The authors would like to acknowledge the GWU Harlan Fellowship for support of Paaqua Grant and the GWU Luther Rice Fellowship for support of Mona Herold. This work was supported by the National Science Foundation grant MCB-1121711.

1. Preparation of Instruments, Culture Media and Dishes
<ol>
 <li>Sharpen four forceps using Alumina abrasive film. Do this under a dissecting microscope to monitor the tip size. One pair of forceps serves as a back-up in case a tip is damaged during the procedure. Autoclave all four forceps and store in a sterile container.</li>
 <li>Make 500 ml each of 0.1X and 1.0X culture medium (either Marc's Modified Ringers [MMR] or Modified Barth's Saline [MBS]; recipes in 15) and filter sterilize. We routinely use MBS (1X = 88 mM NaCl; 1 mM KCl, 0.7 mM CaCl2, 1 mM MgSO4, 5 mM HEPES [pH 7.8], 2.5 mM NaHCO3). Store at 14-18 &deg;C for months.</li>
 <li>Make a 2% agarose solution in culture medium (2 g electrophoresis grade agarose in 100 ml 1X culture medium in a screw cap glass bottle). Autoclave to dissolve agarose. This can be stored at 4 &deg;C for months, and microwaved to liquefy the agarose when it is next needed.</li>
 <li>While the agarose is liquid, pour about 0.5 ml onto the bottom of each well of a sterile 24-well culture plate. This will prevent the explants from sticking to the plastic.</li>
 <li>While the agarose is liquid, pour about 2-3 ml onto the bottom of two to three 60 mm Petri dishes, and swirl gently to make sure the bottom is covered. These will serve as dissection dishes and the agarose prevents embryos from sticking to the plastic once their membranes are removed.</li>
 <li>When the agarose has cooled and hardened, flame the tip of a 6" Pasteur pipette until it melts into a ball, and lightly touch it on the surface of the agarose in each well of the culture plate. This will create a shallow depression into which the explant will be placed.</li>
 <li>Fill each well of the culture plate with 1X sterile culture medium.</li>
 <li>When the agarose has cooled and hardened, fill the dissection dish with diluted culture medium (0.1X MMR or 0.1X MBS) to facilitate blastomere separation.</li>
</ol>
2. Selection of Embryos
<ol>
 <li>Obtain fertilized eggs and remove the jelly coats according to standard protocols 15. Transfer them to 0.5X culture medium in a 100 mm Petri dish.</li>
 <li>When embryos reach the 2-cell stage, sort those in which the first cleavage furrow bisects the lightly pigmented area in the animal hemisphere (Figure 1) into a separate dish. These will be the donors for the explants. Keep the remaining 2-cell embryos in a separate dish next to the donor embryos throughout the procedure to serve as sibling controls to stage the explants.</li>
</ol>
3. Preparation of Explants
<ol>
 <li>Place 5-10 embryos in a dissection dish, but only work on one embryo at a time.</li>
 <li>Position the first embryo so the transparent vitelline membrane can be seen; it is separated by a clear space (perivitelline space) above the surface of the animal pole of the embryo. Using a sharpened forceps in one's subdominant hand (left hand if one is right handed), grasp the vitelline membrane above the perivitelline space. Using a sharpened forceps in the other hand, grasp the membrane close to the first forceps tip, and gently pull in opposite directions to peel the membrane away. Make the initial grasp at a distance from the cell that is to be dissected, so that the target cell is not damaged during removal of the membrane. One can tell that the membrane has been removed because the embryo will flatten.</li>
 <li>Grab one neighboring cell with the forceps in the subdominant hand and use this cell as a "handle" so the cell to be dissected is not directly touched. With the forceps in the other hand, gently pull the remaining neighboring cells away from the desired blastomere. If midline cells, which share the same fate, are targeted, they can be removed together as a pair. Finally, dissect away the "handle" cell.</li>
 <li>Pick up the blastomere (or blastomere pair), with a sterile, glass Pasteur pipette, avoiding air bubbles and excessive suction. The pipette can be fire-polished to remove sharp edges that can damage the cell, but we do not routinely do this. Place the tip of the Pasteur pipette under the surface of the culture medium in a well of the explant culture dish, and gently expel the blastomere. It should slide into the shallow depression by gravity. The same blastomere from more than one embryo can be combined into a single explant.</li>
 <li>Repeat this procedure with the remaining embryos in the dissection dish, one-by-one. After about 10 embryos, the dissection dish will be filled with cellular debris. When this occurs, change to a fresh dissection dish.</li>
 <li>After all dissections are done, check whether the explants are in the shallow depressions. If not, they can be gently pushed into the depression with a hair loop that has been sterilized in 70% ethanol and air dried.</li>
 <li>About an hour after the last dissection, remove debris surrounding the healed explants with a sterile, glass Pasteur pipette.</li>
 <li>Culture the plate of explants at 14-20 &deg;C next to the Petri dish containing the sibling, control embryos. Sibling embryos will indicate the stage of development of the blastomere explants.</li>
</ol>
4. Harvesting Explants for Analysis
<ol>
 <li>Harvest the explants when the siblings reach the desired developmental stage. These explants survive quite well even if some of the cells disintegrate. Therefore, if there is a cloudy mass in the culture well, explore it with a hair loop or forceps to determine if a healthy explant is buried within.</li>
 <li>Pick up explants in a small volume of culture solution with a glass Pasteur pipette, and gently expel them into a fixative or lysis buffer appropriate for the assay to be conducted.</li>
</ol>

