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PubMed articles by:
Lengner, C.
Camargo, F.
Hochedlinger, K.
Jaenisch, R.
Cell Stem Cell.Author manuscript; available in PMC 2008 October 11.
Published in final edited form as:
PMCID: PMC2151746
NIHMSID: NIHMS32596
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Oct4 expression is not required for mouse somatic stem cell self-renewal
Christopher J Lengner,1 Fernando D Camargo,1 Konrad Hochedlinger,3 G Grant Welstead,1 Samir Zaidi,2 Sumita Gokhale,1 Hans R Scholer,4 Alexey Tomilin,5 and Rudolf Jaenisch1,2
1 Whitehead Institute for Biomedical Research, Massachusetts Institute of Technology, Nine Cambridge Center, Cambridge, MA 02142
2 Department of Biology, Massachusetts Institute of Technology, Nine Cambridge Center, Cambridge, MA 02142
3 Massachusetts General Hospital Cancer Center and Center for Regenerative Medicine, Harvard Stem Cell Institute, 185 Cambridge Street, Boston, Massachusetts 02114, USA.
4 Department of Cell and Developmental Biology, Max Planck Institute for Molecular Biomedicine, Munster, Germany
5 Institute of Cytology, Russian Academy of Science, 4 Tikhoretski Ave. 194064 St-Petersburg, Russia
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>Abstract
Abstract
The Pou domain containing transcription factor Oct4 is a well-established regulator of pluripotency in the inner cell mass of the mammalian blastocyst as well as in embryonic stem cells. While it has been shown that the Oct4 gene is inactivated through a series of epigenetic modifications following implantation, recent studies have detected Oct4 activity in a variety of somatic stem cells and tumor cells. Based on these observations it has been suggested that Oct4 may also function in maintaining self-renewal of somatic stem cells and, in addition, may promote tumor formation. We employed a genetic approach to determine whether Oct4 is important for maintaining pluripotency in the stem cell compartments of several somatic tissues including the intestinal epithelium, bone marrow (hematopoietic and mesenchymal lineages), hair follicle, brain, and liver. Oct4 gene ablation in these tissues revealed no abnormalities in homeostasis or regenerative capacity. We conclude that Oct4 is dispensable for both self-renewal and maintenance of somatic stem cells in the adult mammal.
>Introduction
Introduction
The transcriptional regulator Oct4 is a Pou domain containing protein expressed in pluripotent embryonic cells and cells of the germline where its inactivation results in loss of pluripotency and apoptosis, respectively (Kehler et al., 2004; Nichols et al., 1998). In embryonic stem (ES) cells Oct4, along with transcriptional co-regulators Nanog and Sox-2, orchestrates a program of gene activity that suppresses differentiation while endowing self-renewal (Boyer et al., 2005; Loh et al., 2006; Wang et al., 2006). Upon induction of differentiation, the Oct4 locus undergoes a series of repressive epigenetic modifications mediated by the histone methyltransferase G9a and the de novo DNA methyltransferases DNMT3a/b (Feldman et al., 2006). Recent studies have, however, shown that Oct4 is active in a variety of somatic stem cell compartments and in cultured multipotent somatic progenitor cells where it has been suggested to function in a manner analogous to its role in ES cells (Supplementary Table 1). Consistent with this possibility is the observation that ectopic expression of Oct4 in mice harboring a single-copy, doxycycline-inducible transgene is sufficient to prevent differentiation of intestinal epithelium while concomitantly stimulating expansion of the progenitor cell compartment (Hochedlinger et al., 2005).
