionarily conserved between mammals and aves. In ovo electroporation is largely limited to early stages at E1 - E3. Due to the relative late development of the auditory organ at E5, manipulations of the auditory organ by in ovo electroporation past E3 are difficult due to the advanced development of the chicken embryo at later stages. The method presented here is a transient gene transfer method for targeting genes of interest at stage E4 - E4.5 in the developing chicken auditory sensory organ via in ovo micro-electroporation. This method is applicable for gain- and loss-of-functions with conventional plasmid DNA-based expression vectors and can be combined with in ovo cell proliferation assay by adding EdU (5-ethynyl-2&#180;-deoxyuridine) to the whole embryo at the time of electroporation. The use of green or red fluorescent protein (GFP or RFP) expression plasmids allows the experimenter to quickly determine whether the electroporation successfully targeted the auditory portion of the developing inner ear. In this method paper, representative examples of GFP electroporated specimens are illustrated; embryos were harvested 18 - 96 hr&#160;after electroporation and targeting of GFP to the pro-sensory area of the auditory organ was confirmed by RNA in situ hybridization. The method paper also provides an optimized protocol for the use of the thymidine analog EdU to analyze cell proliferation; an example of an EdU based cell proliferation assay that combines immuno-labeling and click EdU chemistry is provided.

Gene Transfer into the Chicken Auditory Organ by In Ovo Micro-electroporation

The auditory organ mediates hearing. Here we present a modified in ovo micro-electroporation method optimized for studying auditory progenitor cell proliferation and differentiation in the developing chicken auditory organ.

Chicken embryos are ideal model systems for studying embryonic development as manipulations of gene function can be conducted with relative ease in ovo. ...

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of morphogenetic events, such as gastrulation2, neurulation3-5, somitogenesis6, heart bending7,8, and brain formation9-13, during early embryogenesis. Key to understanding morphogenetic processes is to follow them dynamically by time-lapse imaging. The acquisition of time-lapse movies of chick embryogenesis ex ovo has been limited either to short time windows or to the need for an incubator to control temperature and humidity around the embryo14. Here, we present a new technique to culture chick embryos ex ovo for high-resolution time-lapse imaging using transmitted light microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique (EC-culture)1 and allows for the culturing of chick embryos without the need for a climate chamber. The embryo is sandwiched between two identical filter paper carriers and is kept fully submerged in a simple, temperature-controlled medium covered by a layer of light mineral oil. Starting from the primitive streak stage (Hamburger-Hamilton stage 5, HH5)15 up to at least the 28-somite stage (HH16)15, embryos can be cultured with either their ventral or dorsal side up. This allows the acquisition of time-lapse movies covering about 30 hr of embryonic development. Representative time-lapse frames and movies are shown. Embryos are compared morphologically to an embryo cultured in the standard EC-culture.
The submerged filter paper sandwich provides a stable environment to study early dorsal and ventral morphogenetic processes. It also allows for live fluorescence imaging and micromanipulations, such as microsurgery, bead implantation, microinjection, gene silencing, and electroporation, and has a strong potential to be combined with immersion objectives for laser-based imaging (including light-sheet microscopy).

A Submerged Filter Paper Sandwich for Long-term Ex Ovo Time-lapse Imaging of Early Chick Embryos

Here, we present a new technique to culture chick embryos ex ovo for time-lapse imaging using transmitted light and fluorescence microscopy. The submerged filter paper sandwich is a variant of the well-established filter paper carrier technique1 that allows for the culturing of chick embryos fully-submerged in a liquid culture medium.

Due to its availability, low cost, flat geometry, and transparency, the ex ovo chick embryo has become a major vertebrate animal model for the study of ...

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. The observation of Tribolium embryos with light sheet-based fluorescence microscopy has multiple advantages over conventional widefield and confocal fluorescence microscopy. Due to the unique properties of a light sheet-based microscope, three dimensional images of living specimens can be recorded with high signal-to-noise ratios and significantly reduced photo-bleaching as well as photo-toxicity along multiple directions over periods that last several days. With more than four years of methodological development and a continuous increase of data, the time seems appropriate to establish standard operating procedures for the usage of light sheet technology in the Tribolium community as well as in the insect community at large. This protocol describes three mounting techniques suitable for different purposes, presents two novel custom-made transgenic Tribolium lines appropriate for long-term live imaging, suggests five fluorescent dyes to label intracellular structures of fixed embryos and provides information on data post-processing for the timely evaluation of the recorded data. Representative results concentrate on long-term live imaging, optical sectioning and the observation of the same embryo along multiple directions. The respective datasets are provided as a downloadable resource. Finally, the protocol discusses quality controls for live imaging assays, current limitations and the applicability of the outlined procedures to other insect species.
This protocol is primarily intended for developmental biologists who seek imaging solutions that outperform standard laboratory equipment. It promotes the continuous attempt to close the gap between the technically orientated laboratories/communities, which develop and refine microscopy methodologically, and the life science laboratories/communities, which require 'plug-and-play' solutions to technical challenges. Furthermore, it supports an axiomatic approach that moves the biological questions into the center of attention.