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	<li>Dale, L., Slack, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fate+map+of+the+32-cell+stage+of+Xenopus+laevis.">Fate map of the 32-cell stage of Xenopus laevis.</a> Development. 100, 279-295 (1987).</li><li>Masho, R., Kubota, H. Y. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+fates+of+blastomeres+of+the+eight-cell+stage+Xenopus+embryo.">Developmental fates of blastomeres of the eight-cell stage Xenopus embryo.</a> Dev. Growth, Diff. 30, 347-359 (1988).</li><li>Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fates+of+the+blastomeres+of+the+16-cell+stage+Xenopus+embryo.">Fates of the blastomeres of the 16-cell stage Xenopus embryo.</a> Dev. Biol. 119, 560-578 (1987).</li><li>Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fates+of+the+blastomeres+of+the+32-cell+stage+Xenopus+embryo.">Fates of the blastomeres of the 32-cell stage Xenopus embryo.</a> Dev. Biol. 122, 300-319 (1987).</li><li>Moody, S. A., Kline, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Segregation+of+fate+during+cleavage+of+frog+(Xenopus+laevis)+blastomeres.">Segregation of fate during cleavage of frog (Xenopus laevis) blastomeres.</a> Anat. Embryol. 182, 347-362 (1990).</li><li>Takasaki, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fates+and+roles+of+the+presumptive+organizer+region+in+the+32-cell+embryos+in+normal+development+of+Xenopus+laevis.">Fates and roles of the presumptive organizer region in the 32-cell embryos in normal development of Xenopus laevis.</a> Dev. Growth, Diff. 29, 141-152 (1987).</li><li>Sullivan, S. A., Moore, K. B., Moody, S. A., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+20+Early+events+in+frog+blastomere+fate+determination.">Chapter 20 Early events in frog blastomere fate determination.</a> Cell Fate and Lineage Determination. , 297-321 (1999).</li><li>Moore, K. B., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Animal-vegetal+asymmetries+influence+the+earliest+steps+in+retinal+fate+commitment+in+Xenopus.">Animal-vegetal asymmetries influence the earliest steps in retinal fate commitment in Xenopus.</a> Dev. Biol. 212, 25-41 (1999).</li><li>Yan, B., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+competence+of+Xenopus+blastomeres+to+produce+neural+and+retinal+progeny+is+repressed+by+two+endo-mesoderm+promoting+pathways.">The competence of Xenopus blastomeres to produce neural and retinal progeny is repressed by two endo-mesoderm promoting pathways.</a> Dev. Biol. 305, 103-119 (2007).</li><li>Gallagher, B. C., Hainski, A. M., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Autonomous+differentiation+of+dorsal+axial+structures+from+an+animal+cap+cleavage+stage+blastomere+in+Xenopus.">Autonomous differentiation of dorsal axial structures from an animal cap cleavage stage blastomere in Xenopus.</a> Development. 112, 1103-1114 (1991).</li><li>Hainski, A. M., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Xenopus+maternal+mRNAs+from+a+dorsal+animal+blastomere+induce+a+secondary+axis+in+host+embryos.">Xenopus maternal mRNAs from a dorsal animal blastomere induce a secondary axis in host embryos.</a> Development. 116, 347-3345 (1992).</li><li>Godsave, S. F., Slack, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Single+cell+analysis+of+mesoderm+formation+in+the+Xenopus+embryo.">Single cell analysis of mesoderm formation in the Xenopus embryo.</a> Development. 111, 523-530 (1991).</li><li>Hainski, A. M., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+activin-like+signal+activates+a+dorsal-specifying+RNA+between+the+8-+and+16-cell+stages+of+Xenopus.">An activin-like signal activates a dorsal-specifying RNA between the 8- and 16-cell stages of Xenopus.</a> Dev. Genetics. 19, 210-221 (1996).</li><li>Pandur, P. D., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+maternal+influences+on+dorsal-ventral+fate+of+Xenopus+animal+blastomeres.">Multiple maternal influences on dorsal-ventral fate of Xenopus animal blastomeres.</a> Dev. Dyn. 225, 581-587 (2002).</li><li>Sive, H. L., Grainger, R. M., Harland, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Early+Development+of+Xenopus+laevis.">Early Development of Xenopus laevis.</a> A Laboratory Manual. , (2000).</li><li>Sullivan, S. A., Akers, L., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=foxD5a,+a+Xenopus+winged+helix+gene,+maintains+an+immature+neural+ectoderm+via+transcriptional+repression+that+is+dependent+upon+the+C-terminal+domain.">foxD5a, a Xenopus winged helix gene, maintains an immature neural ectoderm via transcriptional repression that is dependent upon the C-terminal domain.</a> Dev. Biol. 232, 439-457 (2001).</li><li>Yan, B., Neilson, K. M., Moody, S. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=FoxD5+plays+a+critical+upstream+role+in+regulating+neural+fate+and+onset+of+differentiation.">FoxD5 plays a critical upstream role in regulating neural fate and onset of differentiation.</a> Dev. Biol. 329, 80-95 (2009).</li><li>Vincent, J. -. P., Gerhart, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcortical+rotation+in+Xenopus+eggs:+an+early+step+in+embryonic+axis+specification.">Subcortical rotation in Xenopus eggs: an early step in embryonic axis specification.</a> Dev. Biol. 123, 526-539 (1987).</li><li>Peng, H. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Appendix+A:+Solutions+and+Protocols.">Appendix A: Solutions and Protocols.</a> Methods in Cell Biology. 36, 657-662 (1991).</li><li>Klein, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+first+cleavage+furrow+demarcates+the+dorsal-ventral+axis+in+Xenopus+embryos.">The first cleavage furrow demarcates the dorsal-ventral axis in Xenopus embryos.</a> Dev. Biol. 120, 299-304 (1987).</li><li>Masho, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Close+correlation+between+the+first+cleavage+plane+and+the+body+axis+in+early+Xenopus+embryos.">Close correlation between the first cleavage plane and the body axis in early Xenopus embryos.</a> Dev. Growth Diff. 32, 57-64 (1990).</li></ol>

No conflicts of interest declared.