Oct4 gene expression in the adult has been most frequently reported in the bone marrow of both humans and mice, particularly in hematopoietic and mesenchymal stem cells (HSCs and MSCs) (D’Ippolito et al., 2004; D’Ippolito et al., 2006; Goolsby et al., 2003; Izadpanah et al., 2006; Jiang et al., 2002; Johnson et al., 2005; Lamoury et al., 2006; Moriscot et al., 2005; Nayernia et al., 2006; Pallante et al., 2007; Pochampally et al., 2004; Ren et al., 2006), as well as in various subpoputlations of multipotent progenitors (D’Ippolito et al., 2004; D’Ippolito et al., 2006; Jiang et al., 2002; Nayernia et al., 2006; Pallante et al., 2007; Serafini et al., 2007; Zhang et al., 2005). Additionally, Oct4 expression has been detected in progenitor cells from other tissues including pancreatic islets (Wang et al., 2004), kidney (Gupta et al., 2006; Sagrinati et al., 2006), peripheral blood (Johnson et al., 2005; Romagnani et al., 2005; Tondreau et al., 2005), mammary epithelium (Tai et al., 2005), endometrium of the uterus (Cervello et al., 2006; Matthai et al., 2006), thyroid (Thomas et al., 2006), lung (Ling et al., 2006), brain (Davis et al., 2006; Okuda et al., 2004), liver (Beltrami et al., 2007), dermis, and hair follicles (Dyce et al., 2006; Dyce et al., 2004; Kues et al., 2005; Mongan et al., 2006; Redvers et al., 2006; Yu et al., 2006). In addition to these tissues, a number of studies have reported Oct4 expression in primary tumors, transformed cells, amniotic fluid, and umbilical cord blood (see Table S1 for references). Given its expression in a variety of normal tissues and transformed cells it has been suggested that Oct4 may not only be crucial for the maintenance of pluripotency in embryonic cells, but may also play an important role for the self-renewal of somatic stem cells and in maintaining tissue homeostasis. The goal of the current study was to test this hypothesis by deleting Oct4 from somatic stem cell compartments in vivo.
>Results
Results
To investigate a possible function of Oct4 expression in somatic cells we used a Cre-lox based recombination approach to achieve tissue-specific inactivation of a conditional Oct4 allele (Oct4-2lox, Fig 1a) (Kehler et al., 2004). Oct4-2lox mice were crossed to mice carrying tissue specific or inducible Cre transgenes to inactivate Oct4 in organs that have a well-defined somatic stem cell population. We deleted Oct4 in the intestine, bone marrow, brain, liver and hair follicles since Oct4 expression has previously been documented in these tissues (Table S1). The Cre transgenes that were used to achieve cell-type specific deletion of Oct4 in these tissues are described in Table 1.
Figure 1Figure 1
Regeneration of the Intestinal Epithelium after Villin-CreER Mediated Oct4 Inactivation
Intestinal Epithelium
Inactivation of Oct4 in the intestinal epithelium (including progenitor cells residing in the intestinal crypt) was achieved through the tamoxifen-induced activation of a transgene encoding a Cre recombinase/estrogen receptor fusion protein under transcriptional control of the Villin promoter (Villin-CreER) (el Marjou et al., 2004) in 8-week old mice. Recombination of the Oct4-2lox allele was confirmed by Southern Blot analysis (data not shown). Weight gain was monitored in adult mice for a period of nine months with no differences observed between control (Oct4 1lox/+, transgenic) and Oct4 mutant (Oct4 1lox/1lox, transgenic) mice (Fig S1). Histological analysis revealed a normal tissue architecture containing differentiated cell types (Paneth and goblet cells), a normal distribution of proliferating cells marked by Ki67 expression, and an absence of any Oct4 positive cells in the crypts (Fig S1). In order to test the regenerative capacity of intestinal crypt progenitor cells in the absence of Oct4, mice were subjected to a 14 Gy dose of γ-irradiation. This resulted in widespread apoptosis and cell death coupled with a growth arrest within three days. Five days after γ-irradiation, intestinal progenitor cells became highly proliferative in the presence or absence of Oct4, giving rise to new crypts that re-established a normal intestinal epithelium by the eighth day after irradiation (Fig 1c–e). In order to exclude the possibility that a small number of progenitor cells escaped recombination and were responsible for the observed regeneration and maintenance of the intestinal epithelium, we performed PCR analysis of the Oct4 genomic locus. This analysis revealed no evidence of non-recombined Oct4-2lox alleles in the newly generated tissue (Fig 1f). Finally we examined Oct4 gene expression throughout the time course of epithelial recovery after irradiation and found little to no Oct4 mRNA. Oct4 expression in the intestine was found to be 105 times lower in comparison to ES cells (Fig 1g). Further purification of crypt progenitor cells by EDTA-based fractionation of intestinal epithelium followed by gene expression analysis revealed no preference for Oct4 expression at the base of the crypt (Fig S1). These findings indicate that Oct4 function is not required for maintenance of the intestinal stem cell niche.
