We would like to acknowledge the financial support from the Monash Larkins Program as well as from a NHMRC CDF and a NHMRC project grant. Furthermore, we would like to thank Sue Mei Lim for her constructive suggestions, Edwina McGlinn for her support and the Monash Flowcore team (in particular Adam Dinsdale) for their help in producing the video.

1. Instrument Settings/Reagent Preparation/Genotyping
<ol>
	<li>Prepare iPS cell medium: Supplement 500 ml Knockout DMEM media with 75 ml fetal bovine serum (FBS), 5 ml L-glutamine, 5 ml Non-essential amino acids, 500 &#181;l &#946;-Mercaptoethanol, 5 x 105 units Leukaemia inhibitory factor (LIF). Please refer to Materials List for product and purchasing information for reagents used in the context of this manuscript.</li>
	<li>Prepare MEF medium: Supplement 500 ml DMEM media with 50 ml FBS, 5 ml L-glutamine, 5 ml Non-essential amino acids, 5 ml Sodium pyruvate, 500 &#181;l &#946;-Mercaptoethanol.</li>
	<li>Prepare freezing medium: For 10 ml, combine 9 ml of FBS with 1 ml of DMSO.</li>
	<li>Prepare Gelatin coated dishes Add 3-5 ml 0.1% porcine gelatin solution to a T75 flask, incubate at RT for at least 10 min and aspirate right before use.</li>
	<li>Prepare FACS labeling medium: For 10 ml, combine 9.9 ml of PBS with 0.1 ml of FBS.</li>
	<li>Perform a genotyping PCR for the Rosa26 m2rtTA locus: Perform reactions with Taq DNA polymerase according to manufacturer&#8217;s instructions. Use a modified MgCl2-concentration of 3.5 mM and the following 3 primers: oIMR8052: 5&#8217; GCG AAG AGT TTG TCC TCA ACC 3&#8217;, oIMR8545: 5&#8217;AAA GTC GCT CTG AGT TGT TAT 3&#8217;, oIMR8546: 5&#8217; GGA GCG GGA GAA ATG GAT ATG 3&#8217;. Cycling conditions: 94 &#176;C for 3 min; 35 cycles of (94 &#176;C for 30 sec, 65 &#176;C for 1 min, 72 &#176;C for 1 min); 72 &#176;C for 2 min; hold at 12 &#176;C. Expected product size: 650 bp for wild type allele and 340 bp for the mutant allele.</li>
	<li>Perform a genotyping PCR for the Collagen-StemCa/OKSM locus: Perform reactions with Taq DNA polymerase as per instructions. Use a modified MgCl2-concentration of 2.5 mM and the following 3 primers: col/frt-B: 5&#8217; CCCTCCATGTGTGACCAAGG 3&#8217;, col/frtA1: 5&#8217; GCACAGCATTGCGGACATGC 3&#8217;, col/frtC1: 5&#8217; GCAGAAGCGCGGCCGTCTGG 3&#8217;. Cycling conditions: 95 &#176;C for 1 min: 2 cycles of (94 &#176;C for 30 sec, 70 &#176;C for 45 sec), 6 cycles of (94 &#176;C for 30 sec, 68 &#176;C for 45 sec) and 30 cycles of (94 &#176;C for 20 sec, 60 &#176;C for 1 min); 72 &#176;C for 5 min; hold at 12 &#176;C. Expected product size: 331 bp for wild type allele and 551 bp for mutant allele.</li>
	<li>Carry out all centrifugation steps at 400 x g for 3 min at 4 &#176;C.</li>
</ol>
2. Generation of Mouse Embryonic Fibroblasts
<ol>
	<li>Perform a timed mating between an animal of the OKSM strain with a member of the opposite gender of the m2rtTA strain. Harvest embryos at embryonic day 13.5 by culling the mother and removing the uterine horn. Prior to removal of the uterine horn, spray the abdominal region with an 80% ethanol solution, to avoid microbial contamination.</li>
	<li>Transfer the uterine horn into a 10 cm tissue culture dish containing 10 ml of sterile Phosphate Buffered Saline (PBS). Use sterilized surgical grade scissors to cut the uterine horn into pieces containing one embryo (see Figure 3A,B).</li>
	<li>Use a dissection microscope (ideally placed within a tissue culture hood) and sterilized forceps to carefully remove the uterine envelope and the extra-embryonic membranes surrounding each embryo. Transfer each embryo to a separate 10 cm dish with 10 ml of PBS.</li>
	<li>Proceed by removing the embryo&#8217;s head, limbs, tail and internal organs (heart, liver, intestine etc.) with the forceps (see Figure 3C). Transfer the embryo&#8217;s head into a 1.5 ml tube and freeze down so it can be used for genotyping if necessary.</li>
	<li>Transfer the embryo&#8217;s torso into an empty 10cm plate and use two surgical blades to mince the embryo for 2 min. Add 200 &#181;l of trypsin/EDTA solution on top of the minced embryo and incubate for 3-5 min at RT. Continue mincing for an additional 2 min, add 2 ml of MEF media to inactivate trypsin, and then transfer into a 15 ml tube.</li>
	<li>Use a 1,000 &#181;l pipette to further mechanically dissociate the tissue by gentle pipetting. Transfer to a gelatin coated 10 cm dish and add an additional 10 ml of MEF media. Note: Both normoxic and hypoxic incubators can be used for culture, refer to discussion for more information.</li>
	<li>After 24-48 hr the plate should be densely covered with MEFs.</li>
	<li>Alternatively, the cells can then be further propagated or frozen down. For the freezing of cells, remove the culture media, rinse the dish with 10 ml PBS to remove traces of media and overlay with 3 ml Trypsin/EDTA solution. Incubate for 3-5 min at 37 &#176;C, inactivate trypsin digestion by adding 5 ml of MEF media and transfer to a 15 ml centrifugation tube. Spin down and resuspend the pellet in 3 ml freezing media. Transfer to 3 cryovials and freeze down in a cell freezing container.</li>
</ol>
3. Reprogramming of MEFs
NOTE: Pellet cells by centrifugation at 200 x g for 5 min at 4 &#176;C and use normoxic tissue culture incubators for reprogramming. It can be beneficial to derive and expand MEFs under hypoxic conditions (5% oxygen, see discussion for more information).
<ol>
	<li>Quickly thaw a low passage cryovial (P0-P1, 2-3 Mio cells) in a water bath (37 &#176;C) and transfer the contents into a tube with 10 ml of pre-warmed MEF media. Pellet cells, resuspend in 12 ml fresh MEF media, transfer into a gelatin coated T75 culture vessel, and allow cells to recover for 24-48 hr before proceeding. Note: MEFs can also be used directly after derivation.</li>
	<li>Following the removal of culture media, wash MEFs with PBS and cellularize by overlaying with a Trypsin/EDTA solution (3 ml for 3-5 min at 37 &#176;C). Quench trypsin by adding 5 ml of MEF media and further dissociate cells by gentle mixing with a 10 ml pipette. Determine cell numbers using a hemocytometer.</li>
	<li>For reprogramming experiments, seed MEFs onto gelatin coated T75 flasks at densities of 6.7 x 103 cells/cm2 (~0.5 Mio cells per flask) in iPSC media containing 2 &#181;g/ml doxycycline. Refer to Table 1 regarding seeding recommendations to obtain around 2 Mio reprogramming intermediates for each time point. Note: Reprogramming commences with the addition of doxycycline (2 &#956;g/ml) supplemented media</li>
	<li>For the first 6 days replace media every other day with fresh doxycycline supplemented iPSC media (12 ml of media per T75 flask). Beyond this point renew media on a daily basis if there are high cell numbers. Double the culture volume if media changes can only be performed every alternate day.</li>
