We thank members of the Mariani lab for help editing the protocol. This work was funded by a CIRM Postdoctoral Training Grant (JLF), a CIRM BRIDGES internship (TZTT), the Robert E. and May R. Wright Foundation (FVM), and the University of Southern California (FVM).

1. LysoTracker Staining of Mouse Embryos
 <ol>
 <li>Generate mouse embryos by placing young (5-6 week old) females into male stud cages. Monitor on following mornings for the appearance of a vaginal plug indicating that mating has occurred. If the female is pregnant, noon on that day is called 0.5 dpc (days post coitus). The procedure presented here is ideal for embryos 7-13 dpc.</li>
 <li>On the embryonic day of interest, euthanize the female according to approved protocols and remove the uterus. Remove the embryos from the decidua in a 10 cm Petri dish with Hanks BSS (without phenol red). Remove the extra-embryonic membranes and do at least one rinse with Hanks BSS to remove extraneous tissues and blood.</li>
 <li>Prepare LysoTracker Embryo Staining Solution (5 &mu;M LysoTracker in Hanks BSS). A convenient volume for 1-2 mouse litters is 5 ml. Gently mix to disburse the dye evenly.</li>
 <li>Add embryos to the LysoTracker Embryo Staining Solution in a microfuge tube or vial and incubate at 37 &deg;C for 45 min.</li>
 <li>Rinse very gently in Hanks BSS four times, 5 min each wash.</li>
 <li>Fix in 4% Paraformaldehyde overnight.</li>
 <li>Rinse once in Hanks BSS, 10 min, to remove the fix.</li>
 <li>Dehydrate through a methanol series (50%, 75%, 80%, 100%, 5 min each step) to eliminate background staining. This is important for achieving good signal-to-noise during imaging.</li>
 <li>At this point you can store the embryo samples indefinitely in methanol at -20 &deg;C, protected from light.</li>
 </ol>
 Note: A tissue sample can be taken at any step for DNA genotyping, however, it can be most convenient to take a sample just before storage so that up to this step all the samples can be batch-processed. Collect a sample of tissue from the tail, limb, or head from each embryo and put all into separate tubes. Rehydrate the tissue sample and prepare for genotyping.
 2. LysoTracker Staining of Differentiating ES Cultures
 <ol>
 <li>Prepare embryoid bodies via the Hanging Drop Method15 with a starting density of 500 cells per 20 &mu;l drop.</li>
 <li>After 3 days, transfer the embryoid bodies to suspension culture on 10 cm non-adherent Petri dishes (i.e. those used for bacteriological work). Culture for 5 days, changing the media every 2 days.</li>
 <li>On day 7, transfer the embryoid bodies to gelatinized 8-well chamber slides (add 0.1% Gelatin to the chambers, wait 5 min, remove, and add media). Transfer 1-2 embryoid bodies per chamber.</li>
 <li>Culture for 10 days, changing media every other day. Be careful to place the pipette at the corner of the chamber slide so as to not perturb the embryoid body attaching to the glass surface. Also when adding fresh media also place the pipette at the corner of the chamber and release the media slowly.</li>
 <li>After 10 days of culture, to induce apoptosis as a positive control, treat some of the wells with 0.1 mM and 1.0 mM H2O2 (add the H2O2 to fresh media, mix, then add to the cells) for 60 min. Rinse twice with D-PBS and culture overnight in regular media.</li>
 <li>Gently aspirate existing media and add LysoTracker Cell staining solution (500 nM LysoTracker (final concentration) in media, 300 &mu;l per chamber).</li>
 <li>Incubate at 37 &deg;C for 15 min.</li>
 <li>Rinse twice with D-PBS 2 times, 5 min each.</li>
 <li>Fix in 4% paraformaldehyde for 15 min at RT.</li>
 <li>Rinse twice with D-PBS, 5 min.</li>
 </ol>
 3. Viewing LysoTracker Stained Embryos or ES Cultures
 <ol>
 <li>Prepare the cell samples for microscopy by aspirating the D-PBS; removing the chamber off the slide; air drying for about 5 min; and then adding an aqueous mounting medium (Vectashield with DAPI or similar). Place a foil barrier while drying to decrease bleaching of the fluorophore. Mount embryos in a deep-depression glass dish or small Petri dish in Vectashield or similar. </li>
 <li>Visualize with a dissecting or compound microscope outfitted with a rhodamine or Texas Red filter (LysoTracker RED DND-99, Excitation/Emission: 577/590 nm). The staining is typically quite bright so long exposures are usually not necessary.</li>
 </ol>
 4. Quantification of Results Using Imaging Techniques
 <ol>
 <li>Take digital photos of your samples under the same exposure conditions in manual mode.</li>