Mesenchymal Stem Cells
In order to examine the effects of Oct4 deletion in the bone marrow we employed an interferon-inducible Mx1-Cre transgene (Table 1) (Kuhn et al., 1995; Schneider et al., 2003). We confirmed that activation of Mx1-Cre resulted in 100% recombination in the whole marrow by PCR analysis (Fig. 2a). Subsequent separation of mesenchymal and hematopoietic cells confirmed recombination in both cell lineages (Data not shown). MSCs known to give rise to cells of chondrogenic, adipogenic, myogenic, and osteogenic lineages (Keating, 2006) were derived from both control and mutant bone marrow. These cells, as well as multipotent subpopulations of MSCs, have been the most frequently cited source of Oct4 expression in somatic tissues (Table S1). The proliferative capacity and potential for lineage commitment of MSCs was addressed using in vitro colony formation assays, osteoblast and chondrocyte differentiation assays. Individual MSCs derived from both control and Oct4 mutant MSCs were able to proliferate to form clonal colonies (Fig. 2b). Upon reaching confluence, these cells underwent osteogenic differentiation in the presence of ascorbic acid and inorganic phosphate, forming multilayered nodules containing mineralized extracellular matrix- hallmarks of mature osteoblasts (Fig. 2d). No Oct4 positive cells were observed during colony formation (Figure 2c) Gene expression analysis throughout the time course of osteogenic lineage commitment revealed no significant Oct4 gene activity in whole marrow, proliferating MSCs or in differentiated osteoblasts (Fig. 2e). Similarly, Oct4 mutant MSCs were able to activate Sox9 gene expression and enter the chondrogenic lineage in BMP treated micromass cultures (Fig. 2f & Fig. S2).
Figure 2Figure 2
Proliferative and Lineage Commitment Capacity of Bone Marrow-derived MSCs after Mx1-Cre Mediated Inactivation of Oct4
Hematopoietic Lineages
Oct4 function was addressed in the well-defined hematopoietic stem cell (Kondo et al., 2003). Examination of the peripheral blood circulation of control and Oct4 mutant mice 5, 12, and 35 days after Mx1-Cre induced recombination revealed no deficiency in the ability of Oct4 mutant mice to maintain white blood cell (WBC), red blood cell (RBC) or platelet counts at control levels throughout the experiment (Fig. 3a&b). Flow cytometric analysis of the bone marrow for monocyte, B-cell, and granulocyte lineages at 12 and 35 days after recombination showed little difference in the representation of these cell types between control and Oct4 mutant animals (Fig. 3c and not shown). Furthermore, Oct4 deletion did not affect the lineage negative, c-kit+, sca1+ population of HSCs in the bone marrow (Fig. 3d) (Camargo et al., 2006). Subsequent analysis of Flt3 negative long-term HSCS and Flt3 positive short-term HSCs showed no variation in these populations in the absence of Oct4 (not shown). Quantitative RT-PCR analysis of flow-sorted HSCs, common lyphoid progenitors, granulocytes, and monocytes revealed no preference for Oct4 expression in stem cells, and expression in all of these cell types was negligible in comparison to ES cells (Fig S2). PCR analysis for the 2-lox and recombined 1-lox Oct4 alleles 8 weeks after Mx1-Cre activation verified the persistence of Oct4 null cells in the marrow ruling out potential functional contribution of any cells escaping Cre-mediated recombination (Fig 3e). Ultimately, we tested the regenerative potential of the HSCs by performing a competitive reconstitution assay in which equal numbers of either control or Oct4 mutant marrow cells bearing a CD45.1 cell surface antigen were isolated 5 days after Mx1-Cre activation and co-injected along with CD45.2 competitor cells into lethally irradiated recipient mice. The CD45.1 and CD45.2 cells were allowed to compete for re-establishment of the recipient’s hematopoietic system and the relative contribution of each donor cell was assessed (Fig 3f). We found no significant differences in the ability of control or Oct4 mutant cells to repopulate the ablated hematopoietic systems of recipient animals. Moreover, the observed reconstitution was stable over time demonstrating that HSCs lacking functional Oct4 alleles fully retain long-term pluripotency.