	<li>Harvest reprogramming intermediates at the required time points (suggestion: Days 3, 6, 9, 12) by removing media, rinsing with appropriate volume of PBS (10 ml for a T75 flask) and overlaying with Trypsin-EDTA solution (3 ml for a T75 flask). Incubate for 3-5 min at 37 &#176;C and quench by adding 5 ml of iPSC media followed by gentle pipetting.</li>
	<li>Count cell suspension with a hemocytometer (see above), pellet by centrifugation, aspirate the supernatant and resuspend cells in 10 ml of PBS to remove any traces of trypsin. Pellet cells once again, aspirate supernatant and proceed with step 4.1 to isolate the reprogramming intermediates. Note: Cells undergoing reprogramming can be cryopreserved and thawed for FACS isolation at a later stage.</li>
	<li>To establish fully reprogrammed iPS cultures, grow cells in dox-free iPSC media for a further 4-7 days. Note: Successfully reprogramming cultures will contain a large number of colonies by day 12 (see Figure 4). During this time, aberrant iPS cells that still require OKSM expression will disappear.</li>
	<li>Alternatively, to propagate the remaining fully reprogrammed iPS cultures: initially, seed irradiated MEFs at a density of 15 x 103 cell/cm2 in 12 ml of iPSC media onto a gelatin coated T75 flask one day or 6 hr prior to experiments.</li>
	<li>Next, cellularize iPS cell cultures by removing media, rinsing the T75 flask with 10 ml of PBS, overlaying with 3 ml of Trypsin-EDTA solution and incubating for 3-5 min at 37 &#176;C. Quench trypsin by adding 5 ml of iPSC media followed by gentle mixing, transfer to a 15 ml conical tube, spin down and then resuspended in 10 ml iPS media.</li>
	<li>Transfer 500 &#181;l of this cell suspension onto the irradiated MEFs (T75 flask) and allow the cultures to grow for 4-5 days. Note: Established iPS cultures can be cryopreserved or used for experiments. Clonal cell lines can be derived by picking individual iPSC colonies under a dissecting microscope 22.</li>
	<li>Cryo preserve confluent iPSC cultures as described in 2.8 for MEFs.</li>
</ol>
4. Antibody Labeling
<ol>
	<li>Resuspend cell pellets of reprogramming cultures, as prepared in Section 3, in FACS labeling media supplemented with antibodies (anti-Thy-1.2 pacific blue 1:400, anti-Ssea-1 biotin 1:400 and anti-Epcam Fitc 1:400). Use 200 &#181;l of supplemented labeling mix per 5 million cells and incubate on ice.</li>
	<li>After 10 min gently tap the tube to resuspend the cells and incubate for an additional 10 min on ice. Add 10 ml of cold PBS per tube and spin down.</li>
	<li>For each tube to be stained prepare 200 &#181;l of labeling media supplemented with 1 &#181;l Streptavidin-PeCy7 (1:200) per 5 million cells. When cells have pelleted, carefully aspirate the supernatant and resuspend in an appropriate volume of labeling media supplemented with Streptavidin-PeCy7.</li>
	<li>Keep cells on ice for 10 min, tap to resuspend, incubate for another 10 min on ice, add 10 ml of cold PBS per tube, spin down and aspirate supernatant.</li>
	<li>Resuspend cell pellets in labeling media supplemented with propidium iodide (PI) (2 &#181;g/ml) at approximately 1 x 107 cells/ml. Remove cell clumps by passing the suspension through a 70 &#181;m strainer. Transfer cells to an appropriate FACS tube and keep them on ice.</li>
	<li>Prepare separate samples with individual antibodies (anti-Thy-1.2, anti-Ssea-1 and anti-Epcam). NOTE: These are required to perform color compensation in the three channels used in the analysis. In addition, unlabeled cells are needed to set voltages and gates for sorting. Ideally iPS cells are used for compensation: stocks of 0.5-1 x 106 iPS cells maintained on irradiated MEFs are stored in cryovials in liquid nitrogen. The presence of MEFs is essential to get a signal in the Thy-1.2 channel.</li>
	<li>Thaw a cryovial of iPS cells as described for MEFs (section 3). Following centrifugation, resuspend cells in 800 &#181;l of labeling buffer and 200 &#181;l of this sample is set aside as the unlabeled control.</li>
	<li>Use the remaining cells as compensation controls. Divide cells between three 15 ml tubes and add antibodies as follows: Pacific Blue compensation control: add 0.5 &#181;l of the anti-Thy-1.2 Pacific Blue antibody. Fitc compensation tube: add 0.5 &#181;l of the anti-Epcam-Fitc antibody. Pe-Cy7 compensation control: 0.5 &#181;l of the anti Ssea-1-Biotin antibody.</li>
	<li>Keep cell-antibody samples on ice for 10 min, mix by tapping and incubate for 10 min on ice.</li>
	<li>Wash samples with 10 ml cold PBS to remove unbound antibody, spin cells down and aspirate the supernatant. Resuspend cell pellets in 200 &#181;l of FACS labeling media.</li>
	<li>For anti-Ssea-1-Biotin labeled samples, add a Streptavidin-PeCy7 conjugate (1 &#181;l per 200 &#181;l cells). Label cells as described for primary antibody (Ssea-1-Biotin).</li>
	<li>Pass all samples including the unlabeled control through a 70 &#181;m strainer and add PI (2 &#181;g/ml). Transfer cells to FACS tubes and keep on ice until required.</li>
</ol>
5. FACS Isolation of Intermediates
NOTE: Cells are sorted using a Fluorescence Activated Cell Sorter with 405 nm, 488 nm and 560 nm excitation lasers and a 100 &#181;m nozzle.
<ol>
	<li>Set up voltages for forward and side scatter so that the cell population is properly visualized. Draw a gate around cells as indicated in Figure 5A to exclude debris. Note: The correct set up of voltages on the cell sorter can be complex and requires an experienced FACS operator.</li>
	<li>Use Forward scatter (FSC) height versus FSC area to exclude aggregates (Figure 5B).</li>
	<li>Visualize the PI channel versus FSC to gate in on the PI negative live cells (Figure 5C).</li>
	<li>Use the unlabeled cells to adjust the voltages for the fluorescent channels of PB, Fitc/GFP, Pe-Cy7. Ideally position the cell population at the lower end of the respective channel.</li>
	<li>Set gates with the unlabeled control cells as shown in Figure 6A,B to sub fraction the reprogramming cultures into day 3, 6 and 9 populations: Ssea-1-/Thy-1.2+ cells; Ssea-1-/Thy-1.2- cells; and the reprogramming intermediates Ssea-1+/Thy-1.2- cells.</li>
	<li>Further subdivide the Day 9 Ssea-1+/Thy-1.2- fraction using the Pe-Cy7 versus Fitc blot; draw gates around the Sse-a1+/Epcam+ cells and the Ssea-1+/Epcam- cells. Set gates with the unlabeled control cells as shown in Figure 6C,D.</li>
	<li>Prefill appropriate collection tubes with iPS cell media (1-2 ml) before sorting the desired subpopulations (see Figure 7 for representative FACS profiles for MEFs, Day 3/Day 6/Day 9/Day 12 cultures and iPS cells).</li>
	<li>Perform FACS sorting according to manufactures protocol.</li>
	<li>After FACS isolation submit cells to molecular profiling (RNA/ protein extraction, chromatin immunoprecipitation, DNA methylome analysis etc.). Alternatively, the various cell fractions may be cultured.</li>
</ol>

