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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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><tbody><tr><td>Forceps</td><td>Dumostar</td><td>T5390</td><td></td></tr><tr><td>Dissecting stereomicroscope&amp;nbsp;</td><td>&amp;nbsp;Zeiss</td><td>Stemi 2000-C</td><td></td></tr><tr><td>Stereomicroscope system with fluorescence&amp;nbsp;</td><td>Nikon</td><td>AZ100</td><td></td></tr><tr><td>Inverted microscope with a digital camera</td><td>Olympus</td><td>Olympus BX61</td><td></td></tr><tr><td>Inverted confocal microscope&amp;nbsp;</td><td>Leica Microsystems</td><td>Leica TCS SP8</td><td></td></tr><tr><td>Low melting point agarose&amp;nbsp;</td><td>Life Technologies</td><td>16520-050</td><td></td></tr><tr><td>Pasteur pipettes</td><td>Fisher Scientific</td><td>11397863</td><td></td></tr><tr><td>30mm Petri dishes&amp;nbsp;</td><td>Fisher Scientific</td><td>121V</td><td></td></tr><tr><td>4-well plates</td><td>Thermo scientific</td><td>179820&amp;nbsp;</td><td></td></tr><tr><td>M2 medium&amp;nbsp;</td><td>Sigma-Aldrich</td><td>M7167</td><td></td></tr><tr><td>Phosphate Buffered Saline (PBS)</td><td>Life Technologies</td><td>10010015&amp;nbsp;</td><td></td></tr><tr><td>Paraformaldehyde (PFA)&amp;nbsp;</td><td>Sigma-Aldrich</td><td>P6148</td><td></td></tr><tr><td>Pipettes&amp;nbsp;</td><td>NICHIRYO</td><td>Nichipet</td><td></td></tr><tr><td>tips&amp;nbsp;</td><td>Greiner Bio One</td><td>685280</td><td></td></tr><tr><td>Cell culture incubator&amp;nbsp;</td><td>SANYO</td><td>MCO-17AIC</td><td></td></tr><tr><td>Roller culture apparatus&amp;nbsp;</td><td>BTC Engineering</td><td></td><td></td></tr><tr><td>Syringe filters 0.45&amp;micro;m, sterile&amp;nbsp;</td><td>Sigma-Aldrich</td><td>10462100</td><td></td></tr><tr><td>Glasgow Minimum Essential Medium (GMEM)</td><td>Sigma-Aldrich</td><td>G5154</td><td></td></tr><tr><td>non-essential amino acids (NEAA)</td><td>Life Technologies</td><td>11140050</td><td></td></tr><tr><td>L-glutamine</td><td>Fisher Scientific</td><td>SH30549.01</td><td></td></tr><tr><td>Sodium pyruvate solution</td><td>Fisher Scientific</td><td>SH30239.01</td><td></td></tr><tr><td>Penicillin and Streptomycin 10.000UI/ml</td><td>Lonza</td><td>DE17-602E</td><td></td></tr><tr><td>Gas Cartridge for Portable Meker Burner</td><td>COLEMAN&amp;nbsp;</td><td>COLEMAN 250</td><td></td></tr><tr><td>Thin Wall Borosilicate Capillary Glass with Fillament, OD 1.0 mm, ID 0.78 mm</td><td>Harvard Apparatus</td><td>640798</td><td></td></tr><tr><td>Aspirator tube assemblies for calibrated microcapillary pipettes&amp;nbsp;</td><td>Sigma-Aldrich</td><td>A5177-5EA&amp;nbsp;</td><td></td></tr><tr><td>Flaming/Brown micropipette puller&amp;nbsp;</td><td>Sutter Instrument Company</td><td>P-97&amp;nbsp;</td><td></td></tr><tr><td>Microforge&amp;nbsp;</td><td>De Fonbrune</td><td>BS030301</td><td></td></tr><tr><td>Pneumatic pico pump&amp;nbsp;</td><td>World Precision Instruments</td><td>PV830</td><td></td></tr><tr><td>Microloader tips&amp;nbsp;</td><td>Eppendorf</td><td>5242956.003</td><td></td></tr><tr><td>ECM 830 square wave pulse generator&amp;nbsp;</td><td>BTX</td><td>45-0002</td><td></td></tr><tr><td>Green food coloring dye</td><td>Sigma-Aldrich</td><td>C.I. 42053</td><td></td></tr><tr><td>A far-red cell membrane-impermeable nuclear dye</td><td>Biotium</td><td>40060-T</td><td></td></tr><tr><td>pCAG-Cre:GFP&amp;nbsp;</td><td>Addgene</td><td>#13776&amp;nbsp;</td><td></td></tr><tr><td>pCAG-GFP&amp;nbsp;</td><td>Addgene</td><td>#16664</td><td></td></tr><tr><td>Multimeter</td><td>Excel</td><td>XL830L</td><td></td></tr><tr><td>Micromanipulators&amp;nbsp;</td><td>Leitz&amp;nbsp;</td><td></td><td></td></tr><tr><td>0.2mm diameter platinum wire&amp;nbsp;</td><td>Agar Scientific</td><td>E404-2</td><td></td></tr><tr><td>Anti-GFP antibody</td><td>Abcam</td><td>ab13970</td><td></td></tr><tr><td>Goat anti-Chicken IgY, HRP</td><td>Santa Cruz</td><td>sc-2428</td><td></td></tr><tr><td>Liquid DAB+ Substrate Chromogen System&amp;nbsp;</td><td>Dako</td><td>K3467</td><td></td></tr><tr><td>4&#039;,6-diamidino-2-phenylindole (DAPI)</td><td>Life Technologies</td><td>D21490</td><td></td></tr><tr><td>A far-red fluorescence nuclear counterstain</td><td>Life Technologies</td><td>T3605</td><td></td></tr></tbody></table>