Figure 3Figure 3
Hematopoietic Lineage Analysis in Mx1-Cre, Oct4 Conditional Mice
Liver
While a clearly defined stem cell niche has not been identified in the liver, several candidate progenitor cells have been described (Walkup and Gerber, 2006) and Oct4 has been observed in liver derived stem cells (Beltrami et al., 2007). To test whether Oct4 functions in liver regeneration, we performed partial hepatectomies after inactivation of Oct4 using Mx1-Cre (Kuhn et al., 1995; Schneider et al., 2003) (Fig. 4a). Two months after removal of 75% of the liver, both control and Oct4 mutant mice were able to fully regenerate lost tissue. The newly generated liver was histologically normal, containing mature hepatocytes, bile ducts, and vascularization (Fig. 4b). In addition, Oct4 protein was undetectable by immunostaining and little to no Oct4 mRNA was expressed in the regenerated liver (Fig. 4c&d). As in the intestinal epithelium, the newly regenerated liver of Oct4 mutant mice lacked non-recombined Oct4-2lox alleles (Fig 4e), demonstrating that cells lacking functional Oct4 alleles are capable of tissue regeneration.
Figure 4Figure 4
Liver Regeneration after Mx1-Cre Mediated Inactivation of Oct4
Brain
The Oct4 zebrafish ortholog spiel ohne grenzen (spg) has been shown to be necessary for formation of the mid- and hindbrain (Belting et al., 2001) and cultures of mammalian neural progenitors exhibit Oct4 expression (Davis et al., 2006; Okuda et al., 2004). We therefore examined the effect of Oct4 deletion in neural progenitor cells. A paternally transmitted Nestin-Cre transgene was used to excise the Oct4 conditional allele in all neural progenitor cells of the developing brain (Table 1)(Bates et al., 1999; Fan et al., 2001). Oct4 mutant mice exhibited no apparent behavioral abnormalities and brain morphology appeared normal more than one year after Oct4 deletion (Fig. 5a). Ki67-positive neural stem cells were seen in the subventricular zone of the lateral ventricles of both control and Oct4 mutant mice (Fig 5a&b). Oct4 expression in this region was undetectable by immunostaining (Fig 5a&b), and Oct4 mRNA nearly undetectable in both control and mutant brain extract, embryonic day 13 brain extract, as well as in purified primary cultures of neural progenitor cells (Fig 5c).
Figure 5Figure 5
CNS Analysis in Oct4 conditional, Nestin-Cre mice and Hair Follicle Analysis in Oct4 conditional, K15-CrePr1 mice
Hair Follicle
The function of Oct4 was examined in the hair follicle through activation of a Keratin1-15 CrePr1 transgene (K15-CrePr1). This transgene encodes a Cre-progesterone receptor fusion protein and is expressed exclusively in the hair follicle bulge stem cells (Morris et al., 2004). These cells give rise to all differentiated cell types in the follicle, enter the dermal lineage during wound healing (Ito et al., 2005) and have previously been shown to express Oct4 (Yu et al., 2006). The hair follicles of 8-week old mice were examined after activation of K15-CrePr1 through dermal administration of the progesterone antagonist mifepristone. We observed differentiated cell types and sebaceous glands in both control and Oct4 mutant follicles (Fig. 5d). Immunostaining with Oct4 and Ki67 antibodies revealed no Oct4 positive cells in or around the follicle and a normal distribution of proliferating cells (Fig. 5d), and qRT-PCR analysis of mRNA isolated from skin revealed no significant Oct4 expression relative to ES cells (Fig 4d). To test the regenerative capacity of the skin in control and Oct4 mutant mice, we performed wound-healing assays in which 8 mm, full-thickness dermal biopsies were monitored for 2 weeks and found no significant differences in wound-healing capacity (Fig. 5e). While recent studies have demonstrated that an additional epidermal stem cell, which may or may not have inactivated Oct4 in our model, can contribute to wound healing and follicle formation after wounding, our Oct4 K15-CrePr1 mice were able to continue hair growth during homeostatic conditions, a process known to be dependent on the K15-positive stem cells of the follicular bulge (Morris et al., 2004).
