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 μM 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 °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 °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 μl 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 μl per chamber).</li>
 <li>Incubate at 37 °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="Embryo analysis with Shh mutation; red fluorescence microscopy; LysoT, TUNEL, merged cell imaging." data-altupdatedat="1747911167000" data-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="LysoT and TUNEL fluorescence microscopy of cells under H2O2 treatment and serum starvation, colocalization." data-altupdatedat="1747911167000" data-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="LysoTracker staining of nuclei with H2O2 treatments; fluorescence microscopy and bar graph results." data-altupdatedat="1747911167000" data-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><tbody><tr><td>LysoTracker Red DND-99</td><td>Invitrogen</td><td>#L-7528</td><td></td></tr><tr><td>Hanks BSS</td><td>Invitrogen</td><td>14025-076</td><td></td></tr><tr><td>Paraformaldehyde</td><td>EMD</td><td>EM-PX0055-3</td><td></td></tr><tr><td>Vectashield</td><td>VECTOR</td><td>H-1200</td><td></td></tr><tr><td>DMEM</td><td>Cellgro</td><td>10-013-CV</td><td></td></tr><tr><td>Non-essential amino acids</td><td>Cellgro</td><td>25-025-CI</td><td></td></tr><tr><td>Sodium pyruvate</td><td>Cellgro</td><td>25-000-CI</td><td></td></tr><tr><td>FBS</td><td>Hyclone</td><td>SH30071.02</td><td></td></tr><tr><td>Pen-Strep</td><td>Invitrogen</td><td>15140-122</td><td></td></tr><tr><td>b-Mercapt&amp;#8211;thanol, 50 mM </td><td>Invitrogen</td><td>21985-023</td><td></td></tr><tr><td>LabTek-II Chamber slides &lt;br/&gt; (8-well)</td><td>Nalge Nunc International</td><td>154534</td><td></td></tr><tr><td>0.1% Gelatin</td><td>Millipore</td><td>ES-006-B</td><td></td></tr><tr><td>Dulbecco&#039;s PBS (D-PBS) </td><td>Cellgro</td><td>21-031-CV</td><td></td></tr><tr><td colspan="4">&lt;p class=&quot;jove_step&quot;&gt;Solution Recipes&lt;/p&gt; &lt;p class=&quot;jove_step&quot;&gt;4% Paraformaldehyde&lt;/p&gt; &lt;p class=&quot;jove_content1&quot;&gt;For 100 ml:&lt;/p&gt; &lt;ol&gt; &lt;li&gt;Mix 4 g paraformaldehyde, 90 ml H&lt;sub&gt;2&lt;/sub&gt;O, and NaOH (a drop of 2N NaOH). The paraformaldehyde will not go into solution until you have added some NaOH to increase the pH.&lt;/li&gt; &lt;li&gt;Stir and heat at 60 &amp;deg;C until all the powder is in solution (~10-20 min). &lt;strong&gt;Do not overheat.&lt;/strong&gt;&lt;/li&gt; &lt;li&gt;Add ~10 ml 10x PBS to achieve a final volume of 100 ml.&lt;/li&gt; &lt;li&gt;Store at -20 &amp;deg;C in convenient (~10 ml or ~40 ml) aliquots.&lt;/li&gt; &lt;/ol&gt; &lt;p class=&quot;jove_content1&quot;&gt;&lt;strong&gt;WARNING: Paraformaldehyde in &#039;frill&#039; 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.&lt;/strong&gt;&lt;/p&gt; &lt;p class=&quot;jove_step&quot;&gt;EB Culture Media&lt;/p&gt; &lt;p class=&quot;jove_content1&quot;&gt;For 500 ml EB Media:&lt;/p&gt; &lt;table border=&quot;1&quot;&gt; &lt;tbody&gt; &lt;tr&gt; &lt;td&gt;&lt;strong&gt;DMEM:&lt;/strong&gt;&lt;/td&gt; &lt;td&gt;&lt;strong&gt;404.5 ml&lt;/strong&gt;&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;FBS:&lt;/td&gt; &lt;td&gt;75.0 ml&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;L-Glutamine:&lt;/td&gt; &lt;td&gt;5.0 ml&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;Penicillin/Streptomycin:&lt;/td&gt; &lt;td&gt;5.0 ml&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;Non-essential amino acids:&lt;/td&gt; &lt;td&gt;5.0 ml&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;Sodium pyruvate:&lt;/td&gt; &lt;td&gt;5.0 ml&lt;/td&gt; &lt;/tr&gt; &lt;tr&gt; &lt;td&gt;&amp;#946;-Mercapt&amp;#8211;thanol:&lt;/td&gt; &lt;td&gt;500 &amp;mu;l&lt;/td&gt; &lt;/tr&gt; &lt;/tbody&gt; &lt;/table&gt;</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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Unit 1