Light Sheet-based Fluorescence Microscopy of Living or Fixed and Stained Tribolium castaneum Embryos

Imaging the morphogenesis of insect embryos with light sheet-based fluorescence microscopy has become state of the art. This protocol outlines and compares three mounting techniques appropriate for Tribolium castaneum embryos, introduces two novel custom-made transgenic lines well suited for live imaging, discusses essential quality controls and indicates current experimental limitations.

The red flour beetle Tribolium castaneum has become an important insect model organism in developmental genetics and evolutionary developmental biology. ...

A Novel Use of Three-dimensional High-frequency Ultrasonography for Early Pregnancy Characterization in the Mouse

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is routinely used as an in vivo model to study embryo implantation and pregnancy progression. Unfortunately, such murine studies require pregnancy interruption to enable follow-up phenotypic analysis. To address this issue, we used three-dimensional (3-D) reconstruction of HFUS imaging data for early detection and characterization of murine embryo implantation sites and their individual developmental progression in utero. Combining HFUS imaging with 3-D reconstruction and modeling, we were able to accurately quantify embryo implantation site number as well as monitor developmental progression in pregnant C57BL6J/129S mice from 5.5 days post coitus (d.p.c.) through to 9.5 d.p.c. with the use of a transducer. Measurements included: number, location, and volume of implantation sites as well as inter-implantation site spacing; embryo viability was assessed by cardiac activity monitoring. In the immediate post-implantation period (5.5 to 8.5 d.p.c.), 3-D reconstruction of the gravid uterus in both mesh and solid overlay format enabled visual representation of the developing pregnancies within each uterine horn. As genetically engineered mice continue to be used to characterize female reproductive phenotypes derived from uterine dysfunction, this method offers a new approach to detect, quantify, and characterize early implantation events in vivo. This novel use of 3-D HFUS imaging demonstrates the ability to successfully detect, visualize, and characterize embryo-implantation sites during early murine pregnancy in a non-invasive manner. The technology offers a significant improvement over current methods, which rely on the interruption of pregnancies for gross tissue and histopathologic characterization. Here we use a video and text format to describe how to successfully perform ultrasounds of early murine pregnancy to generate reliable and reproducible data with reconstruction of the uterine form in mesh and solid 3-D images.

Mice are widely used to study gestational biology. However, pregnancy termination is required for such studies which precludes longitudinal investigations and necessitates the use of large numbers of animals. Therefore, we describe a non-invasive technique of high-frequency ultrasonography for early detection and monitoring of post-implantation events in the pregnant mouse.

High-frequency ultrasonography (HFUS) is a common method to non-invasively monitor the real-time development of the human fetus in utero. The mouse is ...

Visualization of Cellular Electrical Activity in Zebrafish Early Embryos and Tumors

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling processes of excitable neuronal and muscular cells and many other biological processes, such as embryonic developmental patterning. However, there is a need for in vivo electrical activity monitoring in vertebrate embryogenesis. The advances of genetically encoded fluorescent voltage indicators (GEVIs) have made it possible to provide a solution for this challenge. Here, we describe how to create a transgenic voltage indicator zebrafish using the established voltage indicator, ASAP1 (Accelerated Sensor of Action Potentials 1), as an example. The Tol2 kit and a ubiquitous zebrafish promoter, ubi, were chosen in this study. We also explain&#160;the processes of Gateway site-specific cloning, Tol2 transposon-based zebrafish transgenesis, and the imaging process for early-stage fish embryos and fish tumors using regular epifluorescent microscopes. Using this fish line, we found that there are cellular electric voltage changes during zebrafish embryogenesis, and fish larval movement. Furthermore, it was observed that in a few zebrafish malignant peripheral nerve sheath tumors, the tumor cells were generally polarized compared to the surrounding normal tissues.

Here, we show the process of creating a cellular electric voltage reporter zebrafish line to visualize embryonic development, movement, and fish tumor cells in vivo.

Bioelectricity, endogenous electrical signaling mediated by ion channels and pumps located on the cell membrane, plays important roles in signaling ...