The most critical steps for successful culturing of individual blastomeres are: 1) correctly identifying the blastomere of interest; 2) maintaining a sterile, healthy culture; 3) dissecting cells at the correct part of the cell cycle; and 4) developing the necessary manual dexterity to prevent damage during dissection and transfer to the culture well.
To manipulate specific blastomeres, it is essential to be able to identify the cardinal axes. In Xenopus embryos, the dorsal side can be identified by three methods: (1) marking the sperm entry point with a vital dye 18, 19; (2) tipping in vitro fertilized eggs and marking one side 18, 19; or (3) selecting embryos during the 2-cell stage in which the first cleavage furrow bisects the lightly pigmented region in the animal hemisphere 20, 21. We use the third method, which accurately identifies blastomeres from naturally-fertilized 16-cell embryos in about 90% of cases 20, 21. At fertilization, the animal hemisphere pigmentation begins to contract towards the sperm entry point on the ventral side, causing the dorsal animal region to become less pigmented in a majority of embryos (Figure 1). There is considerable variation in the extent of this lightly pigmented region across clutches and even within clutches of Xenopus eggs. If the first cleavage furrow bisects this lighter area equally between the two daughter cells, then that lighter area can be used as the indicator of the dorsal side, and the first cleavage furrow will indicate the mid-sagittal plane. If the first cleavage furrow separates the daughters into one dark cell and one light cell, do not use it for the explant because the pigmentation at later stages will not accurately identify the dorsal side. Previous studies showed that in these kinds of embryos the first cleavage furrow remained a marker for the mid-sagittal plane whereas the lightly pigmented region no longer indicated the dorsal side 20, 21. Embryos can also be selected at the early part of the 4-cell cleavage (left embryo in Figure 2B), when the first and the second furrows at the vegetal pole can still be distinguished; the first furrow should be complete and the second furrow not yet complete. If, however, one waits until the end of the 4-cell stage to select embryos (right embryo in Figure 2B), the first and second cleavage furrows can no longer be distinguished, and the lightly pigmented cells may be dorsal ones in only about 70% of embryos 20, 21. This will render targeting specific blastomeres much less accurate. It should be noted that the percentage of embryos in which the first cleavage furrow bisects the lightly pigmented region varies a great deal across clutches. In our experience they can account for &lt;50% of the eggs in a clutch to &gt;80% of the eggs in a clutch. Regardless, there almost always are enough eggs within a clutch of healthy eggs that meet this criterion to support an explant experiment.
The two recommended culture media (MMR, MBS) are virtually interchangeable; the preferences of different labs are mostly historical. MMR can be stored as a 10X solution for a year or more. MBS has a shorter shelf life because it is buffered with bicarbonate. Because embryos and explants can succumb to bacterial infection without the protective jelly and vitelline membranes, forceps and culture media should be sterile. If explants do not survive well, antibiotic can be added to the culture medium (e.g., 0.1% gentamicin). Solutions containing gentamicin have a short storage life, and thus should be made fresh on the day they are to be used. Cellular debris contains proteases that will damage the unprotected explants, and promotes bacterial growth. Therefore, it should be removed from the culture wells after the explants have had a chance to heal.
Blastomere dissections are performed in diluted culture medium (0.1X MBS or 0.1X MMR) in order to lessen the strength of cell-to-cell adhesions. During Xenopus cleavage stages, cell adhesion is mostly accomplished by calcium-/magnesium-dependent cadherins. Therefore, it is not necessary to use enzymatic dissociation; in fact this should be avoided because it removes surface proteins that may be important in fate determination events. Because embryonic cells from different batches of eggs adhere with different strengths, the dilution of the dissection medium may need to be adjusted on an ad hoc basis. If the cells are too adherent to dissect without tearing them, lower the concentration of the dissection medium stepwise from 0.1X. Conversely, if the cells fall apart when the vitelline membrane is removed and are very leaky, increase the concentration of the dissection medium stepwise. However, do not use calcium-/magnesium-free media because the cells become too fragile to transfer to the culture wells. Successful separation of blastomeres also depends on when during the cell cycle one performs the dissection. As shown in Figure 2, early in the cell cycle the cleavage furrows are shallow and the blastomeres are flattened. Later in the cycle the cleavage furrows are deeper and the cells are more rounded. Early in the cell cycle, there are cytoplasmic bridges that will cause cytoplasmic leakage if broken. Therefore, cells are more easily dissected late in the cell cycle. 4-cell embryos, 8-cell embryos, and vegetal cells at all stages are harder to dissect because the cytoplasmic bridges persist longer. One can anticipate a high death rate for these explants, meaning one needs to perform more dissections to recover a reasonable sample size. One can wait until the beginning of the next cleavage furrow to begin the dissection of these cells to ensure that the cytoplasmic bridges have closed. While single blastomeres taken from 8- and 16-cell embryos survive well in culture, in our experience older blastomeres fare less well as single cells. Although we do not know the cause of this, survival of older blastomere explants can be improved by dissecting several (2-6) of the same blastomere from different embryos and culturing them together in a single culture well 10, 13, 14.
This procedure requires practice and some manual dexterity. Because the time between cleavages is temperature dependent, it is convenient to store embryos at 14-16 &deg;C to slow down cleavage and allow more time to perform the dissections. However, embryos will not tolerate temperatures lower than 14 &deg;C. The first challenge in this procedure is removing the vitelline membrane. It is best not to move on to blastomere dissection until this first step is mastered. When dissecting out the blastomere, the "handle" cell is likely to leak and fall apart. This is not a concern because once all the other cells are peeled away from the desired cell, the "handle" cell can be removed with forceps. The handle cell remnants as well as other debris usually fall away when the blastomere is transferred to the culture well (Figure 3B). Blastomeres stick avidly to plastic, so use only glass pipettes to transfer them. Use minimal sucking pressure on the cell because it can disintegrate from the shear forces in the pipette. Also, avoid air bubbles in the pipette and make sure the tip of the pipette is below the surface of the culture medium when the explant is expelled because cells will explode if they touch an air-water interface.
The protocol described herein uses unmanipulated embryos as donors for the explants and simple salt media for culturing. These allow one to test the intrinsic ability of the explant to express particular fates. This method can be experimentally expanded in two ways. First, the culture media can be supplemented with signaling/growth factors to test whether these molecules can cause the explant to express alternative fates. Second, the gene expression of the donor embryos can be changed by injecting either mRNAs (gain-of-function) or anti-sense morpholino oligonucleotides (loss-of-function) into specific precursors prior to blastomere dissection. This can be done at the 1-cell stage, or in a more targeted manner at 1-2 cell divisions prior to dissection. With these approaches, one can test whether a particular exogenously supplied gene causes a blastomere to acquire an alternate fate, or whether a particular endogenous gene is necessary for the blastomere to express its autonomous fate (e.g., Figure 4B). These are powerful experimental approaches because they eliminate the confounding influence of the other cells and signaling centers present in the intact embryo.