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The authors have nothing to disclose.

In order to successfully reprogram MEFs into iPS cells and to purify reprogramming intermediates at high quantity it is essential to be aware of factors that have an impact on the overall efficiency. In particular the batch of FBS used to supplement iPS media can have a detrimental effect. In general positive experiences were made with Embryonic stem (ES) cell qualified FBS but batch testing of sera from a variety of vendors might identify a cheaper alternative.
Another factor that impacts on reprogramming efficiency is the genotype of MEFs 15,20. Although MEFs derived from mice heterozygous (het) for both the OKSM and the m2rtTA locus do reprogram, homozygousity at one or both loci results in higher reprogramming efficiencies. Indeed, MEFs derived from the offspring of OKSM and m2rtTA crosses show an increase in reprogramming efficiency in the following order (OKSM/m2rtTA): het/het &#60; homo/het &#60; het/homo &#60; homo/homo (please note that the numbers given in Table 1 are for het/het animals). On the rare occasion a male homo/homo animal is identified, a harem mating with multiple m2rtTA females can be set up. All offspring from these crosses will be het/homo for the OKSM/m2rtTA loci, respectively. In addition, the rapid genotyping of litters is recommended as larger litters are obtained when using younger mice for breeding (6-15 weeks of age).
Moreover, the use of low passage MEFs for reprogramming is emphasized 23. If MEFs were derived and expanded in a normoxic tissue culture incubator we strongly recommend to only use these cells up to passage 2. However, if the experimenter has access to a low oxygen incubator (5% oxygen) for the derivation and expansion of MEFs, passage numbers as high as 3 will still yield good results 23.
In case the experimenter encounters problems associated with reprogramming, as outlined, the most likely explanations are high passage numbers at the time of dox addition, incorrect genotype of the mice or the use of FBS that is not conducive for iPS cell generation. However, in rare cases we observed that MEFs were not able to reprogram even under ideal conditions. In these cases a spontaneous de novo deletion within the m2rtTA&#8217;s open reading frame was identified as the underlying reason.
This methodology was successfully adapted to extract Ssea-1+ intermediates via magnetic cell isolation (MACS). There is a clear time advantage in using MACS, however, the Ssea-1+ cell populations&#8217; purity is reduced to 80-90%, rather than more than 95%, which depending on the type of experiment the cells are destined for might pose a problem. Thus, an option is to use MACS to enrich for Ssea-1+ cells followed by FACS to increase purity and/or fraction them into Ssea-1+/Epcam+ and Ssea-1+/Epcam- cells.
While the iPS cells produced by the outlined protocol and mouse model have been shown to be pluripotent and effectively contribute to all tissues of chimeric animals 15,20, a reduced capacity to produce embryos entirely composed of these iPS cells via tetraploid complementation has been demonstrated 24. Aberrant methylation patterns at the Dlk-Dio3 gene cluster have been identified as the underlying cause 24. However, addition of ascorbic acid at a concentration of 50 &#181;g/ml to the media during reprogramming will produce tetraploid complementation competent iPS cells 24. Please be advised that an alternative reprogrammable mouse model engineered to express the reprogramming factors at a different stoichiometry can produce tetraploid competent iPS cells even in the absence of ascorbic acid treatment 25. However, in our hands cells from this strain reprogram at considerably lower frequency compared to the herein used mouse model.
The methodology described in this manuscript allows the separation of reprogramming intermediates from the refractory bulk of the population and should be seen as valuable tool to dissect and understand the reprogramming process, removing the previous limitations of having to use an unfractioned population for profiling (e.g., high signal noise from the refractory cells). The antibody combination described herein allows isolation of intermediates from mouse cells at high purity, however it is possible that additional cell surface markers will be uncovered in the future that will allow to obtain intermediates at even higher purity. Please note that Ssea-1 expression is not a hallmark of human induced pluripotent stem cells and as such this protocol cannot be used on reprogramming cells of human origin. In conclusion, reprogramming intermediates once isolated with the described method can be used for molecular analyses including expression profiling, chromatin immunoprecipitation, methylome analysis, and protein assays.