 <li>Open the images in Adobe Photoshop or a similar imaging program. To determine the amount of LysoTracker staining, select the red channel and threshold the image (converts the image to black-and-white) such that the red staining is now represented by white pixels. Select the white pixels and use the histogram function to obtain the total count.</li>
 <li>Use the blue channel to count the number of nuclei in the field. It can be useful to keep a record of the count by annotating the image.</li>
 <li>Record the values and then calculate the average staining level per cell. Repeat this analysis with the same threshold setting for all images you wish to sample.</li>
 </ol>
 5. Representative Results
 There is a normal pattern of PCD during embryonic development that can change when a growth factor gene, such as Sonic hedgehog (Shh), is removed16,17 . Since LysoTracker is a small molecule dye that is highly diffusible, the whole embryo can be visualized to assess areas of PCD even deep within the mesenchyme12,13. Figure 1 shows the LysoTracker staining pattern for normal and Shh-/- mutant mouse embryos at 10.5dpc. At this stage, PCD is greatly increased in Shh-/- mutant embryos, particularly in the limb and somite regions. Portions of the embryo can be imaged in whole-mount at higher magnification (1B) to assess the overall pattern. Dissected portions can also be sectioned with a vibrating microtome (1C) or a cryostat (D-F) to reveal PCD regions in greater detail.
 TUNEL analysis is best done on thin sections where penetration of the terminal transferase is less problematic. LysoTracker staining and a TUNEL assay can be done in the same tissue (Figure 1D-G). These results show that although there is not a strict 1:1 correlation between LysoTracker and TUNEL staining, the general regions of PCD overlap considerably. At a higher magnification, it is possible to see puncta that stain positive with the TUNEL assay but not with the LysoTracker dye, puncta that stain only with the LysoTracker dye, and puncta that have overlapping staining.
 When an EB is plated on a gelatin-coated surface, differentiating cells will migrate outward. After 10 days of culture, cells will have differentiated into a variety of cell types including neurons, muscle cells, and cardiomyocytes. The cultures shown in Figure 2 contain mixed populations of these cell types, however one could adapt our protocol to study PCD of a particular cell-type of interest. For example, the EB could be treated with Retinoic Acid to enhance the differentiation of neurons19 , and then the LysoTracker assay could be used to assess neuronal cell death under different conditions. In the normal culture of differentiated ES cells, there are a few cells that undergo PCD and these can be detected with LysoTracker and/or TUNEL assays (Figure 2A). PCD can be induced by a number of different cytotoxic agents such as hydrogen peroxide (H2O2) which induces oxidative damage and initiates the apoptotic pathway20,21 . A low dose of H2O2 increases both LysoTracker and TUNEL staining compared to no treatment, while a high dose results in cell shrinkage and widespread LysoTracker and TUNEL positivity. Similar to the results in the embryo, it is possible to see puncta that stain positive with the TUNEL assay but not with the LysoTracker dye, puncta that stain only with the LysoTracker dye, and puncta that have overlapping staining. The consequence of the application of a cytotoxic agent that induces apopotosis is compared to serum starvation conditions which induce autophagy (Figure 2D). Cells that have undergone a period of serum starvation (1 hour) exhibit a different pattern where LysoTracker and TUNEL staining do not highly correlate. LysoTracker staining is much increased compared to untreated cultures, however TUNEL positivity is at normal levels.
 Employing standard tools available in Adobe Photoshop, LysoTracker staining can be quantified with a simple thresholding method that allows one to calculate the average staining level for a population of cells. Figure 3A-B shows how a sample image looks during the quantification process. Starting with a baseline image with LysoTracker staining in the red channel and the DAPI-stained nuclei in the blue channel (panel 3A), the red channel of this image was thresholded to convert the image to black-and-white and the white pixels were then selected and counted (Figure 3A'). A thresholding level was chosen that represents the LysoTracker staining pattern. As with any quantification method that uses images, it is important to avoid overexposing your photos as this could potentially result in over-emphasizing differences. The number of the nuclei in the field can then be counted by looking at just the blue channel (Figure 3B) and marking the nuclei using a drawing tool such as a paintbrush while counting (Figure 3B'). One can then express the level of staining by calculating the average level of staining per cell (number of pixels/nuclei count). If one wishes to use this algorithm for high through-put screening, most screening packages can measure brightness in a particular channel and also have an object identifier function that will count nuclei in a field, however for most typical applications the manual method is quick and simple. To give an example, a sample region from Figures 2A-C was quantified and the results confirm that increasing concentrations of H2O2 results in a higher level of LysoTracker staining (Figure 3C).