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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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.remotexs.ntu.edu.sg/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, ...

methods-for-precisely-localized-transfer-cells-or-dna-into-early

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

10.4K Views.  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

Quantitative Analysis of Protein Expression to Study Lineage Specification in Mouse Preimplantation Embryos

This protocol presents a method to perform quantitative, single-cell in situ analyses of protein expression to study lineage specification in mouse preimplantation embryos. The procedures necessary for embryo collection, immunofluorescence, imaging on a confocal microscope, and image segmentation and analysis are described. This method allows quantitation of the expression of multiple nuclear markers and the spatial (XYZ) coordinates of all cells in the embryo. It takes advantage of MINS, an image segmentation software tool specifically developed for the analysis of confocal images of preimplantation embryos and embryonic stem cell (ESC) colonies. MINS carries out unsupervised nuclear segmentation across the X, Y and Z dimensions, and produces information on cell position in three-dimensional space, as well as nuclear fluorescence levels for all channels with minimal user input. While this protocol has been optimized for the analysis of images of preimplantation stage mouse embryos, it can easily be adapted to the analysis of any other samples exhibiting a good signal-to-noise ratio and where high nuclear density poses a hurdle to image segmentation (e.g., expression analysis of embryonic stem cell (ESC) colonies, differentiating cells in culture, embryos of other species or stages, etc.).

This protocol presents a method to perform quantitative, single-cell in situ analysis of protein expression to study lineage specification in mouse preimplantation embryos. The procedures necessary for collection of blastocysts, whole-mount immunofluorescent detection of proteins, imaging of samples on a confocal microscope, and nuclear segmentation and image analysis are described.

This protocol presents a method to perform quantitative, single-cell in situ analyses of protein expression to study lineage specification in mouse ...

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 ...