Embryonic stem (ES) cells are derived from the inner cell mass of blastocyst stage embryos1. Under appropriate culture conditions they self-renew and remain pluripotent. In 2006 Yamanaka and colleagues demonstrated that mature cells can be reprogrammed into so called induced pluripotent stem (iPS) cells by forced expression of the transcription factors Oct-4, Klf-4, Sox-2, c-Myc (OKSM) 2. iPS cells, like ES cells, can give rise to all cell types of the body, however, they are free of the ethical constraints surrounding the generation of ES cells3. Furthermore, iPS cells carry the promise of personalized regenerative medicine and hold tremendous potential for applications like disease modeling and in vitro drug screening4,5. In order for reprogramming technology to fulfill this potential, the basic mechanism of nuclear reprogramming needs to be fully understood. However, efforts to dissect the reprogramming pathway have been hampered by the fact that only a very small number of cells reprogram (0.1-1%). Successfully reprogramming fibroblasts have been reported to undergo a distinct series of events including a mesenchymal to epithelial transition 6-10 and, in the final stages of reprogramming, activation of the endogenous core pluripotency network 11-14. We and others 12,13,15-17 have recently identified a set of cell surface markers that allows for the separation of rare intermediates from the refractory bulk population. Reprogramming mouse embryonic fibroblasts (MEFs) undergo changes in the expression of Thy-1.2, Ssea1 and Epcam (among others) during the 2-week-long reprogramming process15. Early during reprogramming a subset of MEFs down-regulate expression of fibroblast identity marker (Thy-1.2) and then start expressing the pluripotency-associated marker Ssea-112. During the final stages of reprogramming Ssea1-positive cells reactivate endogenous pluripotency genes such as Oct-410-13,15. This last transition is marked on the cell surface by detectable expression of Epcam (see Figure 1) or in a later stage Pecam 15. Recently, O&#8217;Malley et al. reported the use of CD44 and iCAM1 as alternatives or complementary to Thy-1.2 and Ssea-1 for the identification of reprogramming intermediates.&#160;We have previously FACS extracted reprogramming intermediates from Day 0, Day 3, Day 6, Day 9 and Day 12 reprogramming cultures, as well as from established iPS cell lines based on these cell surface markers 15,18. For the below described reprogramming system and conditions we have shown at the single cell level that although the populations are quiet homogenous, there is a certain degree of heterogeneity in the identified intermediate populations. It should be noted that only a subset of cells within these populations are able to progress to the respective next stage of the reprogramming process and give rise to iPS cell colonies at different efficiencies, which have been extensively characterized previously15,19. Moreover, the reprogramming efficiency of these populations will depend as well on the re-plating and culture conditions. To increase experimental reproducibility we use a reprogrammable mouse strain that has been engineered to express a transcriptional transactivator (m2rtTA) under control of the Rosa26 locus and a polycistronic OKSM cassette under control of a doxycycline responsive promoter20,21. Using this mouse model circumvents the unwanted side effects of traditional viral methods of iPS cell generation, i.e., a heterogeneous starting population with cell to cell variability in number and location of integration sites of viral inserts. Two transgenic mouse strains (OKSM, m2rtTA), available as homozygous founder animals at the Jackson Laboratory, have to be crossed in order to establish the reprogrammable mouse model (see Figure 2). In this manuscript we describe in detail how to derive MEFs, generate iPS cells, and isolate the reprogramming intermediates at various stages of the conversion process by FACS.

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cell surface marker mediated purification of ips cell intermediates from a reprogrammable mouse model

Mature cells can be reprogrammed to a pluripotent state. These so called induced pluripotent stem (iPS) cells are able to give rise to all cell types of the body and consequently have vast potential for regenerative medicine applications. Traditionally iPS cells are generated by viral introduction of transcription factors Oct-4, Klf-4, Sox-2, and c-Myc (OKSM) into fibroblasts. However, reprogramming is an inefficient process with only 0.1-1% of cells reverting towards a pluripotent state, making it difficult to study the reprogramming mechanism. A proven methodology that has allowed the study of the reprogramming process is to separate the rare intermediates of the reaction from the refractory bulk population. In the case of mouse embryonic fibroblasts (MEFs), we and others have previously shown that reprogramming cells undergo a distinct series of changes in the expression profile of cell surface markers which can be used for the separation of these cells. During the early stages of OKSM expression successfully reprogramming cells lose fibroblast identity marker Thy-1.2 and up-regulate pluripotency associated marker Ssea-1. The final transition of a subset of Ssea-1 positive cells towards the pluripotent state is marked by the expression of Epcam during the late stages of reprogramming. Here we provide a detailed description of the methodology used to isolate reprogramming intermediates from cultures of reprogramming MEFs. In order to increase experimental reproducibility we use a reprogrammable mouse strain that has been engineered to express a transcriptional transactivator (m2rtTA) under control of the Rosa26 locus and OKSM under control of a doxycycline responsive promoter. Cells isolated from these mice are isogenic and express OKSM homogenously upon addition of doxycycline. We describe in detail the establishment of the reprogrammable mice, the derivation of MEFs, and the subsequent isolation of intermediates during reprogramming into iPS cells via fluorescent activated cells sorting (FACS).