 <img src="/files/ftp_upload/4254/4254fig1.jpg" alt="Figure 1"/> 
 Figure 1. LysoTracker staining in normal and Shh-/- embryos. A,A'. Normal and Shh-/- mouse embryos at 10.5 dpc, stained with LysoTracker Red, mounted in Vectashield and photographed under an epiflourescence dissecting microscope. Brackets indicate the location of the forelimbs. B,B'. Forelimb of normal and Shh-/- embryos imaged at higher magnification. Note the normal domains of cell death including the 'Posterior Necrotic Zone' (marked with an asterisk) and a region in the proximal mesenchyme of the limb bud (marked with an arrow). Shh null limbs exhibit stronger staining in the medial and distal portion. C,C'. Medial sections through the forelimb bud of both normal and null limb buds. Note the presence of staining in the proximal mesenchyme (arrow) and AER (arrowhead) of the normal limb bud and the strong staining within the dorsal and ventral and distal mesenchyme within the null limb bud. D-F. The same section (12 micron, frozen section) was imaged for LysoTracker and TUNEL staining (Roche 1684795). The section passes through the distal portion of the Shh null mesenchyme. The arrowhead points to the AER. Note how the overall pattern is very similar using both methods, however the staining does not overlap completely. G,G'. At a higher magnification of the AER and mesenchyme, one can see both the presence of puncta that are LysoTracker or TUNEL-positive only as well as puncta that have overlapping staining. 
 <img src="/files/ftp_upload/4254/4254fig2.jpg" alt="Figure 2"/> 
 Figure 2. LysoTracker staining in differentiated ES cells. Differentiating cells were plated on 8-chamber slides in the absence of LIF. To show how LysoTracker and TUNEL staining compare, the apoptosis pathway was induced with 2 different doses of H2O2, while the autophagy pathway was induced using 1 hour of serum starvation. These images were taken with a Zeiss LSM5 Confocal Microscope. A-A'': Untreated cultures show some normal apoptosis as indicated by LysoTracker and TUNEL staining. The pattern using both techniques mostly overlaps (white arrows), however, one can see a few puncta that are LysoTracker positive only (red arrows). B-B'', C-C": Treatment with increasing amounts of H2O2 stimulates apoptosis. The lower dose shows more LysoTracker and TUNEL positive puncta when compared to untreated cells. The higher dose induces in even more LysoTracker and TUNEL positive puncta. White arrows point to overlapping staining, while red arrows point to puncta that are positive for LysoTracker only. It is also possible to see puncta that are only positive for TUNEL staining (green arrows). D-D": Serum starvation can induce autophagy, which can stimulate an accumulation of material in lysosomes and autophagolysosomes. Note how many more LysoTracker puncta are visible compared to untreated cultures. TUNEL positive puncta are present (most of which have overlapping LysoTracker staining, white arrows), however, the number is not greatly increased compared to untreated conditions. In addition, under serum starvation conditions, LysoTracker-only positive puncta are highly prevalent (red arrows).
 <img src="/files/ftp_upload/4254/4254fig3.jpg" alt="Figure 3"/> 
 Figure 3. Quantification method to assess the level of staining in a population of cells using Adobe Photoshop. A: A portion of Figure 2A was chosen and the LysoTracker (red channel) and DAPI staining (nuclei, blue channel) is shown in this panel. A': The red channel of Figure 3A was thresholded to convert it to a black-and-white image. The white pixels can then be selected and counted using the histogram function. B-B': The blue channel shows the nuclei and while counting them, the image can be annotated with a drawing function such as a paintbrush. C: This is an example quantification of a sample from panels 2A-C. The data is represented as pixels per nuclei. <a href="https://www.jove.com/files/ftp_upload/4254/4254fig3large.jpg" target="_blank">Click here to view larger figure</a>.