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance molecules, local gene transfer or knock- out is required. Gene targeting knock-in or knock-out into local regions is possible to perform with combination with a specific CRE line, which is laborious, costly, and time consuming. Therefore, a simple transfection method, an in utero electroporation technique, which can be performed with short time, will be handy to test the possible function of candidate genes prior to the generation of transgenic animals 1,2. In addition to this, in utero electroporation targets areas of the brain where no specific CRE line exists, and will limit embryonic lethality 3,4. Here, we present a method of in utero electroporation combining two different types of electrodes for simple and convenient gene transfer into target areas of the developing brain. First, a unique holding method of embryos using an optic fiber optic light cable will make small embryos (from E9.5) visible for targeted DNA solution injection into ventricles and needle type electrodes insertion to the targeted brain area 5,6. The patterning of the brain such as cortical area occur at early embryonic stage, therefore, these early electroporation from E9.5 make a big contribution to understand entire area patterning event. Second, the precise shape of a capillary prevents uterine damage by making holes by insertion of the capillary. Furthermore, the precise shape of the needle electrodes are created with tungsten and platinum wire and sharpened using sand paper and insulated with nail polish 7, a method which is described in great detail in this protocol. This unique technique allows transfection of plasmid DNA into restricted areas of the brain and will enable small embryos to be electroporated. This will help to, open a new window for many scientists who are working on cell differentiation, cell migration, axon guidance in very early embryonic stage. Moreover, this technique will allow scientists to transfect plasmid DNA into deep parts of the developing brain such as thalamus and hypothalamus, where not many region-specific CRE lines exist for gain of function (GOF) or loss of function (LOF) analyses.

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

A gene transfer method into the developing mouse brain is described by using a unique surgical method and special shape of electrodes. This unique technique allows transfection of plasmid DNA temporally and spatially, which will aid many neuroscientists in studying brain development.

In order to understand the function of genes expressed in specific region of the developing brain, including signaling molecules and axon guidance ...

Neuroscience

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. The CRISPR/Cas9 system allows researchers to modify the genome with unprecedented efficiency, fidelity, and simplicity. Harnessing this technology, researchers are seeking a rapid, efficient, and easy protocol for generating GM mice. Here we introduce an improved method for cryopreservation of one-cell embryos that leads to a higher developmental rate of the freeze-thawed embryos. By combining it with optimized electroporation conditions, this protocol allows for the generation of knockout and knock-in mice with high efficiency and low mosaic rates within a short time. Furthermore, we show a step-by-step explanation of our optimized protocol, covering CRISPR reagent preparation, in vitro fertilization, cryopreservation and thawing of one-cell embryos, electroporation of CRISPR reagents, mouse generation, and genotyping of the founders. Using this protocol, researchers should be able to prepare GM mice with unparalleled ease, speed, and efficiency.

Here, we present a modified method for cryopreservation of one-cell embryos as well as a protocol that couples the use of freeze-thawed embryos and electroporation for the efficient generation of genetically modified mice.

The use of genetically modified (GM) mice has become crucial for understanding gene function and deciphering the underlying mechanisms of human diseases. ...

Genetics

Spatiotemporal Subcellular Manipulation of the Microtubule Cytoskeleton in the Living Preimplantation Mouse Embryo using Photostatins

The microtubule cytoskeleton forms the framework of a cell and is fundamental for intracellular transport, cell division, and signal transduction. Traditional pharmacological disruption of the ubiquitous microtubule network using, for instance, nocodazole can have devastating consequences for any cell. Reversibly photoswitchable microtubule inhibitors have the potential to overcome the limitations by enabling drug effects to be implemented in a spatiotemporally-controlled manner. One such family of drugs is the azobenzene-based photostatins (PSTs). These compounds are inactive in dark conditions, and upon illumination with UV light, they bind to the colchicine-binding site of &#946;-tubulin and block microtubule polymerization and dynamic turnover. Here, the application of PSTs in the 3-dimensional (3D) live preimplantation mouse embryo is set out to disrupt the microtubule network on a subcellular level. This protocol provides instructions for the experimental setup, as well as light activation and deactivation parameters for PSTs using live-cell confocal microscopy. This ensures reproducibility and enables others to apply this procedure to their research questions. Innovative photoswitches like PSTs may evolve as powerful tools to advance the understanding of the dynamic intracellular microtubule network and to non-invasively manipulate the cytoskeleton in real-time. Furthermore, PSTs may prove useful in other 3D structures such as organoids, blastoids, or embryos of other species.