Cell Death

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23.2K Views.  University of Southern California. Wij presenteren een eenvoudig protocol om gebieden van geprogrammeerde celdood (PCD) in muizenembryo's en onderscheidende embryonale stamcellen (ES) celkweken met een sterk oplosbare kleurstof genoemd LysoTracker visualiseren.

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

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

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

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

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

Developmental Biology

Labeling Stem Cells with Fluorescent Dyes for non-invasive Detection with Optical Imaging

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo labeling of stem cells with a fluorescent dye, subsequent intravenous injection of the labeled cells and visualization of their accumulation in specific target organs or pathologies. The presented technique demonstrates how we label human mesenchymal stem cells (hMSC) by simple incubation with the lipophilic fluorescent dye DiD (C67H103CIN2O3S) and how we label human embryonic stem cells (hESC) with the FDA approved fluorescent dye Indocyanine Green (ICG). The uptake mechanism is via adherence and diffusion of the lypophilic dye across the phospholipid cell membrane bilayer. The labeling efficiency is usually improved if the cells are incubated with the dye in serum-free media as opposed to incubation in serum-containing media. Furthermore, the addition of the transfection agent Protamine Sulfate significantly improves contrast agent uptake.

This video shows techniques for labeling of human embryonic stem cells and mesenchymal stem cells with fluorescent dyes. This technique can be used for an in vivo tracking of transplanted stem cells with optical imaging and for histopathological correlations with fluorescence microscopy.

Optical imaging (OI) is an easy, fast and inexpensive tool for in vivo monitoring of new stem cell based therapies. The technique is based on ex vivo ...

In vitro and in vivo Bioluminescence Reporter Gene Imaging of Human Embryonic Stem Cells

The discovery of human embryonic stem cells (hESCs) has dramatically increased the tools available to medical scientists interested in regenerative medicine. However, direct injection of hESCs, and cells differentiated from hESCs, into living organisms has thus far been hampered by significant cell death, teratoma formation, and host immune rejection. Understanding the in vivo hESC behavior after transplantation requires novel imaging techniques to longitudinally monitor hESC localization, proliferation, and viability. Molecular imaging has given investigators a high-throughput, inexpensive, and sensitive means for tracking in vivo cell proliferation over days, weeks, and even months. This advancement has significantly increased the understanding of the spatio-temporal kinetics of hESC engraftment, proliferation, and teratoma-formation in living subjects.

A major advance in molecular imaging has been the extension of noninvasive reporter gene assays from molecular and cellular biology into in vivo multi-modality imaging platforms. These reporter genes, under control of engineered promoters and enhancers that take advantage of the host cell s transcriptional machinery, are introduced into cells using a variety of vector and non-vector methods. Once in the cell, reporter genes can be transcribed either constitutively or only under specific biological or cellular conditions, depending on the type of promoter used. Transcription and translation of reporter genes into bioactive proteins is then detected with sensitive, noninvasive instrumentation (e.g., CCD cameras) using signal-generating probes such as D-luciferin.