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No conflicts of interest declared.

Important hallmarks of apoptosis include the detection of DNA damage with the TUNEL assay, along with the detection of caspase activity (detected with mAbs or a binding molecule such as ZVAD-fmk), the observation of cellular changes, and the presentation of phosphatidylserine (PS) on the membrane surface (detected by Annexin V binding). There are some disadvantages with each of these assays. For example, the terminal transferase used in the TUNEL assay does not penetrate beyond a few cell layers. Also Annexin V can...

<table border="1">
	<tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>&nbsp;</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>LysoTracker Red DND-99</td>
 <td>Invitrogen</td>
 <td>#L-7528</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Hanks BSS</td>
 <td>Invitrogen</td>
 <td>14025-076</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Paraformaldehyde</td>
 <td>EMD</td>
 <td>EM-PX0055-3</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Vectashield</td>
 <td>VECTOR</td>
 <td>H-1200</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>DMEM</td>
 <td>Cellgro</td>
 <td>10-013-CV</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Non-essential amino acids</td>
 <td>Cellgro</td>
 <td>25-025-CI</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Sodium pyruvate</td>
 <td>Cellgro</td>
 <td>25-000-CI</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>FBS</td>
 <td>Hyclone</td>
 <td>SH30071.02</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Pen-Strep</td>
 <td>Invitrogen</td>
 <td>15140-122</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>b-Mercaptoethanol, 50 mM </td>
 <td>Invitrogen</td>
 <td>21985-023</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>LabTek-II Chamber slides 
 (8-well)</td>
 <td>Nalge Nunc International</td>
 <td>154534</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>0.1% Gelatin</td>
 <td>Millipore</td>
 <td>ES-006-B</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Dulbecco's PBS (D-PBS) </td>
 <td>Cellgro</td>
 <td>21-031-CV</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>Solution Recipes
 4% Paraformaldehyde
 For 100 ml:
 <ol>
 <li>Mix 4 g paraformaldehyde, 90 ml H2O, and NaOH (a drop of 2N NaOH). The paraformaldehyde will not go into solution until you have added some NaOH to increase the pH.</li>
 <li>Stir and heat at 60 &deg;C until all the powder is in solution (~10-20 min). Do not overheat.</li>
 <li>Add ~10 ml 10x PBS to achieve a final volume of 100 ml.</li>
 <li>Store at -20 &deg;C in convenient (~10 ml or ~40 ml) aliquots.</li>
 </ol>
 WARNING: Paraformaldehyde in 'frill' form (compressed small pellets) is less powdery and can therefore be measured outside of a hood. However you should still wear a protective dust mask (N95 at least) during handling.
 EB Culture Media
 For 500 ml EB Media:
 <table border="1">
 <tbody>
 <tr>
 <td>DMEM:</td>
 <td>404.5 ml</td>
 </tr>
 <tr>
 <td>FBS:</td>
 <td>75.0 ml</td>
 </tr>
 <tr>
 <td>L-Glutamine:</td>
 <td>5.0 ml</td>
 </tr>
 <tr>
 <td>Penicillin/Streptomycin:</td>
 <td>5.0 ml</td>
 </tr>
 <tr>
 <td>Non-essential amino acids:</td>
 <td>5.0 ml</td>
 </tr>
 <tr>
 <td>Sodium pyruvate:</td>
 <td>5.0 ml</td>
 </tr>
 <tr>
 <td>&beta;-Mercaptoethanol:</td>
 <td>500 &mu;l</td>
 </tr>
 </tbody>
 </table></td>
 </tr>
	 </tbody>
 </table>