Xenopus laevis embryos have been utilized extensively to identify the mechanisms by which embryonic cells acquire their specific fates because their eggs are large enough to permit microsurgical approaches. Additionally, they develop externally without the need for nutritional supplementation of the culture medium because each cell contains a rich intracellular supply of yolk platelets that provide an intrinsic energy store. An important asset for studying the mechanisms by which cell fate is determined is the comprehensive set of fate maps of the cleavage stage blastomeres (from 2- through 32-cell stages) 1, 2, 3, 4, 5, 6. These maps were constructed by microinjecting a detectable molecule into a single, identifiable blastomere and monitoring later in development which tissues are populated by the labeled progeny. Consistent fate maps are possible because the cardinal axes of the embryos can be reliably identified in many embryos. First, in all wild type embryos the animal hemisphere is pigmented, whereas the vegetal hemisphere is not. Second, the entry of the sperm at fertilization causes a contraction of the animal hemisphere pigmentation towards the future ventral side; in many embryos a pigmentation difference therefore can be used to discriminate between dorsal and ventral sides. Third, the first cleavage furrow approximates the mid-sagittal plane in most embryos, and thus can be used to identify the right and left sides of the embryo. The fate maps also rely on the fact that naturally fertilized eggs frequently cleave in regular patterns that make each blastomere identifiable across a large population of embryos. While there is variability within and between clutches of eggs regarding pigmentation and cleavage patterns, using selection procedures described herein allows cells with prescribed fates to be identified with about 90% accuracy.
Cell fates can be determined during embryogenesis by several mechanisms. Intrinsic factors, such as differentially inherited cytoplasmic mRNAs or proteins, contribute to several aspects of early patterning. For example, specific maternal mRNAs determine which cells will contribute to the germ line, become the endoderm, or contribute to the dorsal body axis (reviewed in 7). Extrinsic factors provided locally by neighboring cells or more distantly from an embryonic signaling center, are responsible for inducing specific tissue types and patterning nearly every organ system. Examples of signaling centers include the organizer/node in the gastrula that induces the neural ectoderm and patterns the mesoderm, and the zone of polarizing activity that patterns the anterior-posterior axis of the limb bud. Although fate maps identify the precursors of the different organs and reveal the developmental path taken by their descendants in the normal embryo, they cannot distinguish between intrinsic and extrinsic influences on those cells. They also do not reveal the full developmental potential of a cell, whose descendants differentiate in the complex signaling environment of the embryo. Two experimental approaches can test whether a cell's fate is determined by intrinsic factors or is subsequently influenced by external factors: 1) transplantation of the cell to a novel location in the embryo; or 2) removal of the cell from the embryo followed by culture in the absence of exogenous signals.
Both experimental approaches have been feasible in Xenopus because the cells are large enough to be manually separated. For example, numerous studies have deleted single cells from embryos (to change cell-cell interactions) or transplanted cells to novel locations in host embryos to test for fate changes (reviewed in 7, 8, 9). In addition, the second approach of explanting small numbers of cells from different regions of the embryo into culture to elucidate inductive tissue interactions in the absence of exogenous factors is possible because the cells of the Xenopus embryo are filled with an intracellular nutrient store, the yolk platelets. Therefore, they can be cultured for a few days in a defined salt medium without nutritional or growth factor supplementation of the culture medium. We have used this approach to show that dorsal animal blastomeres have an autonomous ability to produce neural and dorsal mesodermal tissues due to maternally inherited mRNAs 10, 11, and others showed that mesoderm specification of 32-cell blastomeres relies on both intrinsic and extrinsic information 12. An advantage of culturing cells as explants is that the medium can also be supplemented with defined signaling factors to determine which cell-to-cell communication pathway might influence the fate of the explanted cell 12, 13, 14. In addition, one can inject the blastomere prior to culture with mRNA to over-express a gene, or with anti-sense oligonucleotides to prevent the translation of endogenous mRNAs. These, gain- and loss-of-function analyses can identify which molecules are required for an autonomously expressed fate. To analyze the fate of the explant, identification of specific cell types can be performed by standard gene expression (e.g., in situ hybridization, RT-PCR) and immunocytochemical assays. This protocol provides a simple, yet powerful, way to distinguish between intrinsic and extrinsic mechanisms that regulate how an embryonic cell develops into specific tissues.