Oct4 Expression in Somatic Cells
The results described thus far failed to reveal any functional role for Oct4 in the homeostasis or regeneration of a number of somatic tissues. Because Oct4 expression in somatic tissues has been detected in stem cell populations by PCR or immuno-histological methods in a number of published reports (Table S1), we attempted to confirm these observations using another approach. We generated a reporter allele through homologous recombination in ES cells in which EGFP is expressed from the endogenous Oct4 locus (Oct4-EGFP) (Fig 1b). The fidelity of the reporter was confirmed by flow cytometric analysis of targeted ES cells. Over 95% of Oct4-EGFP ES cells strongly expressed EGFP, consistent with their pluripotent state (Fig 6a). We subsequently generated mice from Oct4-EGFP ES cells and examined EGFP expression at the single cell level in a number of tissues. Flow cytometric analysis of whole bone marrow containing multipotent MSCs and HSCs revealed no EGFP positive cell population when compared to wildtype marrow (Fig 6b). Purification of c-kit+, sca1+, lineage- HSCs derived from the bone marrow confirmed the absence of Oct4 expressing hematopoietic progenitors (Fig 6c). In addition to the bone marrow, numerous other tissues, including liver, brain, intestine, stomach, skeletal muscle, skin, lungs, heart, spleen, kidney, bladder, thymus, and prostate were examined using antibodies raised against EGFP and positive signals were never observed in contrast to teratomas derived from Oct4-EGFP ES cells, which exhibit pockets of undifferentiated cells with strong GFP expression (Fig S3 and data not shown). These findings, coupled with the failure to detect Oct4 transcripts by qRT-PCR, indicate that the endogenous Oct4 locus is inactive in the adult mouse. Our results are consistent with the Oct4 locus being effectively silenced in somatic tissues and therefore dispensable for the maintenance of somatic stem cells or tissue regeneration in the adult.
Figure 6Figure 6
Oct4 Gene Expression in Bone Marrow and HSCs
>Discussion
Discussion
Oct4 expression in somatic stem cells and in various cancer cells has been described in numerous recently published reports (Table S1). This has led to the provocative proposition that Oct4 may be involved not only in the maintenance of pluripotency in embryonic stem cells but also in the self-renewal of somatic stem cells and in the genesis of cancers (Tai et al., 2005). To test this possibility, we deleted the Oct4 gene in several tissues that have well-defined stem cell compartments and a high rate of cell turnover. Oct4 deletion in the intestine, bone marrow, hair follicle, liver, or CNS had no effect on tissue maintenance or injury-induced regeneration. Moreover, we observed no significant expression of the Oct4 gene using immunohistochemistry, quantitative RT-PCR, or flow cytometric analysis of cells harboring a targeted Oct4-EGFP reporter allele.
While Oct4 is highly expressed in ES cells it becomes silenced upon differentiation through a two-step mechanism in which histone H3 methylation precedes Dnmt3a/3b-mediated promoter methylation (Feldman et al., 2006). Oct4 activity does, however, persist in the germline where it is required for the viability of germ cells (Kehler et al., 2004). Furthermore, Oct4 is frequently expressed in tumors of germ cell origin where it acts as an oncogenic fate determinant (Gidekel et al., 2003; Looijenga et al., 2003). It is significant for this discussion that ectopic activation of Oct4 in the intestine or skin results in rapid expansion of progenitor cells and invasive tumor formation indicating that Oct4 can also act as a powerful oncogene in somatic cells (Hochedlinger et al., 2005). If Oct4 functioned in somatic stem cells and the Oct4 genomic locus existed in a state permissible for transcriptional activation in these cells, one might expect that it would be a frequently activated oncogene in cancers of somatic origin. This, however does not seem to be the case. Looijenga et al. examined the expression of Oct4 in over 3600 primary tumors representing more than one hundred tumor types and found Oct4 reactivity consistently in germ cell tumors, but only in only 3 tumors of somatic origin (Looijenga et al., 2003). In these rare Oct4 expressing somatic tumors, Oct4 expression could potentially be due to genomic rearrangements transposing the Oct4 coding sequence into the vicinity of active promoter regions rather than reactivation of endogenous Oct4 regulatory elements (Yamaguchi et al., 2005). These observations are consistent with the notion that the epigenetic mechanisms silencing Oct4 in somatic tissues are highly effective and that Oct4 is dispensable in the adult, and is only rarely activated in somatic tumors.