use of lysotracker to detect programmed cell death in embryos and differentiating embryonic stem cells

Programmed cell death (PCD) occurs in adults to maintain normal tissue homeostasis and during embryological development to shape tissues and organs1,2,6,7. During development, toxic chemicals or genetic alterations can cause an increase in PCD or change PCD patterns resulting in developmental abnormalities and birth defects3-5. To understand the etiology of these defects, the study of embryos can be complemented with in vitro assays that use differentiating embryonic stem (ES) cells.
Apoptosis is a well-studied form of PCD that involves both intrinsic and extrinsic signaling to activate the caspase enzyme cascade. Characteristic cell changes include membrane blebbing, nuclear shrinking, and DNA fragmentation. Other forms of PCD do not involve caspase activation and may be the end-result of prolonged autophagy. Regardless of the PCD pathway, dying cells need to be removed. In adults, the immune cells perform this function, while in embryos, where the immune system has not yet developed, removal occurs by an alternative mechanism. This mechanism involves neighboring cells (called "non-professional phagocytes") taking on a phagocytic role-they recognize the 'eat me' signal on the surface of the dying cell and engulf it8-10. After engulfment, the debris is brought to the lysosome for degradation. Thus regardless of PCD mechanism, an increase in lysosomal activity can be correlated with increased cell death.
To study PCD, a simple assay to visualize lysosomes in thick tissues and multilayer differentiating cultures can be useful. LysoTracker dye is a highly soluble small molecule that is retained in acidic subcellular compartments such as the lysosome11-13. The dye is taken up by diffusion and through the circulation. Since penetration is not a hindrance, visualization of PCD in thick tissues and multi-layer cultures is possible12,13. In contrast, TUNEL (Terminal deoxynucleotidyl transferase dUTP nick end labeling) analysis14, is limited to small samples, histological sections, and monolayer cultures because the procedure requires the entry/permeability of a terminal transferase.
In contrast to Aniline blue, which diffuses and is dissolved by solvents, LysoTracker Red DND-99 is fixable, bright, and stable. Staining can be visualized with standard fluorescent or confocal microscopy in whole-mount or section using aqueous or solvent-based mounting media12,13. Here we describe protocols using this dye to look at PCD in normal and sonic hedgehog null mouse embryos. In addition, we demonstrate analysis of PCD in differentiating ES cell cultures and present a simple quantification method. In summary, LysoTracker staining can be a great complement to other methods of detecting PCD.

Watch this Scientific Journal Video about Use of LysoTracker to Detect Programmed Cell Death in Embryos and Differentiating Embryonic Stem Cells at JoVE.com

Use of LysoTracker to Detect Programmed Cell Death in Embryos and Differentiating Embryonic Stem Cells

We present a simple protocol to visualize regions of programmed cell death (PCD) in mouse embryos and differentiating embryonic stem (ES) cell cultures using a highly soluble dye called LysoTracker.