After the dissection, disaggregation and plating of an E13.5 mouse embryo, a 10 cm dish is expected to reach confluency in approximately 1 to 2 days. At this stage, it is normal for the culture to contain some adherent pieces of tissue that have not been cellularized properly. These will disappear after passaging.
Upon doxycycline induction, reprogramming MEFs undergo distinct morphological changes. Around day 6, early colony-like patches should start to emerge (Figure 4C). These will continue to grow in size upon further culture (see Figure 4D,E). A good reprogramming experiment should result in &#62;500 colonies per T75 that was initially seeded with 5 x 105 cells (Figure 4E). Established iPS cultures possess characteristic dome shaped colonies and should mostly be devoid of differentiated cells (Figure 4F). Occasionally, additional passaging of iPS cultures may be required to remove undifferentiated/partially reprogrammed cells; a confluent flask of cells can be split at a ratio of 1:10 onto a feeder layer of irradiated MEFs.
As mentioned above, morphological and molecular changes during reprogramming are reflected by changes in the FACS profiles for Thy-1.2, Ssea-1 and ultimately Epcam (see Figure 7A-F). As reported previously12,15, while MEFs are predominantly positive for Thy-1.2 and negative for the other markers, Day 3 cultures already look markedly different. A large proportion of cells have started to down-regulate the expression of Thy-1.2 and a very small subset of these Thy-1.2 negative cells have become positive for Ssea-1, the actual reprogramming intermediate for this time point. The number of reprogramming intermediates that can be extracted from Day 3 cultures is very unpredictable as the percentage of Ssea-1+/Thy-1.2- usually lies in a range between 1-10% of viable cells. On Days 6 and 9 an increased percentage of Ssea-1+ cells, usually well above 10%, can be detected. Around day 12 a subset of Ssea-1 positive cells can be detected that also label positive for Epcam. Day 12 cultures, like Day 3 cultures, represents a bottleneck in the purification of intermediates, as the percentage of Ssea-1.2+/Epcam+ cells is variable and usually falls in the range of only 2-4% of all live cells. The expected number of reprogramming intermediates presented in Table 1 only serves as a rough approximation. Established bona fide iPS cell cultures will be strongly positive for Ssea-1 and Epcam.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig1highres.jpg" width="500" /> 
Figure 1. Surface marker changes during the reprogramming pathway: Down-regulation of fibroblast identity marker Thy-1.2 is followed by up-regulation of Ssea-1. The transition of a subset of Ssea-1 positive cells towards a pluripotent state is indicated by acquisition of Epcam around day 12. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig2highres.jpg" width="500" /> 
Figure 2. Schematic representation of the reprogrammable mouse model: m2rtTA expression is under control of the ubiquitously active Rosa26 locus. In the presence of doxycycline (dox) the m2rtTA protein binds to a tetracycline dependent promoter (tetOP) at the Collagen 1a1 (Col1a1) locus resulting in the expression of the four-factor cassette. Two bicistronic cassettes are linked by an internal ribosome entry site (IRES). Open reading frames for Oct-4/Klf-4 and Sox-2/c-Myc are fused by self-cleaving F2A and E2A sequences, respectively. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig3highres.jpg" width="500" /> 
Figure 3. Embryo dissection: (A) Transfer uterine horn into a 10 cm dish filled with 10 ml PBS and (B) cut into pieces containing one embryo. (C) Using forceps free the embryos from extra embryonic tissue and remove head, limbs, tail and internal organs. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig4highres.jpg" width="500" /> 
Figure 4. Morphological changes during reprogramming. Colony-like patches become apparent in the cultures from day 6 onwards. Bona fide iPSCs are characterized by a domed shaped morphology. (A) Day 0/MEFs, (B) Day 3, (C) Day 6, (D) Day 9, (E) Day 12, (F) iPS cell culture. Scale bar: 200 &#181;m. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig5highres.jpg" width="500" /> 
Figure 5. Basic FACS setup. (A) Exclude debris with Side Scatter versus Forward Scatter Area blot. (B) Exclude aggregates from non-debris by gating on single cell population with Forward Scatter Area versus Forward Scatter High blot. (C) Exclude dead cells from single cell population by gating on PI low events with PI channel versus Forward Scatter Area blot. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig6highres.jpg" width="500" /> 
Figure 6. Gating. (A, B) Using the unlabeled control cells set up gates for Thy-1.2+/Ssea-1- cells, Thy-1.2-/Ssea-1- cells and Thy-1.2-/Ssea-1+ cells. (C) Use the unlabeled control cells to set gates for Epcam positive and Epcam negative cells. (D) Ssea-1+/Thy-1.2- cells can be separated into an Epcam positive and into an Epcam negative population. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" fo:content-width="5in" src="/files/ftp_upload/51728/51728fig7highres.jpg" width="500" /> 
Figure 7. Thy-1.2 versus Ssea-1 FACS blots at various stages of the reprogramming process. (A) Day 0/MEFs, (B) Day 3, (C) Day 6, (D) Day 9, (E) Day 12, (F) iPS cell culture.
Table 1. Suggested seeding density, flask number and expected outcome for P1 MEFs of an embryo heterozygous for OKSM and m2rtTA. <a href="https://www.jove.com/files/ftp_upload/51728/51728fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Day </td>
			<td>Number of T75 flasks seeded on day 0</td>
			<td>Total number of Ssea1+ cells after FACS </td>
			<td>Total number of Ssea1+/Epcam+ cells after FACS </td>
		</tr>
		<tr>
			<td>3</td>
			<td>8</td>
			<td>2 million </td>
			<td></td>
		</tr>
		<tr>
			<td>6</td>
			<td>4</td>
			<td>2 million </td>
			<td></td>
		</tr>
		<tr>
			<td>9</td>
			<td>4</td>
			<td>2 million </td>
			<td></td>
		</tr>
		<tr>
			<td>12</td>
			<td>10</td>
			<td>10 million </td>
			<td>2 million </td>
		</tr>
	</tbody>
</table>

Watch this Scientific Journal Video about Cell Surface Marker Mediated Purification of iPS Cell Intermediates from a Reprogrammable Mouse Model at JoVE.com

Cell Surface Marker Mediated Purification of iPS Cell Intermediates from a Reprogrammable Mouse Model

Mouse embryonic fibroblast can be reprogrammed into induced pluripotent stem cells at low efficiency by the forced expression of transcription factors Oct-4, Sox-2, Klf-4, c-Myc. The rare intermediates of the reprogramming reaction are FACS isolated via labeling with antibodies against cell surface makers Thy-1.2, Ssea-1, and Epcam.

Mature cells can be reprogrammed to a pluripotent state. These so called induced pluripotent stem (iPS) cells are able to give rise to all cell types of ...

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12.2K ビュー.  Monash University. Mouse embryonic fibroblast can be reprogrammed into induced pluripotent stem cells at low efficiency by the forced expression of transcription factors Oct-4, Sox-2, Klf-4, c-Myc. The rare intermediates of the reprogramming reaction are FACS isolated via labeling with antibodies against cell surface makers Thy-1.2, Ssea-1, and Epcam. 

ビデオ: Cell Surface Marker Mediated Purification of iPS Cell Intermediates from a Reprogrammable Mouse Model

Mouse embryonic fibroblast can be reprogrammed into induced pluripotent stem cells at low efficiency by the forced expression of transcription factors Oct-4, Sox-2, Klf-4, c-Myc. The rare intermediates of the reprogramming reaction are FACS isolated via labeling with antibodies against cell surface makers Thy-1.2, Ssea-1, and Epcam.

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We present a fluorescence assay to demonstrate that Lucifer Yellow (LY) is a robust marker to determine the apparent paracellular permeability of hCMEC/D3 cell monolayers, an in vitro model of the human blood-brain barrier. We used this assay to determine the kinetics of a confluent monolayer formation in cultured hCMEC/D3 cells.

fluorescence-activated cell sorting for purification of plasmacytoid dendritic cells from the mouse bone marrow

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of...

high-quality brain and bone marrow nuclei preparation for single nuclei multiome assays

Author Spotlight: Advancing Biomedical Research Through Single Cell Analysis 

The success of single-cell/single-nuclei transcriptomics and multi-omics largely depends on the quality of cells/nuclei. Therefore, isolating cells/nuclei from tissue and their purification must be highly standardized. This protocol describes the preparation of nuclei from the brain and bone marrow for downstream single-nuclei multiome assay.

High-Quality Sample Preparation for Multiome 10X Genomics Analysis

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trophoblast cell recovery from angiogenesis-tube formation assay for differentiation marker expression analysis

Author Spotlight: Exploring Trophoblast Invasion and Differentiation for Placental Development and Preeclampsia

The present protocol allows the recovery of tube-like structures formed in the classical in vitro angiogenesis-tube formation assay where cells, such as trophoblast cells, organize in capillary-like structures over a layer of extracellular basement membrane matrix. The recovered structures are used for RNA and protein isolation for various biochemical...