Efficient Differentiation of Human Pluripotent Stem Cells into Liver Cells

The liver detoxifies harmful substances, secretes vital proteins, and executes key metabolic activities, thus sustaining life. Consequently, liver failure&#8212;which can be caused by chronic alcohol intake, hepatitis, acute poisoning, or other insults&#8212;is a severe condition that can culminate in bleeding, jaundice, coma, and eventually death. However, approaches to treat liver failure, as well as studies of liver function and disease, have been stymied in part by the lack of a plentiful supply of human liver cells. To this end, this protocol details the efficient differentiation of human pluripotent stem cells (hPSCs) into hepatocyte-like cells, guided by a developmental roadmap that describes how liver fate is specified across six consecutive differentiation steps. By manipulating developmental signaling pathways to promote liver differentiation and to explicitly suppress the formation of unwanted cell fates, this method efficiently generates populations of human liver bud progenitors and hepatocyte-like cells by days 6 and 18 of PSC differentiation, respectively. This is achieved through the temporally-precise control of developmental signaling pathways, exerted by small molecules and growth factors in a serum-free culture medium. Differentiation in this system occurs in monolayers and yields hepatocyte-like cells that express characteristic hepatocyte enzymes and have the ability to engraft a mouse model of chronic liver failure. The ability to efficiently generate large numbers of human liver cells in vitro has ramifications for treatment of liver failure, for drug screening, and for mechanistic studies of liver disease.

This protocol details a monolayer, serum-free method to efficiently generate hepatocyte-like cells from human pluripotent stem cells (hPSCs) in 18 days. This entails six steps as hPSCs sequentially differentiate into intermediate cell-types such as the primitive streak, definitive endoderm, posterior foregut and liver bud progenitors before forming hepatocyte-like cells.

The liver detoxifies harmful substances, secretes vital proteins, and executes key metabolic activities, thus sustaining life. Consequently, liver ...

In Vitro Generation of Somite Derivatives from Human Induced Pluripotent Stem Cells

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give rise to multiple cell types, including the myotome (MYO), sclerotome (SCL), dermatome (D), and syndetome (SYN), which in turn develop into skeletal muscle, axial skeleton, dorsal dermis, and axial tendon/ligament, respectively. Therefore, the generation of SMs and their derivatives from human induced pluripotent stem cells (iPSCs) is critical to obtain pluripotent stem cells (PSCs) for application in regenerative medicine and for disease research in the field of orthopedic surgery. Although the induction protocols for MYO and SCL from PSCs have been previously reported by several researchers, no study has yet demonstrated the induction of SYN and D from iPSCs. Therefore, efficient induction of fully competent SMs remains a major challenge. Here, we recapitulate human SM patterning with human iPSCs in vitro by mimicking the signaling environment during chick/mouse SM development, and report on methods of systematic induction of SM derivatives (MYO, SCL, D, and SYN) from human iPSCs under chemically defined conditions through the presomitic mesoderm (PSM) and SM states. Knowledge regarding chick/mouse&#160;SM development was successfully applied to the induction of SMs with human iPSCs. This method could be a novel tool for studying human somitogenesis and patterning without the use of embryos and for cell-based therapy and disease modeling.

We present here a protocol for the differentiation of human induced pluripotent stem cells into each somite derivative (myotome, sclerotome, dermatome, and syndetome) in chemically defined conditions, which has applications in future disease modeling and cell-based therapies in orthopedic surgery.

In response to signals such as WNTs, bone morphogenetic proteins (BMPs), and sonic hedgehog (SHH) secreted from surrounding tissues, somites (SMs) give ...

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the development of neuronal connectivity. During establishment of the retino-tectal projection, optic axons extend from the eye and navigate through distinct regions of the brain to reach their target tissue, the optic tectum. Once optic axons enter the tectum, they elaborate terminal arbors that function to increase the number of synaptic connections they can make with target interneurons in the tectum. Here, we describe a method to express DNA encoding green fluorescent protein (GFP), and gain- and loss-of-function gene constructs, in optic neurons (retinal ganglion cells) in Xenopus embryos. We explain how to microinject a combined DNA/lipofection reagent into eyebuds of one day old embryos such that exogenous genes are expressed in single or small numbers of optic neurons. By tagging genes with GFP or co-injecting with a GFP plasmid, terminal axonal arbors of individual optic neurons with altered molecular signaling can be imaged directly in brains of intact, living Xenopus tadpoles several days later, and their morphology can be quantified. This protocol allows for determination of cell-autonomous molecular mechanisms that underlie the development of optic axon arborization in vivo.

Microinjection of DNA into Eyebuds in Xenopus laevis Embryos and Imaging of GFP Expressing Optic Axonal Arbors in Intact, Living Xenopus Tadpoles

This protocol aims to demonstrate how to microinject a DNA/DOTAP mixture into eyebuds of one day old Xenopus laevis embryos, and how to image and reconstruct individual green fluorescent protein (GFP) expressing optic axonal arbors in tectal midbrains of intact, living Xenopus tadpoles.

The primary visual projection of tadpoles of the aquatic frog Xenopus laevis serves as an excellent model system for studying mechanisms that regulate the ...