What then could be the significance of somatic Oct4 expression observed in numerous studies? We note that the Oct4 expression levels described in many of these studies were very low, usually detected only after 30–40 cycles of PCR amplification (Table S1). We confirmed published results in that we were able to detect low levels of Oct4 message in various progenitor cells and somatic tissue (including HSCs, MSCs, intestinal crypt progenitors, and neuroprogenitors) by quantitative RT-PCR. The amount of Oct4 detected was many fold lower than what was observed in ES cells and present even in samples that had genetically inactive Oct4 loci, raising the question as to whether the observed Oct4 expression is functionally significant or merely represents the noise of the detection method. Furthermore, the genome contains at least six Oct4 pseudogenes, the expression of which could potentially be mistaken for that of the endogenous Oct4 gene (Pain et al., 2005; Suo et al., 2005). In addition to Oct4 pseudogenes, a number of Pou-domain family members are expressed in somatic tissue, and similarities in both gene and protein sequence may also contribute to false detection of Oct4 in a number of assays. This caveat is evident in the immunohistochemistry presented in the current study. Using an Oct4 antibody, we were able to detect positive cells in several tissues, particularly the stroma of the intestine. It is clear that this staining is non-specific as it appeared in tissues lacking a functional Oct4 locus, and was not reproducible when using expression analysis or a GFP antibody in tissue sections isolated from Oct4-GFP mice. This also raises the issue of functional redundancy within the Oct family, such that the observed lack of phenotype in Oct4 mutant tissue could be due to compensation by other Oct proteins, although we believe this possibility to be unlikely due to the evolutionary divergence between Oct4 and these family members. Finally, the detection of Oct4 expression in tumor cell lines and in vitro cultures of tissue-derived cells may be of little relevance to somatic stem cell function or carcinogenesis in vivo. It is possible, for example, that Oct4 gene activation in cultured cells is a rare event and that Oct4 expressing cells may have a proliferative advantage in culture. In summary, our data strongly argue that Oct4, even if expressed at low levels in somatic cells, is dispensable for the self-renewal of somatic stem cells, for tissue homeostasis, and for the regeneration of tissue in the adult.
>Materials and Methods
Materials and Methods
Generation and Maintenance of Mice
Conditional Oct4 mice were maintained and genotyped as described previously (Kehler et al., 2004). Nestin-Cre mice were maintained and genotyped as previously described (Bates et al., 1999). In order to achieve 100% recombination of the target allele in neural progenitor cells, the imprinted Nestin-Cre transgene was passed through the male germline. Mx1-Cre mice were obtained from Jackson Labs (Bar Harbor, ME, USA). Activation of Mx1-Cre was achieved through five consecutive daily intraperitoneal injections of 0.5mg poly dI/dC (polyinosinic-polycytidylic acid, Sigma, St. Louis, MO. USA). Partial hepatectomies were performed after Mx1-Cre activation on 8-week old mice anesthetized with Avertin at a concentration of 0.5 mg/g. Regenerated hepatic tissue was isolated after an 8 week period during which mice were kept in sterile cages with antibiotic-containing water. Villin-CreER mice were a generous gift from Dr. Sylvie Robine at the Institut Curie, Paris, France. The Villin-CreER transgene was activated through five consecutive daily intraperitoneal injections of Tamoxifen (1mg in corn oil) (Sigma). K15-CrePr1Pr1 mice were a generous gift from Dr. George Cotsarelis at the University of Pennsylvania, Philadelphia, PA, USA. The K15-CrePr1Pr1 transgene was activated through administration of 1% w/w Mifepristone (Sigma) in hand cream directly to the dermis for five consecutive days. Dermal biopsies were performed 1–2 weeks after K15-CrePr1Pr1 activation and monitored for 2 weeks for wound healing ability with a micrometer.
Oct4-GFP mice were generated by subcloning a 6.5kb genomic fragment encompassing exons 2 to 5 from a BAC library into Bluescript via HindIII and EcoRI sites and termed pBS-Oct4. A loxP flanked pgk-Neo resistance cassette was cloned into the NotI site of pIRES-EGFP and subsequently the portion containing IRES-EGFP-floxed NEO (lacking the polyA) was released via BamHI and EcoRV and subcloned into the unique BclI site of pBS-Oct4 which lies between the stop codon and the polyadenylation signal. The targeting construct was linearized with SacII and electroporated into V6.5 ES cells. GFP positive colonies were picked and correct integration was verified by Southern blot analysis on genomic DNA digested with BamHI and probed with a fragment released from pBS-Oct4 with HindIII and SacI (5′ probe) resulting in a 6kb wild type band and a 9kb targeted band. Targeted ES cells were transiently electroporated with Pac-Cre to eliminate the floxed Neo cassette. Clones that had excised the neo cassette gave a 7.4kb band after digestion of genomic DNA with BamHI using the 5′ probe. All animals were treated in accordance with IACUC guidelines under current approved protocols.