Trophoblast Cell Recovery from Angiogenesis-Tube Formation Assay for Differentiation Marker Expression Analysis

"cell surface capture" workflow for label-free quantification of the cell surface proteome

&quot;Cell Surface Capture&quot; Workflow for Label-Free Quantification of the Cell Surface Proteome

Here, we describe a proteomics workflow for characterization of the cell surface proteome of various cell types. This workflow includes cell surface protein enrichment, subsequent sample preparation, analysis using an LC-MS/MS platform, and data processing with specialized...

"Cell Surface Capture" Workflow for Label-Free Quantification of the Cell Surface Proteome

purification of platelets from mouse blood

Purification of Platelets from Mouse Blood

Here, mouse blood was collected in the presence of an anti-coagulant. The platelets were purified by iohexol gradient medium using low speed centrifugation. The platelets were activated with thrombin to investigate if they were viable. The quality of the purified platelets was analyzed by flow cytometry and...

cd8+ t-cell purification from mouse lymph nodes and spleen for kinetic studies

CD8+ T-Cell Purification from Mouse Lymph Nodes and Spleen for Kinetic Studies

visualization and quantification of mesenchymal cell adipogenic differentiation potential with a lineage specific marker

Visualization and Quantification of Mesenchymal Cell Adipogenic Differentiation Potential with a Lineage Specific Marker

Traditional methods of assessing adipogenic differentiation are cheap and easy to use, but are not specific to changes in gene expression. We have developed an assay to quantify mesenchymal cell differentiation into mature adipocytes using a lineage specific marker. This assay has diverse applications across basic research and clinical...

dissection of adult mouse utricle and adenovirus-mediated supporting-cell infection

Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Mechanosensory hair cells are the receptor cells of the inner ear.  The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice.  We present the dissection of the adult mouse utricle, and we demonstrate adenovirus-mediated infection of supporting cells in cultured...

cell-type specific protein purification and identification from complex tissues using a mutant methionine trna synthetase mouse line

Cell-Type Specific Protein Purification and Identification from Complex Tissues Using a Mutant Methionine tRNA Synthetase Mouse Line

This protocol describes how to perform cell-type-specific protein labeling with azidonorleucine (ANL) using a mouse line expressing a mutant L274G-Methionine tRNA synthetase (MetRS*) and the necessary steps for labeled cell-type-specific proteins isolation. We outline two possible ANL administration routes in live mice by (1) drinking water and (2) intraperitoneal injections.

isolation and enrichment of liver progenitor subsets identified by a novel surface marker combination

Isolation and Enrichment of Liver Progenitor Subsets Identified by a Novel Surface Marker Combination

Liver injuries are accompanied by progenitor cell expansion that represents a heterogeneous cell population. Novel classification of this cellular compartment allows for the distinguishing of multiple subsets. The method described here illustrates the flow cytometry analysis and high purity isolation of various subsets that can be used for further...

preparation of a single-cell suspension from mouse carotid arteries for single-cell sequencing

Author Spotlight: Vascular Tissue Dissociation and Exploring Single-Cell Subclusters for Targeted Therapy 

Here we describe a two-step cell digestion protocol for preparing a single-cell suspension of mouse carotid...

Preparation of a Single-Cell Suspension from Mouse Carotid Arteries for Single-Cell Sequencing

a method of targeted cell isolation via glass surface functionalization

A Method of Targeted Cell Isolation via Glass Surface Functionalization

This protocol describes customizable surface functionalization of the desthiobiotin, streptavidin, and APTES system in order to isolate specific cell types of interest. In addition, this manuscript covers the applications, optimization, and verification of this process.

purification of specific cell population by fluorescence activated cell sorting (facs)

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting FACS

For many scientific studies requiring a biological and chemical analysis of cell populations the cells must be in a high state of purity.  Fluorescence activated cell sorting (FACS) is a superior method in which to obtain pure cell populations.

Purification of Specific Cell Population by Fluorescence Activated Cell Sorting (FACS)

localized surface plasmon resonance imaging to detect protein secretions from a single cell

This video demonstrates localized surface plasmon resonance imaging to detect protein secretions from a single cell. Using a microfluidic setup and a peptide-bound plasmonic nanostructure chip, changes in reflected light intensity at the resonance angle reveal interactions with secreted proteins from a single...

Localized Surface Plasmon Resonance Imaging to Detect Protein Secretions from a Single Cell

Chromatographic Purification of Highly Active Yeast Ribosomes

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers. Particularly troublesome is the fact that lysis of cells releases a large number of proteases and nucleases which can degrade ribosomes. Thus, it is important to separate ribosomes from these enzymes as quickly as possible. Unfortunately, conventional differential ultracentrifugation methods leaves ribosomes exposed to these enzymes for unacceptably long periods of time, impacting their structural integrity and functionality. To address this problem, we utilize a chromatographic method using a cysteine charged Sulfolink resin. This simple and rapid application significantly reduces co-purifying proteolytic and nucleolytic activities, producing high yields of intact, highly biochemically active yeast ribosomes. We suggest that this method should also be applicable to mammalian ribosomes. The simplicity of the method, and the enhanced purity and activity of chromatographically purified ribosome represents a significant technical advancement for the study of eukaryotic ribosomes.

Contamination of preparations of eukaryotic ribosomes purified by traditional methods by co-purifying nucleases and proteases negatively impacts on downstream biochemical and structural analyses. A rapid and simple chromatographic purification method is used to solve this problem using yeast ribosomes as a model system.