Programmed cell death (PCD) occurs in adults to maintain normal tissue homeostasis and during embryological development to shape tissues and ...

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23.0K Views.  University of Southern California. We present a simple protocol to visualize regions of programmed cell death (PCD) in mouse embryos and differentiating embryonic stem (ES) cell cultures using a highly soluble dye called LysoTracker.

Video: Use of LysoTracker to Detect Programmed Cell Death in Embryos and Differentiating Embryonic Stem Cells

Dissection of 6.5 dpc Mouse Embryos

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

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

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

Derivation of Hematopoietic Stem Cells from Murine Embryonic Stem Cells

A stem cell is defined as a cell with the capacity to both self-renew and generate multiple differentiated progeny.  Embryonic stem cells (ESC) are derived from the blastocyst of the early embryo and are pluripotent in differentiative ability.  Their vast differentiative potential has made them the focus of much research centered on deducing how to coax them to generate clinically useful cell types.  The successful derivation of hematopoietic stem cells (HSC) from mouse ESC has recently been accomplished and can be visualized in this video protocol.  HSC, arguably the most clinically exploited cell population, are used to treat a myriad of hematopoietic malignancies and disorders.  However, many patients that might benefit from HSC therapy lack access to suitable donors.  ESC could provide an alternative source of HSC for these patients.  The following protocol establishes a baseline from which ESC-HSC can be studied and inform efforts to isolate HSC from human ESC.  In this protocol, ESC are differentiated as embryoid bodies (EBs) for 6 days in commercially available serum pre-screened for optimal hematopoietic differentiation.  EBs are then dissociated and infected with retroviral HoxB4.  Infected EB-derived cells are plated on OP9 stroma, a bone marrow stromal cell line derived from the calvaria of  M-CSF-/- mice, and co-cultured in the presence of hematopoiesis promoting cytokines for ten days.  During this co-culture, the infected cells expand greatly, resulting in the generation a heterogeneous pool of 100s of millions of cells.  These cells can then be used to rescue and reconstitute lethally irradiated mice.

This protocol details the derivation of transplantable hematopoietic stem cells from mouse embryonic stem cells (ESC) and their subsequent injection into lethally irradiated recipient mice. Briefly, ESC are differentiated as embryoid bodies, which are then infected with retroviral HoxB4 and co-cultured with OP9 stromal cells and hematopoietic cytokines.

A stem cell is defined as a cell with the capacity to both self-renew and generate multiple differentiated progeny.  Embryonic stem cells (ESC) are ...

Derivation of Human Embryonic Stem Cells by Immunosurgery

The ability of human embryonic stem cells to self-renew and differentiate into all cell types of 
the body suggests that they hold great promise for both medical applications and as a research 
tool for addressing fundamental questions in development and disease. Here, we provide a 
concise, step-by-step protocol for the derivation of human embryonic stem cells from embryos 
by immunosurgical isolation of the inner cell mass. 


The ability of human embryonic stem cells to self-renew and differentiate into all cell types of 
the body suggests that they hold great promise for both medical applications and as a research 
tool for addressing fundamental questions in development and disease. Here, we provide a 
concise, step-by-step protocol for the derivation of human embryonic stem cells from embryos 
by immunosurgical isolation of the inner cell mass.

The ability of human embryonic stem cells to self-renew and differentiate into all cell types of 
the body suggests that they hold great promise for both ...

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 ...