Histological Analysis
Isolated tissues were fixed in 4% paraformaldehyde in phosphate-buffered saline (brain) or 10% phosphate buffered formalin (all other tissue) overnight and subsequently embedded in paraffin wax using a Tissue-Tek VIP embedding machine (Miles Scientific, Napervielle, IL) and a Thermo Shandon Histocentre 2 (Thermo FIsher Scientific, Waltham, MA). Sections were cut at a thickness of 2 um using a Leica RM2065 (Leica, Wetzlar, Germany) and stained with hematoxylin and eosin or antibodies against Oct4 or Ki67 as previously described (Hochedlinger et al., 2005). Tissues isolated from mice harboring an inducible Oct4 cDNA under control of the reverse tetracycline transactivator were used as a positive control for Oct4 immunohistochemistry (Hochedlinger et al., 2005).
For immunofluorescence, cells were fixed in a 6-well plate with 4% PFA for 10 minutes. After one wash with PBS, the cells were blocked and permeabilized in 5% FBS, .1% Triton-X for 15 minutes. Cells were subsequently incubated with a primary antibody against Oct4 at a 1:100 dilution for 1 hour at room temperature (Rabbit polyclonal, H-134; Santa Cruz Biotech, Santa Cruz, California). After 3 washes with PBS, the cells were incubated with an anti-rabbit secondary antibody labeled with Cy3 for 1 hour in the dark. After a 5 minute incubation with DAPI, the cells were washed 3 times with PBS.
Tissue Isolation and Marrow Cell Culture
Total intestinal epithelium was dissociated using a solution of 3mM EDTA and 0.05mM DTT for 30 minutes at room temperature. The musculature was discarded and purified crypts/villi were used for DNA and RNA isolation. Fractionation of the crypt-villus structure to enrich for stem cells was performed as described in (Ferraris et al., 1992). Ablation of intestinal epithelium was achieved through γ-irradation of mice in a Gammacell-40 Exactor Low-Dose Research Irradiator using a dosage of 14Gy. After irradiation mice were monitored daily and euthanized no more than eight days after irradiation. Neural progenitor cells were isolated by mechanical dissociation of E12.5 brains and cultured on polyornithine-coated tissue culture plastic in N3 media containing FGF and EGF (R&D Systems, Minneapolis, USA) as described in (Gritti et al, 2001). Whole marrow was isolated from 8 week old mice from the femur and tibia after removal of the condyles at the growth plate by flushing with a syringe and 30-gauge needle containing DMEM + 5% Fetal Calf Serum (FCS) (Hyclone, Thermo Fisher Scientific). Whole marrow was separated into mesenchymal and hematopoietic fractions through differential plating on tissue culture plates for 72 hours in α-MEM supplemented with 15% FCS (HyClone). Colony formation (CFU-F) assays were carried out by plating 4×106 nucleated cells from freshly isolated whole marrow onto 10cm plates and allowed to expand for 5 days, at which time cultures were stained with 0.1% coomassie blue to identify colonies. For osteogenic differentiation, MSCs were plated at a density of 106 cells/6-well plate and allowed to proliferate until confluent at which time osteogenesis was initiated by the addition of ascorbic acid and β-glycerol phosphate to the culture media (Owen et al., 1990). Alkaline phosphatase staining and silver nitrate staining for mineral by the method of Von Kossa were performed as previously described (Lengner et al., 2006). Chondrogenic differentiation of MSCs was carried out in high-density micromass cultures in the presence of 100ng/mL BMP2 (R&D Systems) and analyzed as previously described (Lengner et al., 2004).