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 <td>Comments (optional)</td>
 </tr>
 <tr>
 <td>Alumina abrasive film</td>
 <td>Thomas Scientific</td>
 <td>#6775E-38 </td>
 <td>Course (12 &mu;m), for major repairs of forceps tips</td>
 </tr>
 <tr>
 <td>Alumina abrasive film</td>
 <td>Thomas Scientific</td>
 <td>#6775E-46 </td>
 <td>Medium (3 &mu;m), for fine sharpening of forceps tips</td>
 </tr>
 <tr>
 <td>Alumina abrasive film</td>
 <td>Thomas Scientific</td>
 <td>#6775E-54 </td>
 <td>Fine (0.3 &mu;m), for polishing of forceps tips</td>
 </tr>
 <tr>
 <td>Forceps: Dumont, Dumoxel Biologie #5</td>
 <td>Fine Science Tools</td>
 <td>#11252-30</td>
 <td>These have the fine tips that do not need sharpening when first purchased. Corrosive resistant so they can be autoclaved.</td>
 </tr>
 <tr>
 <td>Gentamicin solution</td>
 <td>Sigma</td>
 <td>G1397</td>
 <td>Add to medium on same day as use</td>
 </tr>
 </tbody>
</table>

blastomere explants to test for cell fate commitment during embryonic development

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal which tissues descend from each cell of the embryo. Although fate maps are very useful for identifying the precursors of an organ and for elucidating the developmental path by which the descendant cells populate that organ in the normal embryo, they do not illustrate the full developmental potential of a precursor cell or identify the mechanisms by which its fate is determined. To test for cell fate commitment, one compares a cell's normal repertoire of descendants in the intact embryo (the fate map) with those expressed after an experimental manipulation. Is the cell's fate fixed (committed) regardless of the surrounding cellular environment, or is it influenced by external factors provided by its neighbors? Using the comprehensive fate maps of the Xenopus embryo, we describe how to identify, isolate and culture single cleavage stage precursors, called blastomeres. This approach allows one to assess whether these early cells are committed to the fate they acquire in their normal environment in the intact embryo, require interactions with their neighboring cells, or can be influenced to express alternate fates if exposed to other types of signals.