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

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

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

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

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

Isolation and Derivation of Mouse Embryonic Germinal Cells

The ability of embryonic germinal cells (EG) to differentiate into primordial germinal cells (PGCs) and later into gametes during early developmental stages is a perfect model to address our hypothesis about cancer and infertility. This protocol shows how to isolate primordial germinal cells from developing gonads in 10.5-11.5 days post coitum (dpc) mouse embryos.  Developing gonadal ridges from mouse embryos (C57BL6J) were dissociated by mechanical disruption with collagenase, then plated in a mouse embryo fibroblast feeder layer (MEF-CF1) that was previously mitotically inactivated with mitomycin C in the presence of knockout media and supplemented with Leukemia Inhibitor Factor (LIF), basic Fibroblast Growth Factor (bFGF), and Stem Cell Factor (SCF).  Using these optimized methods for PCG identification, isolation, and establishment of culture conditions permits long term cultures of EG cells for more than 40 days.  The embryonic germinal cell lines showed embryonic phenotype and expression of common used markers of the pluripotent state.  Isolation and derivation of germinal cells in culture provide a tool to understand their development in vitro and offer the opportunity to monitor cumulative damage at genetic and epigenetic levels after exposure to oxidative stress.

The ability of embryonic germinal cells to differentiate into primordial germinal cells during early development stages is a perfect model to address our hypothesis about cancer and infertility. This protocol shows how to isolate primordial germinal cells from developing gonads in 10.5-11.5 days post coitum mouse embryos.

The ability of embryonic germinal cells (EG) to differentiate into primordial germinal cells (PGCs) and later into gametes during early developmental ...

Video Bioinformatics Analysis of Human Embryonic Stem Cell Colony Growth

Because video data are complex and are comprised of many images, mining information from video material is difficult to do without the aid of computer software.  Video bioinformatics is a powerful quantitative approach for extracting spatio-temporal data from video images using computer software to perform dating mining and analysis. In this article, we introduce a video bioinformatics method for quantifying the growth of human embryonic stem cells (hESC) by analyzing time-lapse videos collected in a Nikon BioStation CT incubator equipped with a camera for video imaging. In our experiments, hESC colonies that were attached to Matrigel were filmed for 48 hours in the BioStation CT.  To determine the rate of growth of these colonies, recipes were developed using CL-Quant software which enables users to extract various types of data from video images.  To accurately evaluate colony growth, three recipes were created. The first segmented the image into the colony and background, the second enhanced the image to define colonies throughout the video sequence accurately, and the third measured the number of pixels in the colony over time.  The three recipes were run in sequence on video data collected in a BioStation CT to analyze the rate of growth of individual hESC colonies over 48 hours. To verify the truthfulness of the CL-Quant recipes, the same data were analyzed manually using Adobe Photoshop software. When the data obtained using the CL-Quant recipes and Photoshop were compared, results were virtually identical, indicating the CL-Quant recipes were truthful. The method described here could be applied to any video data to measure growth rates of hESC or other cells that grow in colonies. In addition, other video bioinformatics recipes can be developed in the future for other cell processes such as migration, apoptosis, and cell adhesion.  

Video bioinformatics is the automated processing, analysis, understanding, and data mining of biological spatio-temporal data extracted from microscopic videos. The purpose of this article is to demonstrate a method for measuring human embryonic stem cell colony growth using a video bioinformatics method.

Because video data are complex and are comprised of many images, mining information from video material is difficult to do without the aid of computer ...

Differentiation of Embryonic Stem Cells into Oligodendrocyte Precursors

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant interest in deriving a pure population of lineage-committed oligodendrocyte precursor cells (OPCs) for transplantation. OPCs are characterized by the activity of the transcription factor Olig2 and surface expression of a proteoglycan NG2. Using the GFP-Olig2 (G-Olig2) mouse embryonic stem cell (mESC) reporter line, we optimized conditions for the differentiation of mESCs into GFP+Olig2+NG2+ OPCs. In our protocol, we first describe the generation of embryoid bodies (EBs) from mESCs. Second, we describe treatment of mESC-derived EBs with small molecules: (1) retinoic acid (RA) and (2) a sonic hedgehog (Shh) agonist purmorphamine (Pur) under defined culture conditions to direct EB differentiation into the oligodendroglial lineage. By this approach, OPCs can be obtained with high efficiency (&gt;80%) in a time period of 30 days. Cells derived from mESCs in this protocol are phenotypically similar to OPCs derived from primary tissue culture. The mESC-derived OPCs do not show the spiking property described for a subpopulation of brain OPCs in situ. To study this electrophysiological property, we describe the generation of spiking mESC-derived OPCs by ectopically expressing NaV1.2 subunit. The spiking and nonspiking cells obtained from this protocol will help advance functional studies on the two subpopulations of OPCs.