Eukaryotic ribosomes are much more labile as compared to their eubacterial and archael counterparts, thus posing a significant challenge to researchers.  ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Avidity-based Extracellular Interaction Screening (AVEXIS) for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ensuring cohesion within multicellular organisms. Proteins predicted to form extracellular interactions are encoded by approximately a quarter of human genes1, but despite their importance and abundance, the majority of these proteins have no documented binding partner. Primarily, this is due to their biochemical intractability: membrane-embedded proteins are difficult to solubilise in their native conformation and contain structurally-important posttranslational modifications. Also, the interaction affinities between receptor proteins are often characterised by extremely low interaction strengths (half-lives &lt; 1 second) precluding their detection with many commonly-used high throughput methods2. 
Here, we describe an assay, AVEXIS (AVidity-based EXtracellular Interaction Screen) that overcomes these technical challenges enabling the detection of very weak protein interactions (t1/2 &le; 0.1 sec) with a low false positive rate3. The assay is usually implemented in a high throughput format to enable the systematic screening of many thousands of interactions in a convenient microtitre plate format (Fig. 1). It relies on the production of soluble recombinant protein libraries that contain the ectodomain fragments of cell surface receptors or secreted proteins within which to screen for interactions; therefore, this approach is suitable for type I, type II, GPI-linked cell surface receptors and secreted proteins but not for multipass membrane proteins such as ion channels or transporters.
The recombinant protein libraries are produced using a convenient and high-level mammalian expression system4, to ensure that important posttranslational modifications such as glycosylation and disulphide bonds are added. Expressed recombinant proteins are secreted into the medium and produced in two forms: a biotinylated bait which can be captured on a streptavidin-coated solid phase suitable for screening, and a pentamerised enzyme-tagged (&beta;-lactamase) prey. The bait and prey proteins are presented to each other in a binary fashion to detect direct interactions between them, similar to a conventional ELISA (Fig. 1). The pentamerisation of the proteins in the prey is achieved through a peptide sequence from the cartilage oligomeric matrix protein (COMP) and increases the local concentration of the ectodomains thereby providing significant avidity gains to enable even very transient interactions to be detected. By normalising the activities of both the bait and prey to predetermined levels prior to screening, we have shown that interactions having monomeric half-lives of 0.1 sec can be detected with low false positive rates3.

Avidity-based Extracellular Interaction Screening AVEXIS for the Scalable Detection of Low-affinity Extracellular Receptor-Ligand Interactions

AVEXIS is a high throughput protein interaction assay developed to systematically screen for novel extracellular receptor-ligand pairs involved in cellular recognition processes. It is specifically designed to detect transient protein interactions that are difficult to identify using other high throughput approaches.

Extracellular protein:protein interactions between secreted or membrane-tethered proteins are critical for both initiating intercellular communication and ...

Generation of Mice Derived from Induced Pluripotent Stem Cells

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also produce a source of patient-matched cells for regenerative therapies. iPSCs may be generated using multiple protocols and derived from multiple cell sources. Once generated, iPSCs are tested using a variety of assays including immunostaining for pluripotency markers, generation of three germ layers in embryoid bodies and teratomas, comparisons of gene expression with embryonic stem cells (ESCs) and production of chimeric mice with or without germline contribution2. Importantly, iPSC lines that pass these tests still vary in their capacity to produce different differentiated cell types2. This has made it difficult to establish which iPSC derivation protocols, donor cell sources or selection methods are most useful for different applications.
The most stringent test of whether a stem cell line has sufficient developmental potential to generate all tissues required for survival of an organism (termed full pluripotency) is tetraploid embryo complementation (TEC)3-5. Technically, TEC involves electrofusion of two-cell embryos to generate tetraploid (4n) one-cell embryos that can be cultured in vitro to the blastocyst stage6. Diploid (2n) pluripotent stem cells (e.g. ESCs or iPSCs) are then injected into the blastocoel cavity of the tetraploid blastocyst and transferred to a recipient female for gestation (see Figure 1). The tetraploid component of the complemented embryo contributes almost exclusively to the extraembryonic tissues (placenta, yolk sac), whereas the diploid cells constitute the embryo proper, resulting in a fetus derived entirely from the injected stem cell line.
Recently, we reported the derivation of iPSC lines that reproducibly generate adult mice via TEC1. These iPSC lines give rise to viable pups with efficiencies of 5-13%, which is comparable to ESCs3,4,7 and higher than that reported for most other iPSC lines8-12. These reports show that direct reprogramming can produce fully pluripotent iPSCs that match ESCs in their developmental potential and efficiency of generating pups in TEC tests. At present, it is not clear what distinguishes between fully pluripotent iPSCs and less potent lines13-15. Nor is it clear which reprogramming methods will produce these lines with the highest efficiency. Here we describe one method that produces fully pluripotent iPSCs and "all- iPSC" mice, which may be helpful for investigators wishing to compare the pluripotency of iPSC lines or establish the equivalence of different reprogramming methods.

Generating induced pluripotent stem cell (iPSC) lines produces lines of differing developmental potential even when they pass standard tests for pluripotency. Here we describe a protocol to produce mice derived entirely from iPSCs, which defines the iPSC lines as possessing full pluripotency1.

The production of induced pluripotent stem cells (iPSCs) from somatic cells provides a means to create valuable tools for basic research and may also ...

MicroRNA Expression Profiles of Human iPS Cells, Retinal Pigment Epithelium Derived From iPS, and Fetal Retinal Pigment Epithelium

The objective of this report is to describe the protocols for comparing the microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, retinal pigment epithelium (RPE) derived from human iPS cells (iPS-RPE), and fetal RPE. The protocols include collection of RNA for analysis by microarray, and the analysis of microarray data to identify miRNAs that are differentially expressed among three cell types. The methods for culture of iPS cells and fetal RPE are explained. The protocol used for differentiation of RPE from human iPS is also described. The RNA extraction technique we describe was selected to allow maximal recovery of very small RNA for use in a miRNA microarray. Finally, cellular pathway and network analysis of microarray data is explained. These techniques will facilitate the comparison of the miRNA profiles of three different cell types.

The microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, retinal pigment epithelium (RPE) derived from human induced-pluripotent stem (iPS) cells (iPS-RPE), and fetal RPE, were compared.

The objective of this report is to describe the protocols for comparing the microRNA (miRNA) profiles of human induced-pluripotent stem (iPS) cells, ...

This manuscript illustrates a protocol for efficiently creating integration-free human induced pluripotent stem cells (iPSCs) from peripheral blood using episomal plasmids and histone deacetylase (HDAC) inhibitors. The advantages of this approach include: (1) the use of a minimal amount of peripheral blood as a source material; (2) nonintegrating reprogramming vectors; (3) a cost effective method for generating vector free iPSCs; (4) a single transfection; and (5) the use of small molecules to facilitate epigenetic reprogramming. Briefly, peripheral blood mononuclear cells (PBMCs) are isolated from routine phlebotomy samples and then cultured in defined growth factors to yield a highly proliferative erythrocyte progenitor cell population that is remarkably amenable to reprogramming. Nonintegrating, nontransmissible episomal plasmids expressing OCT4, SOX2, KLF4, MYCL, LIN28A, and a p53 short hairpin (sh)RNA are introduced into the derived erythroblasts via a single nucleofection. Cotransfection of an episome that expresses enhanced green fluorescent protein (eGFP) allows for easy identification of transfected cells. A separate replication-deficient plasmid expressing Epstein-Barr nuclear antigen 1 (EBNA1) is also added to the reaction mixture for increased expression of episomal proteins. Transfected cells are then plated onto a layer of irradiated mouse embryonic fibroblasts (iMEFs) for continued reprogramming. As soon as iPSC-like colonies appear at about twelve days after nucleofection, HDAC inhibitors are added to the medium to facilitate epigenetic remodeling. We have found that the inclusion of HDAC inhibitors routinely increases the generation of fully reprogrammed iPSC colonies by 2 fold. Once iPSC colonies exhibit typical human embryonic stem cell (hESC) morphology, they are gently transferred to individual iMEF-coated tissue culture plates for continued growth and expansion.