Typical microtubule inhibitors, used widely in basic and applied research, have far-reaching effects on cells. Recently, photostatins emerged as a class of photoswitchable microtubule inhibitors, capable of instantaneous, reversible, spatiotemporally precise manipulation of microtubules. This step-by-step protocol details the application of photostatins in a 3D live preimplantation mouse embryo.

The microtubule cytoskeleton forms the framework of a cell and is fundamental for intracellular transport, cell division, and signal transduction. ...

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the efficiency in developing gene-knockout mice by inducing small indel mutation would be sufficient enough, the efficiency of embryo genome editing for making large-size DNA knock-in (KI) is still low. Therefore, in contrast to the direct KI method in embryos, gene targeting using embryonic stem cells (ESCs) followed by embryo injection to develop chimera mice still has several advantages (e.g., high throughput targeting in vitro, multi-allele manipulation, and Cre and flox gene manipulation can be carried out in a short period). In addition, strains with difficult-to-handle embryos in vitro, such as BALB/c, can also be used for ESC targeting. This protocol describes the optimized method for large-size DNA (several kb) KI in ESCs by applying CRISPR/Cas9-mediated genome editing followed by chimera mice production to develop gene-manipulated mouse models.

Here we present a protocol for developing genetically modified mouse models using embryonic stem cells, especially for large DNA knock-in (KI). This protocol is tuned up using CRISPR/Cas9 genome editing, resulting in significantly improved KI efficiency compared with the conventional homologous recombination-mediated linearized DNA targeting method.

The CRISPR/Cas9 system has made it possible to develop genetically modified mice by direct genome editing using fertilized zygotes. However, although the ...

Chromatin Immunoprecipitation Assay for Tissue-specific Genes using Early-stage Mouse Embryos

Chromatin immunoprecipitation (ChIP) is a powerful tool to identify protein:chromatin interactions that occur in the context of living cells 1-3. This technique has been widely exploited in tissue culture cells, and to a lesser extent, in primary tissue. The application of ChIP to rodent embryonic tissue, especially at early times of development, is complicated by the limited amount of tissue and the heterogeneity of cell and tissue types in the embryo. Here we present a method to perform ChIP using a dissociated embryonic day 8.5 (E8.5) embryo. Sheared chromatin from a single E8.5 embryo can be divided into up to five aliquots, which allows the investigator sufficient material for controls and for investigation of specific protein:chromatin interactions.
We have utilized this technique to begin to document protein:chromatin interactions during the specification of tissue-specific gene expression programs. The heterogeneity of cell types in an embryo necessarily restricts the application of this technique because the result is the detection of protein:chromatin interactions without distinguishing whether the interactions occur in all, a subset of, or a single cell type(s). However, examination of tissue-specific genes during or following the onset of tissue-specific gene expression is feasible for two reasons. First, immunoprecipitation of tissue specific factors necessarily isolates chromatin from the cell type where the factor is expressed. Second, immunoprecipitation of coactivators and histones containing post-translational modifications that are associated with gene activation should only be found at genes and gene regulatory sequences in the cell type where the gene is being or has been activated. The technique should be applicable to the study of most tissue-specific gene activation events.
In the example described below, we utilized E8.5 and E9.5 mouse embryos to examine factor binding at a skeletal muscle specific gene promoter. Somites, which are the precursor tissues from which the skeletal muscles of the trunk and limbs will form, are present at E8.5-9.54,5. Myogenin is a regulatory factor required for skeletal muscle differentiation 6-9. The data demonstrate that myogenin is associated with its own promoter in E8.5 and E9.5 embryos. Because myogenin is only expressed in somites at this stage of development 6,10, the data indicate that myogenin interactions with its own promoter have already occurred in skeletal muscle precursor cells in E8.5 embryos.

We demonstrate a chromatin immunoprecipitation (ChIP) method to identify factor interactions at tissue-specific genes during or after the onset of tissue-specific gene expression in mouse embryonic tissue. This protocol should be widely applicable for the study of tissue-specific gene activation as it occurs during normal embryonic development.