The ability of this assay to accurately assess the developmental potential of the cell relies upon dissecting out the correct blastomere based on the fate maps 1, 2, 3, 4, 5, 6. Therefore, it is critical to choose embryos with the correct pigmentation pattern at the 2-cell stage, as illustrated in Figure 1, that subsequently conform to regular cleavage patterns, as illustrated in Figure 2. If one observes the sorted embryos as they reach the required cleavage stage and separates out the embryos showing irregular cleavages, accuracy is greatly improved. As illustrated in Figure 2, regular cleavages often occur only on one side of the embryo. These embryos can be used as donors if the donor cell originates from the "regular" side. Because regular cleavage patterns are found in fewer embryos as cell division proceeds, sort about five times as many embryos as you intend to dissect. Naturally fertilized eggs tend to have more regular cleavages than in vitro fertilized eggs. Although cleavage patterns vary a great deal within a single clutch of eggs, one can expect at least 10-20% to be useful for this kind of a manipulation.
Blastomeres can be cultured alone or in small groups. When blastomeres are first dissected (Figure 3A), and transferred to the culture well (Figure 3B, C), they are soft and fragile. In our experience, animal and equatorial blastomeres are much easier to dissect and culture than vegetal blastomeres (Figure 3C). Perhaps because they are smaller, they are easier to transfer. They also seem to heal more quickly when nicked with the forceps. Vegetal cells often seem to complete cytokinesis more slowly and thus retain cytoplasmic bridges with sister cells, making them more "leaky" when dissected, and their cell membrane is more fragile and easier to damage. After about 1-2 hr in culture, the explanted blastomere(s) will have divided about 4 times and sloughed off most of the debris remaining from damaged neighboring cells (Figure 3D). After 20 hr, they form small, sturdy balls of cells (Figure 3E). While some may be buried in debris, often a healthy mass can be found (Figure 3E, left).
Although the explants are small, they survive processing for in situ hybridization as well as the more commonly used animal cap explants, which allows one to monitor later gene expression. In the experiment shown in Figure 4A, two dorsal blastomeres (D1.1) or two ventral blastomeres (V1.1) were dissected at the 16-cell stage and cultured in 1X MBS at 14 &deg;C until sibling embryos reached early neural plate stages (st12-13; about 20-22 hr after dissecting). Each sample was assayed for the expression of a zygotic neural plate gene (sox2) by in situ hybridization (the blue reaction product). This assay demonstrates that the majority of dorsal animal blastomeres, which are precursors of the neural plate, can express sox2 autonomously, that is, without additional signaling from other regions of the embryo. Those explants that do not express sox2 may have been incorrectly identified at the time of dissection. In contrast, none of the ventral animal blastomeres, which are precursors of the ventral epidermis, express sox2. Thus, we conclude that dorsal animal blastomere have an intrinsic ability to form neural tissue, whereas the ventral animal blastomeres do not. Note that some of the D1.1 explants have also eIongated, a morphology indicative of dorsal mesoderm formation; these blastomeres also are fated to form the notochord in the intact embryo 3, and in explant culture express a notochord marker protein 10. Blastomere intrinsic fates can be altered by prior microinjection of mRNAs (gain-of-function) or anti-sense morpholino oligonucleotides (MOs; loss-of-function). In Figure 4B, the effect of altering the endogenous levels of a maternally expressed neural gene, foxD5 16, 17, is tested. When the endogenous level of FoxD5 was increased by injecting foxD5 mRNAs into the 8-cell precursor blastomeres of V1.1 (bottom row), the majority of the V1.1 explants (cultured to st 12-13) expressed sox2. Conversely, when the endogenous level of FoxD5 was reduced by injection of MOs into the 8-cell precursor blastomeres of D1.1 (top row), only a few D1.1 explants (cultured to st 12-13) expressed sox2 (compare to top row of Figure 4A). These experiments indicate that the maternal expression of foxD5 plays a role in the intrinsic ability of dorsal blastomeres to acquire a neural fate.
<img src="/files/ftp_upload/4458/4458fig1.jpg" alt="Figure 1"/> 
Figure 1. Pigmentation patterns in 2-cell embryos from naturally fertilized eggs. The first cleavage furrow is indicated by an arrow. A) An example of an embryo that should not be used to identify blastomeres at later stages because the lightly pigmented region of the animal hemisphere of the embryo is contained mostly in one cell (on the left). Since the first cleavage furrow predicts the mid-sagittal plane of the embryo, the lightly pigmented region of this embryo will not identify dorsal. B) Two examples of embryos that should be sorted for explant cultures. In both, the lightly pigmented region of the animal hemisphere (*) is bisected by the first cleavage furrow. The embryo on the left is early in the first cell cycle, and the lightly pigmented region is only visible in about 25% of the cell surface. The embryo on the right is nearing the end of the first cell cycle, and the pigment has moved ventrally so the dorsal halves of the two cells are lighter. As a result, at later stages the lightly pigmented animal cells will predict dorsal (d) and darkly pigmented animal cells will predict ventral (v).
<img src="/files/ftp_upload/4458/4458fig2.jpg" alt="Figure 2"/> 
Figure 2. Examples of cleavage stage embryos from naturally fertilized eggs. All embryos are oriented with the animal pole facing the reader, dorsal to the top. A) 4-cell embryos. The embryo on the left has just entered the second cell cycle so the cleavage furrows are shallow. The embryo on the right has nearly completed the second cell cycle so the cleavage furrows are deeper and the cells appear more rounded. Black arrows point to the first cleavage furrow, and red arrows point to the second cleavage furrow. B) 8-cell embryos. The embryo on the left is early is the third cell cycle and the embryo on the right is near the end of the third cell cycle. C) 16-cell embryos. C' is an example of irregular cleavages to be avoided. C" is an example of regular cleavages on the ventral side, but irregular on the dorsal side. C''' is an example of more regular cleavages on the dorsal side, but not on ventral side. C'''' is an example of regular cleavages at the midline, but not laterally. D) 32-cell embryos. D' is an example of regular cleavages on the ventral side, but irregular on the dorsal side. D'' is an example of regular cleavages on the dorsal side, but not on ventral side. D''' is an example of regular cleavages on the right side, but more irregular cleavages on the left side. E) 64-cell embryo. For a schematic representation of these stages, please refer to the Nieuwkoop and Faber staging series, which can be found on-line (<a href="http://www.xenbase.org/anatomy/alldev.do" target="_blank">http://www.xenbase.org/anatomy/alldev.do</a>).
<img src="/files/ftp_upload/4458/4458fig3.jpg" alt="Figure 3"/> 
Figure 3. Explants from 16-cell embryos from naturally fertilized eggs. A) A ventral animal blastomere shortly after dissection. Arrow denotes remnants of the "handle" cell. B) A ventral animal blastomere transferred to a culture well that has divided once. C) A ventral vegetal blastomere shortly after dissection. Compared to animal blastomeres, these are larger, and more difficult to manipulate. D) Two ventral animal blastomere explants, 2 hr after dissection, have divided about four times. The left explant is viewed from the original, pigmented outer surface of the blastomere; the right explant is viewed from the original, unpigmented inner surface of the blastomere. E) Two ventral animal blastomere explants cultured for 24 hr. The left explant is sitting in a bed of cellular debris (arrows) whereas the right explant is free of any detritus. All images are at 50X magnification.
<img src="/files/ftp_upload/4458/4458fig4.jpg" alt="Figure 4" fo:content-width="4.8in" fo:src="/files/ftp_upload/4458/4458fig4highres.jpg"/> 
Figure 4. In situ hybridization analysis of gene expression in explants derived from 16-cell blastomeres dissected from naturally fertilized eggs. A. The 16-cell embryo on left illustrates the dorsal blastomere pair (D1.1) that was explanted for the results in the top row, and the ventral blastomere pair (V1.1) that was explanted for the results in the bottom row. Dissected blastomere pairs from uninjected embryos were placed in 0.1X MBS in culture wells and incubated at 14 &deg;C for about 20 hr. They were collected in tubes of fixative and processed by in situ hybridization for expression of a zygotic neural gene, sox2. B. Blastomeres of 8-cell stage embryos were injected bilaterally either with foxD5 MOs, to reduce endogenous expression in dorsal blastomeres, or with foxD5 mRNA to increase endogenous expression in ventral blastomeres. When the injected embryos reached the 16-cell stage (20-30 min later), blastomere pairs were dissected, cultured and analyzed by in situ hybridization as in A. The number of explants that express sox2 is significantly reduced in D1.1 explants that were injected with foxD5 MOs (compare to top row in A). The number of explants that express sox2 is significantly increased in V1.1 explants that were injected with foxD5 mRNA (compare to bottom row in A). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/4458/4458fig4large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development at JoVE.com

Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Fate maps, constructed from lineage tracing all of the cells of an embryo, reveal  which tissues descend from each cell of the embryo.  Although fate maps ...

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15.2K Views.  The George Washington University. The fate of an individual embryonic cell can be influenced by inherited molecules and/or by signals from neighboring cells. Utilizing fate maps of the cleavage stage Xenopus embryo, single blastomeres can be identified for culture in isolation to assess the contributions of inherited molecules versus cell-cell interactions.

Video: Blastomere Explants to Test for Cell Fate Commitment During Embryonic Development

Growth Factor-Coated Bead Placement on Dorsal Forebrain Explants

This video demonstrates two methods for preparing and placing beads, which have been coated with growth factor, on explants of the developing cerebral cortex. These beads can be used to induce spatially restricted gene expression on developing neural tissue such as forebrain explants. Methods are given for using both Affi-Gel beads and heparin acryllic beads.

Ole Isacson: Development of New Therapies for Parkinson's Disease

Ole Isacson gives a concise overview of Parkinsons's disease, its causes, therapeutic strategies, and advances in Parkinson's research.

Culture and Maintenance of Human Embryonic Stem Cells  

Human embryonic stem (hES) cells must be monitored and cared for in order to maintain healthy, undifferentiated cultures. At minimum, the cultures must be  fed  every day by performing a complete medium change to replenish lost nutrients and to keep the cultures free of unwanted differentiation factors. Although a small amount of differentiation is normal and expected in stem cell cultures, the culture should be routinely cleaned up by manually removing, or "picking" differentiated areas. Identifying and removing excess differentiation from hES cell cultures are essential techniques in the maintenance of a healthy population of cells.