Here we describe a protocol for generating human induced pluripotent stem cells from peripheral blood using an episome based reprogramming strategy and histone deacetylase inhibitors.

This manuscript illustrates a protocol for efficiently creating integration-free human induced pluripotent stem cells (iPSCs) from peripheral blood using ...

Trans-inner Cell Mass Injection of Embryonic Stem Cells Leads to Higher Chimerism Rates

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current blastocyst injection protocol. Here we report that a simple rotation of the embryo, and injection through Trans-Inner cell mass (TICM) increased the percentage of chimeric mice from 31% to 50%, with no additional equipment or further specialized training. 26 different inbred clones, and 35 total clones were injected over a period of 9 months. There was no significant difference in either pregnancy rate or recovery rate of embryos between traditional injection techniques and TICM. Therefore, without any major alteration in the injection process and a simple positioning of the blastocyst and injecting through the ICM, releasing the ES cells into the blastocoel cavity can potentially improve the quantity of chimeric production and subsequent germline transmission.

Here we present a protocol to increase chimera production without the use of new equipment. A simple orientation change of the embryo for injection can increase the number of embryos produced, and potentially reduce the timeline to germline transmission.

In an effort to increase efficiency in the creation of genetically modified mice via ES Cell methodologies, we present an adaptation to the current ...

Primary Cell Cultures from the Mouse Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the eye. It is a single sheet of pigmented cells that are hexagonally packed and connected by tight junctions. The main functions of the RPE include absorption of light, phagocytosis of the shed photoreceptor outer segments, spatial buffering of ions, transport of nutrients, ions and water as well as active involvement in the visual cycle. With such important and diverse functions, it is critically important to study the biology of RPE cells. A number of RPE cell lines have been established; however, passaged and immortalized cells are known to quickly lose some of the morphological and physiological characteristics of natural RPE cells. Thus, primary cells are more suitable for studying different aspects of RPE cell biology and function. Mouse primary RPE cell culture is very useful to researchers since mouse models are widely used in biological studies, however collecting RPE cells from mouse is also very challenging due to their small size. Here, we present a protocol for establishing primary mouse RPE cell cultures which includes enucleation and dissection of the eyes and isolation of the RPE sheets to yield the cells for culturing. This method enables efficient cell recovery. The RPE cells obtained from two mice&#160;can reach confluency on one 12 mm polyester membrane insert pre-loaded in culture plate after one week of culture and display some of the original properties of bona fide RPE cells such as hexagonal shape and pigmentation after two weeks of culture.

The retinal pigment epithelium (RPE) is a multi-functional epithelium of the eye. Here we present a protocol to establish primary cell cultures derived from the murine RPE.

The retinal pigment epithelium (RPE) is a highly polarized multi-functional epithelium that is located between the neural retina and the choroid of the ...

Several dyes are currently available for use in detecting differentiation of mesenchymal cells into adipocytes. Dyes, such as Oil Red O, are cheap, easy to use and widely utilized by laboratories analyzing the adipogenic potential of mesenchymal cells. However, they are not specific to changes in gene transcription. We have developed a gene-specific differentiation assay to analyze when a mesenchymal cell has switched its fate to an adipogenic lineage. Immuno-labelling against fatty acid binding protein-4 (FABP4), a lineage-specific marker of adipogenic differentiation, enabled visualization and quantification of differentiated cells. The ability to quantify adipogenic differentiation potential of mesenchymal cells in a 96 well microplate format has promising implications for a number of applications. Hundreds of clinical trials involve the use of adult mesenchymal stromal cells and it is currently difficult to correlate therapeutic outcomes within and especially between such clinical trials. This simple high-throughput FABP4 assay provides a quantitative assay for assessing the differentiation potential of patient-derived cells and is a robust tool for comparing different isolation and expansion methods. This is particularly important given the increasing recognition of the heterogeneity of the cells being administered to patients in mesenchymal cell products. The assay also has potential utility in high throughput drug screening, particularly in obesity and pre-diabetes research.

Traditional methods of assessing adipogenic differentiation are cheap and easy to use, but are not specific to changes in gene expression. We have developed an assay to quantify mesenchymal cell differentiation into mature adipocytes using a lineage specific marker. This assay has diverse applications across basic research and clinical medicine.

Several dyes are currently available for use in detecting differentiation of mesenchymal cells into adipocytes. Dyes, such as Oil Red O, are cheap, easy ...

A Mouse Model of Intestinal Partial Obstruction

Intestinal obstructions, that impede or block peristaltic movement, can be caused by abdominal adhesions and most gastrointestinal (GI) diseases including tumorous growths. However, the cellular remodeling mechanisms involved in, and caused by, intestinal obstructions are poorly understood. Several animal models of intestinal obstructions have been developed, but the mouse model is the most cost/time effective. The mouse model uses the surgical implantation of an intestinal partial obstruction (PO) that has a high mortality rate if it is not performed correctly. In addition, mice receiving PO surgery fail to develop hypertrophy if an appropriate blockade is not used or not properly placed. Here, we describe a detailed protocol for PO surgery which produces reliable and reproducible intestinal obstructions with a very low mortality rate. This protocol utilizes a surgically placed silicone ring that surrounds the ileum which partially blocks digestive movement in the small intestine. The partial blockage makes the intestine become dilated due to the halt of digestive movement. The dilation of the intestine induces smooth muscle hypertrophy on the oral side of the ring that progressively develops over 2 weeks until it causes death. The surgical PO mouse model offers an in vivo model of hypertrophic intestinal tissue useful for studying pathological changes of intestinal cells including smooth muscle cells (SMC), interstitial cells of Cajal (ICC), PDGFR&#945;+, and neuronal cells during the development of intestinal obstruction.

Intestinal obstructions are a partial or complete blockage of the intestine that can cause severe abdominal pain, nausea, vomiting, and preventing the passage of stool. This procedure for creating intestinal partial obsructions in mice is reliable in studying the mechanisms underlying pathological cell growth and death in the intestine.

Intestinal obstructions, that impede or block peristaltic movement, can be caused by abdominal adhesions and most gastrointestinal (GI) diseases including ...