To avoid the need for excitatory light to track stem cells in vivo as is required for fluorescence imaging, bioluminescence reporter gene imaging systems require only an exogenously administered probe to induce light emission. Firefly luciferase, derived from the firefly Photinus pyralis, encodes an enzyme that catalyzes D-luciferin to the optically active metabolite, oxyluciferin. Optical activity can then be monitored with an external CCD camera. Stably transduced cells that carry the reporter construct within their chromosomal DNA will pass the reporter construct DNA to daughter cells, allowing for longitudinal monitoring of hESC survival and proliferation in vivo. Furthermore, because expression of the reporter gene product is required for signal generation, only viable parent and daughter cells will create bioluminescence signal; apoptotic or dead cells will not. 

In this video, the specific materials and methods needed for tracking stem cell proliferation and teratoma formation with bioluminescence imaging will be described.

With the growing interest in stem cell therapies, molecular imaging techniques are ideal for monitoring stem cell behavior after transplantation. Luciferase reporter genes have enabled non-invasive, repetitive assessment of cell survival, location, and proliferation in vivo. This video will demonstrate how to track hESC proliferation in a living mouse.

The discovery of human embryonic stem cells (hESCs) has dramatically increased the tools available to medical scientists interested in regenerative ...

Identification Of Erythromyeloid Progenitors And Their Progeny In The Mouse Embryo By Flow Cytometry

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where they act as front line sentinels of infection and tissue damage. While other immune cells are continuously renewed from hematopoietic stem and progenitor cells (HSPC) located in the bone marrow, a lineage of macrophages, known as resident macrophages, have been shown to be self-maintained in tissues without input from bone marrow HSPCs. This lineage is exemplified by microglia in the brain, Kupffer cells in the liver and Langerhans cells in the epidermis among others. The intestinal and colon lamina propria are the only adult tissues devoid of HSPC-independent resident macrophages. Recent investigations have identified that resident macrophages originate from the extra-embryonic yolk sac hematopoiesis from progenitor(s) distinct from fetal hematopoietic stem cells (HSC). Among yolk sac definitive hematopoiesis, erythromyeloid progenitors (EMP) give rise both to erythroid and myeloid cells, in particular resident macrophages. EMP are only generated within the yolk sac between E8.5 and E10.5 days of development and they migrate to the fetal liver as early as circulation is connected, where they expand and differentiate until at least E16.5. Their progeny includes erythrocytes, macrophages, neutrophils and mast cells but only EMP-derived macrophages persist until adulthood in tissues. The transient nature of EMP emergence and the temporal overlap with HSC generation renders the analysis of these progenitors difficult. We have established a tamoxifen-inducible fate mapping protocol based on expression of the macrophage cytokine receptor Csf1r promoter to characterize EMP and EMP-derived cells in vivo by flow cytometry.

While infiltrating macrophages are continuously recruited to adult tissues from circulating precursors, resident macrophages seed their tissue during development, where they are maintained without further input from progenitors. The progenitors for resident macrophages were recently identified. Here, we present methods for the genetic fate mapping of the resident macrophage progenitors.

Macrophages are professional phagocytes from the innate arm of the immune system. In steady-state, sessile macrophages are found in adult tissues where ...

Tracing Gene Expression Through Detection of &#946;-galactosidase Activity in Whole Mouse Embryos

The Escherichia coli LacZ gene, encoding &#946;-galactosidase, is largely used as a reporter for gene expression and as a tracer in cell lineage studies. The classical histochemical reaction is based on the hydrolysis of the substrate X-gal in combination with ferric and ferrous ions, which produces an insoluble blue precipitate that is easy to visualize. Therefore, &#946;-galactosidase activity serves as a marker for the expression pattern of the gene of interest as the development proceeds. Here we describe the standard protocol for the detection of &#946;-galactosidase activity in early whole mouse embryos and the subsequent method for paraffin sectioning and counterstaining. Additionally, a procedure for clarifying whole embryos is provided to better visualize X-gal staining in deeper regions of the embryo. Consistent results are obtained by performing this procedure, although optimization of reaction conditions is needed to minimize background activity. Limitations in the assay should be also considered, particularly regarding the size of the embryo in whole mount staining. Our protocol provides a sensitive and a reliable method for &#946;-galactosidase detection during the mouse development that can be further applied to the cryostat sections as well as whole organs. Thus, the dynamic gene expression patterns throughout development can be easily analyzed by using this protocol in whole embryos, but also detailed expression at the cellular level can be assessed after paraffin sectioning.