We describe a small molecule-based protocol for differentiation of mouse embryonic stem cells into oligodendrocyte precursor cells (OPCs). This protocol generates Olig2+NG2+ OPCs with high efficiency by 30 days of differentiation. We also describe a method to generate "spiking" OPCs that can fire action potentials.

Oligodendrocytes are the myelinating cells of the central nervous system. For regenerative cell therapy in demyelinating diseases, there is significant ...

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.

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 ...

Efficient Derivation of Human Cardiac Precursors and Cardiomyocytes from Pluripotent Human Embryonic Stem Cells with Small Molecule Induction

To date, the lack of a suitable human cardiac cell source has been the major setback in regenerating the human myocardium, either by cell-based transplantation or by cardiac tissue engineering1-3. Cardiomyocytes become terminally-differentiated soon after birth and lose their ability to proliferate. There is no evidence that stem/progenitor cells derived from other sources, such as the bone marrow or the cord blood, are able to give rise to the contractile heart muscle cells following transplantation into the heart1-3. The need to regenerate or repair the damaged heart muscle has not been met by adult stem cell therapy, either endogenous or via cell delivery1-3. The genetically stable human embryonic stem cells (hESCs) have unlimited expansion ability and unrestricted plasticity, proffering a pluripotent reservoir for in vitro derivation of large supplies of human somatic cells that are restricted to the lineage in need of repair and regeneration4,5. Due to the prevalence of cardiovascular disease worldwide and acute shortage of donor organs, there is intense interest in developing hESC-based therapies as an alternative approach. However, how to channel the wide differentiation potential of pluripotent hESCs efficiently and predictably to a desired phenotype has been a major challenge for both developmental study and clinical translation. Conventional approaches rely on multi-lineage inclination of pluripotent cells through spontaneous germ layer differentiation, resulting in inefficient and uncontrollable lineage-commitment that is often followed by phenotypic heterogeneity and instability, hence, a high risk of tumorigenicity6-8 (see a schematic in Fig. 1A). In addition, undefined foreign/animal biological supplements and/or feeders that have typically been used for the isolation, expansion, and differentiation of hESCs may make direct use of such cell-specialized grafts in patients problematic9-11. To overcome these obstacles, we have resolved the elements of a defined culture system necessary and sufficient for sustaining the epiblast pluripotence of hESCs, serving as a platform for de novo derivation of clinically-suitable hESCs and effectively directing such hESCs uniformly towards clinically-relevant lineages by small molecules12 (see a schematic in Fig. 1B). After screening a variety of small molecules and growth factors, we found that such defined conditions rendered nicotinamide (NAM) sufficient to induce the specification of cardiomesoderm direct from pluripotent hESCs that further progressed to cardioblasts that generated human beating cardiomyocytes with high efficiency (Fig. 2). We defined conditions for induction of cardioblasts direct from pluripotent hESCs without an intervening multi-lineage embryoid body stage, enabling well-controlled efficient derivation of a large supply of human cardiac cells across the spectrum of developmental stages for cell-based therapeutics.

We have established a protocol for induction of cardioblasts direct from pluripotent human embryonic stem cells maintained under defined conditions with small molecules, which enables derivation of a large supply of human cardiac progenitors and functional cardiomyocytes for cardiovascular repair.