Chromatin immunoprecipitation (ChIP) is a powerful tool to identify protein:chromatin interactions that occur in the context of living cells 1-3. This ...

Biology

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect their in vivo potential. Preferably, these cells should be placed in a proper embryonic environment in order to acquire their definite phenotype. Furthermore, cell lineage tracing studies in the mouse after labeling cells with dyes or retroviral vectors has remained mostly limited to early stage mouse embryos with still poorly developed organs. To overcome these limitations, we designed standard and ultrasound-mediated microinjection protocols to inject various agents in targeted regions of the heart in mouse embryos at E9.5 and later stages of development. &#160;Embryonic explant or embryos are then cultured or left to further develop in utero. These agents include fluorescent dyes, virus, shRNAs, or stem cell-derived progenitor cells. Our approaches allow for preservation of the function of the organ while monitoring migration and fate of labeled and/or injected cells. These technologies can be extended to other organs and will be very helpful to address key biological questions in biology of development.

We describe a series of methods to inject dyes, DNA vectors, virus, and cells in order to monitor both cell fate and phenotype of endogenous and grafted cells derived from embryonic or pluripotent cells within mouse embryos at embryonic day (E)9.5 and later stages of development.

Testing the fate of embryonic or pluripotent stem cell-derivatives in in vitro protocols has led to controversial outcomes that do not necessarily reflect ...

Laser-assisted Lentiviral Gene Delivery to Mouse Fertilized Eggs

Lentiviruses are efficient vectors for gene delivery to mammalian cells. Following transduction, the lentiviral genome is stably incorporated into the host chromosome and is passed on to progeny. Thus, they are ideal vectors for creation of stable cell lines, in vivo delivery of indicators, and transduction of single cell fertilized eggs to create transgenic animals. However, mouse fertilized eggs and early stage embryos are protected by the zona pellucida, a glycoprotein matrix that forms a barrier against lentiviral gene delivery. Lentiviruses are too large to penetrate the zona and are typically delivered by microinjection of viral particles into the perivitelline cavity, the space between the zona and the embryonic cells. The requirement for highly skilled technologists and specialized equipment has minimized the use of lentiviruses for gene delivery to mouse embryos. This article describes a protocol for permeabilizing the mouse fertilized eggs by perforating the zona with a laser. Laser-perforation does not result in any damage to embryos and allows lentiviruses to gain access to embryonic cells for gene delivery. Transduced embryos can develop into blastocyst in vitro, and if implanted in pseudopregnant mice, develop into transgenic pups. The laser used in this protocol is effective and easy to use. Genes delivered by lentiviruses stably incorporate into mouse embryonic cells and are germline transmittable. This is an alternative method for creation of transgenic mice that requires no micromanipulation and microinjection of fertilized eggs.

Mouse fertilized eggs and early stage embryos are protected by the zona pellucida, a glycoprotein matrix that forms a barrier against gene delivery. This article describes a protocol for perforating the zona with a laser to transduce embryonic cells with lentiviral vectors and to create transgenic mice.

Lentiviruses are efficient vectors for gene delivery to mammalian cells. Following transduction, the lentiviral genome is stably incorporated into the ...

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

Summary

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Chapters in this video

Quantitative Analysis of Protein Expression to Study Lineage Specification in Mouse Preimplantation Embryos

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

Mouse in Utero Electroporation: Controlled Spatiotemporal Gene Transfection

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

Use of Freeze-thawed Embryos for High-efficiency Production of Genetically Modified Mice

Spatiotemporal Subcellular Manipulation of the Microtubule Cytoskeleton in the Living Preimplantation Mouse Embryo using Photostatins

CRISPR/Cas9-Mediated Highly Efficient Gene Targeting in Embryonic Stem Cells for Developing Gene-Manipulated Mouse Models

Chromatin Immunoprecipitation Assay for Tissue-specific Genes using Early-stage Mouse Embryos

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

Laser-assisted Lentiviral Gene Delivery to Mouse Fertilized Eggs