Here we describe the standard protocol for the detection of &#946;-galactosidase activity in early whole mouse embryos and the method for paraffin sectioning and counterstaining. This is an easy and quick procedure to monitor gene expression during development that can also be applied to tissue sections, organs or cultured cells.

The Escherichia coli LacZ gene, encoding &#946;-galactosidase, is largely used as a reporter for gene expression and as a tracer in cell lineage studies. ...

Live Imaging of Mitosis in the Developing Mouse Embryonic Cortex

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the developing cerebral cortex, abnormal mitosis of neural progenitors can cause defects in brain size and function. Hence, there is a critical need for tools to understand the mechanisms of neural progenitor mitosis. Cortical development in rodents is an outstanding model for studying this process. Neural progenitor mitosis is commonly examined in fixed brain sections. This protocol will describe in detail an approach for live imaging of mitosis in ex vivo embryonic brain slices. We will describe the critical steps for this procedure, which include: brain extraction, brain embedding, vibratome sectioning of brain slices, staining and culturing of slices, and time-lapse imaging. We will then demonstrate and describe in detail how to perform post-acquisition analysis of mitosis. We include representative results from this assay using the vital dye Syto11, transgenic mice (histone H2B-EGFP and centrin-EGFP), and in utero electroporation (mCherry-&#945;-tubulin). We will discuss how this procedure can be best optimized and how it can be modified for study of genetic regulation of mitosis. Live imaging of mitosis in brain slices is a flexible approach to assess the impact of age, anatomy, and genetic perturbation in a controlled environment, and to generate a large amount of data with high temporal and spatial resolution. Hence this protocol will complement existing tools for analysis of neural progenitor mitosis.

Neural progenitor mitosis is a critical parameter of neurogenesis. Much of our understanding of neural progenitor mitosis is based on analysis of fixed tissue. Live imaging in embryonic brain slices is a versatile technique to assess mitosis with high temporal and spatial resolution in a controlled environment.

Although of short duration, mitosis is a complex and dynamic multi-step process fundamental for development of organs including the brain. In the ...

Neuroscience

Cell-Lineage Guided Mass Spectrometry Proteomics in the Developing (Frog) Embryo

Characterization of molecular events as cells give rise to tissues and organs raises a potential to better understand normal development and design efficient remedies for diseases. Technologies enabling accurate identification and quantification of diverse types and large numbers of proteins would provide still missing information on molecular mechanisms orchestrating tissue and organism development in space and time. Here, we present a mass spectrometry-based protocol that enables the measurement of thousands of proteins in identified cell lineages in Xenopus laevis (frog) embryos. The approach builds on reproducible cell-fate maps and established methods to identify, fluorescently label, track, and sample cells and their progeny (clones) from this model of vertebrate development. After collecting cellular contents using microsampling or isolating cells by dissection or fluorescence-activated cell sorting, proteins are extracted and processed for bottom-up proteomic analysis. Liquid chromatography and capillary electrophoresis are used to provide scalable separation for protein detection and quantification with high-resolution mass spectrometry (HRMS). Representative examples are provided for the proteomic characterization of neural-tissue fated cells. Cell-lineage-guided HRMS proteomics is adaptable to different tissues and organisms. It is sufficiently sensitive, specific, and quantitative to peer into the spatio-temporal dynamics of the proteome during vertebrate development.

Cell-Lineage Guided Mass Spectrometry Proteomics in the Developing Frog Embryo

Here we describe a mass spectrometry-based proteomic characterization of cell lineages with known tissue fates in the vertebrate Xenopus laevis embryo.

Characterization of molecular events as cells give rise to tissues and organs raises a potential to better understand normal development and design ...

Chemistry

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

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, ...

Cell Labeling and Injection in Developing Embryonic Mouse Hearts

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

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

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

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

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