Culture and Maintenance of Human Embryonic Stem Cells

This protocol demonstrates how to maintain healthy, undifferentiated human embryonic stem (ES) cells.

Human embryonic stem (hES) cells must be monitored and cared for in order to maintain healthy, undifferentiated cultures. At minimum, the cultures must be ...

Freezing and Thawing Human Embryonic Stem Cells

Since James Thomson et al developed a technique in 1998 to isolate and grow hES in culture, freezing cells for later use and thawing and expanding cells from a frozen stock have become important procedures performed in routine hES cell culture. Since hES cells are very sensitive to the stresses of freezing and thawing, special care must taken. Here we demonstrate the proper technique for rapidly thawing hES cells from liquid nitrogen stocks, plating them on mouse embryonic feeder cells, and slowly freezing them for long-term storage.

Since James Thomson et al developed a technique in 1998 to isolate and grow hES in culture, freezing cells for later use and thawing and expanding cells from a frozen stock have become important procedures performed in routine hES cell culture. Since hES cells are very sensitive to the stresses of freezing and thawing, special care must taken. Here we demonstrate the proper technique for rapidly thawing hES cells from liquid nitrogen stocks, plating them on mouse embryonic feeder cells, and slowly freezing them for long-term storage.

Since James Thomson et al developed a technique in 1998 to isolate and grow hES in culture, freezing cells for later use and thawing and expanding cells ...

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing and adult tissue. An advance in this concept is Genetic Inducible Fate Mapping (GIFM), linking gene expression, cell fate, and cell behaviors in vivo, to create fate maps based on genetic lineage.
GIFM exploits X-CreER lines where X is a gene or set of gene regulatory elements that confers spatial expression of a modified bacteriophage protein, Cre recombinase (CreERT). CreERT contains a modified estrogen receptor ligand binding domain which renders CreERT sequestered in the cytoplasm in the absence of the drug tamoxifen. The binding of tamoxifen releases CreERT, which translocates to the nucleus and mediates recombination between DNA sequences flanked by loxP sites. In GIFM, recombination typically occurs between a loxP flanked Stop cassette preceding a reporter gene such as GFP.
Mice are bred to contain either a region- or cell type-specific CreER and a conditional reporter allele. Untreated mice will not have marking because the Stop cassette in the reporter prevents further transcription of the reporter gene. We administer tamoxifen by oral gavage to timed-pregnant females, which provides temporal control of CreERT release and subsequent translocation to the nucleus removing the Stop cassette from the reporter. Following recombination, the reporter allele is constitutively and heritably expressed. This series of events marks cells such that their genetic history is indelibly recorded. The recombined reporter thus serves as a high fidelity genetic lineage tracer that, once on, is uncoupled from the gene expression initially used to drive CreERT.
We apply GIFM in mouse to study normal development and ascertain the contribution of genetic lineages to adult cell types and tissues. We also use GIFM to follow cells on mutant genetic backgrounds to better understand complex phenotypes that mimic salient features of human genetic disorders.
This video article guides researchers through experimental methods to successfully apply GIFM. We demonstrate the method using our well characterized Wnt1-CreERT;mGFP mice by administering tamoxifen at embryonic day (E)8.5 via oral gavage followed by dissection at E12.5 and analysis by epifluorescence stereomicroscopy. We also demonstrate how to micro-dissect fate mapped domains for explant preparation or FACS analysis and dissect adult fate-mapped brains for whole mount fluorescent imaging. Collectively, these procedures allow researchers to address critical questions in developmental biology and disease models.

A Practical Approach to Genetic Inducible Fate Mapping: A Visual Guide to Mark and Track Cells In Vivo

Genetic Inducible Fate Mapping (GIFM) marks and tracks cells with fine spatial and temporal control in vivo and elucidates how cells from a specific genetic lineage contribute to developing and adult tissues. Demonstrated here are the techniques required to fate map E12.5 mouse embryos for epifluorescent and explant analysis.

Fate maps are generated by marking and tracking cells in vivo to determine how progenitors contribute to specific structures and cell types in developing ...

A Reverse Genetic Approach to Test Functional Redundancy During Embryogenesis

Gene function during embryogenesis is typically defined by loss-of-function experiments, for example by targeted mutagenesis (knockout) in the mouse. In the zebrafish model, effective reverse genetic techniques have been developed using microinjection of gene-specific antisense morpholinos. Morpholinos target an mRNA through specific base-pairing and block gene function transiently by inhibiting translation or splicing for several days during embryogenesis (knockdown). However, in vertebrates such as mouse or zebrafish, some gene functions can be obscured by these approaches due to the presence of another gene that compensates for the loss. This is especially true for gene families containing sister genes that are co-expressed in the same developing tissues. In zebrafish, functional compensation can be tested in a relatively high-throughput manner, by co-injection of morpholinos that target knockdown of both genes simultaneously. Likewise, using morpholinos, a genetic interaction between any two genes can be demonstrated by knockdown of both genes together at sub-threshold levels. For example, morpholinos can be titrated such that neither individual knockdown generates a phenotype. If, under these conditions, co-injection of both morpholinos causes a phenotype, a genetic interaction is shown. Here we demonstrate how to show functional redundancy in the context of two related GATA transcription factors. GATA factors are essential for specification of cardiac progenitors, but this is revealed only by the loss of both Gata5 and Gata6. We show how to carry out microinjection experiments, validate the morpholinos, and evaluate the compensated phenotype for cardiogenesis.

Gene function can be obscured in loss-of-function experiments if there is compensation by another gene. The zebrafish model provides a relatively high-throughput means to reveal such functional redundancy in living embryos.

Gene function during embryogenesis is typically defined by loss-of-function experiments, for example by targeted mutagenesis (knockout) in the mouse. In ...

Fate Mapping of Human Embryonic Stem Cells by Teratoma Formation 

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three embryonic germ layers (1). Directed differentiation of hESCs into specific cell types has generated much interest in the field of regenerative medicine (e.g., (2-5)), and methods for determining the in vivo fate of selected or manipulated hESCs are essential to this endeavor. We have adapted a highly efficient teratoma formation assay for this purpose. A small number of specifically selected hESCs is mixed with undifferentiated wild type hESCs and Phaseolus vulgaris lectin to form a cell pellet. This is grafted beneath the kidney capsule in an immunodeficient mouse. As few as 2.5 x 105 hESCs are needed to form a 16 cm3 teratoma within 8-12 weeks. The fate of the originally selected hESCs can then be determined by immunohistochemistry. This method provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Fate Mapping of Human Embryonic Stem Cells by Teratoma Formation

Directed differentiation of hESCs into specific cells has generated much interest in regenerative medicine. We provide a concise, step-by-step protocol for determining the in vivo fate of selected hESCs that provides a valuable tool for characterizing tissue-specific reagents for cell-based therapy.

Human embryonic stem cells (hESCs) have an unlimited capacity for self-renewal, and the ability to differentiate into cells derived from all three ...

Single Cell Fate Mapping in Zebrafish

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

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

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

A System for ex vivo Culturing of Embryonic Pancreas

The pancreas controls vital functions of our body, including the production of digestive enzymes and regulation of blood sugar levels1. Although in the past decade many studies have contributed to a solid foundation for understanding pancreatic organogenesis, important gaps persist in our knowledge of early pancreas formation2. A complete understanding of these early events will provide insight into the development of this organ, but also into incurable diseases that target the pancreas, such as diabetes or pancreatic cancer. Finally, this information will generate a blueprint for developing cell-replacement therapies in the context of diabetes.
 During embryogenesis, the pancreas originates from distinct embryonic outgrowths of the dorsal and ventral foregut endoderm at embryonic day (E) 9.5 in the mouse embryo3,4. Both outgrowths evaginate into the surrounding mesenchyme as solid epithelial buds, which undergo proliferation, branching and differentiation to generate a fully mature organ2,5,6. Recent evidences have suggested that growth and differentiation of pancreatic cell lineages, including the insulin-producing &beta;-cells, depends on proper tissue-architecture, epithelial remodeling and cell positioning within the branching pancreatic epithelium7,8. However, how branching morphogenesis occurs and is coordinated with proliferation and differentiation in the pancreas is largely unknown. This is in part due to the fact that current knowledge about these developmental processes has relied almost exclusively on analysis of fixed specimens, while morphogenetic events are highly dynamic.
 Here, we report a method for dissecting and culturing mouse embryonic pancreatic buds ex vivo on glass bottom dishes, which allow direct visualization of the developing pancreas (Figure 1). This culture system is ideally devised for confocal laser scanning microscopy and, in particular, live-cell imaging. Pancreatic explants can be prepared not only from wild-type mouse embryos, but also from genetically engineered mouse strains (e.g. transgenic or knockout), allowing real-time studies of mutant phenotypes. Moreover, this ex vivo culture system is valuable to study the effects of chemical compounds on pancreatic development, enabling to obtain quantitative data about proliferation and growth, elongation, branching, tubulogenesis and differentiation. In conclusion, the development of an ex vivo pancreatic explant culture method combined with high-resolution imaging provides a strong platform for observing morphogenetic and differentiation events as they occur within the developing mouse embryo.

A System for ex vivo Culturing of Embryonic Pancreas

Here, we describe a method for isolation, culture and manipulation of mouse embryonic pancreas. This represents an excellent ex vivo system for studying various aspects of pancreatic development, including morphogenesis, differentiation and growth. Pancreatic bud explants can be cultured for several days and used in a range of different applications, including whole-mount immunofluorescence and live imaging.

The pancreas controls vital functions of our body,  including the production of digestive enzymes and regulation of blood sugar  levels1. Although in the ...

Cell Tracking Using Photoconvertible Proteins During Zebrafish Development

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of genetically-encoded fluorescent proteins has proven a valuable strategy for elucidating dynamic morphogenetic processes as they occur in the intact organism. During the past decade, the development of photoactivatable and photoconvertible fluorescent proteins has opened the possibility to investigate the fate of discrete subpopulations of tagged proteins1. Unlike photoactivatable proteins, photoconvertible fluorescent proteins (PCFPs) are readily tracked and imaged in their native emission state prior to photoconversion, making it easier to identify and select regions by optical inspection. PCFPs, such as Kaede2, KikGR3, Dendra4 and EosFP5, can be shifted from green to red upon exposure to UV or blue light due to a His-Tyr-Gly tripeptide sequence which forms a green chromophore that can be photoconverted to a red one by a light-catalyzed &beta;-elimination and subsequent extension of a &pi;-conjugated system3. PCFPs and their monomeric variants are useful tools for tracking cells6-10 and studying protein dynamics11-14, respectively. During recent years, PCFPs have been expressed in different animal model, such as zebrafish6, chicken7,8 and mouse9,10 for cell fate tracking. Here we report a protocol for cell-specific photoconversion of PCFPs in the living zebrafish embryo and further tracking of photoconverted proteins at later developmental stages. This methodology allows studying, in a tissue-specific manner, cell biological events underlying morphogenesis in the zebrafish animal model.

Here, we present a method for the photoactivated switch of photoconvertible fluorescent proteins (PCFPs) in the living zebrafish embryo and further tracking of photoconverted protein at specific time points during development. This methodology allows monitoring of cell biological events underlying different developmental processes in a live vertebrate organism.

Embryogenesis is a dynamic process that is best studied by using techniques that allow the documentation of developmental changes in vivo. The use of ...