Hematopoietic Lineage Analysis and Flow Cytometry
For competitive reconstitution assays, C57Bl/6-CD45.1 recipient mice were lethally irradiated (11 Gy) with γ-irradiation in a split dose, with 2 hours between doses. The reconstituting cells were injected retro-orbitally within 24 hours of irradiation. 2×105 CD45.2 whole bone marrow cells from either control or Oct4 mutant mice were transplanted along with the same dose of ‘competitor’ cells from B6-CD45.1 animals. Peripheral blood analysis was performed as previously described (Camargo et al., 2006). At various time points after transplantation, 200μL peripheral blood was collected from the retroorbital plexus of anesthetized transplant recipients. Erythrocytes were lysed and nucleated cells were simultaneously stained with anti-CD45.1, CD45.2, B220, Ly6G and Mac1 antibodies. In order to identify hematopoietic stem cells, BM was stained with antibodies against c-Kit, Sca-1, Flt3 and a lineage cocktail (composed of CD3, B220. Ter119, Mac1 and Gr-1 antibodies). All antibodies were obtained from Ebiosciences (San Diego, CA). Stained blood and bone marrow samples were then analyzed by flow cytometry using a 2-laser instrument, FACSCalibur (Becton Dickinson, Mountain View, CA), or sorted for subsequent RNA purification at the MIT FACs facility using a FACS Aria (Becton Dickinson). Differential blood counts were obtained with a Hemavet 850 (Drew Scientific, Oxford, CT).
Quantitative RT-PCR
Total RNA was isolated using Trizol reagent (Invitrogen, Carlsbad, CA). Five micrograms of total RNA was treated with DNAse1 to remove potential contamination of genomic DNA using a DNA Free RNA kit (Zymo Research, Orange, CA). One mircrogram of DNAse1 treated RNA was reverse transcribed using a First strand synthesis kit (Invitrogen) and ultimately resuspended in 100ul of water. Quantitative PCR analysis was performed in triplicate using 1/50 of the reverse transcription reaction in an ABI Prism 7000 (Applied Biosystems, Foster City, CA) with Platinum SYBR Green qPCR SuperMix-UDG with ROX (Invitrogen). Primers used for Oct4 amplification were F: 5′-ACATCGCCAATCAGCTTGG-3′, R: 5′-AGAACCATACTCGAACCACATCC-3′. To insure equal loading of cDNA into RT reactions, GAPDH mRNA was amplified using F: 5′-TTCACCACCATGGAGAAGGC-3′ and R: 5′-CCCTTTTGGCTCCACCCT-3′. Data was extracted from in the linear range of amplification. All graphs of qRT-PCR data shown represent samples of RNA that were DNAse treated, reverse transcribed, and amplified in parallel to avoid variation inherent in these procedures.
>Supplementary Material
Supplementary Material
01
Supplementary Figure 1. Long Term Effects of Oct4 Loss in the Intestinal Epithelium
(A) Growth of Villin-CreER mice was analyzed for a period of 9 months after inactivation of the Oct4 conditional allele in 8-week old mice. Weight on the final day of Tamoxifen treatment (day5) was set equal to 1. Data are mean +/− SD, n=5. (B) Histological analysis of intestinal epithelium with H&E staining 9 months after inactivation of the Oct4 conditional allele. Top panels show normal intestinal architecture. Center panels reveal goblet cells (arrows) and Paneth cells (arrowheads) in the presence or absence of a functional Oct4 gene. Ki67 staining (lower panels) shows a normal distribution of proliferating cells near the base of the villi. (C) qRT-PCR analysis after fractionation of the intestinal crypt villlus structure with early fractions (1–3) corresponding to the tip of the villi (as evidenced by Trefoil Factor-3 expression) and later fractions (4–6) corresponding to the transit amplifying compartment near the base of the crypt (marked by PCNA expression), with intestinal stem cells most represented in fraction 6 (marked by expression of the putative stem cell marker Msi1). Oct4 expression is negligible in all of these fractions when compared to ESCs. All data are mean +/− SD, relative to GAPDH, n=3. (D) Oct4 immunostaining in the control and mutant intestine reveals no Oct4+ epithelial nuclei in comparison to Oct4 M2rtTA intestinal epithelium in which Oct4 expression is ectopically induced through doxycycline administration. Non-specific staining is seen in the mesenchyme of control and mutant tissue (arrowheads).
Supplementary Figure 2. Oct4 Gene Expression in Progenitor Cells of the Bone Marrow
(A) Quantitative RT-PCR analysis of high-density micromass (MM) cultures of marrow-derived MSCs under chondrogenic culture conditions shows activation of the chondrogenic transcriptional regulator Sox9. (B) Oct4 gene expression is undetectable in chondrogenic MSC cultures in comparison to ESCs. (C) Oct4 expression in hematopoietic cell populations purified by flow sorting. All data are mean +/− SD, relative to GAPDH, n=3
Supplementary Figure 3. Oct4-EGFP in Somatic Tissues
(A) Teratoma derived from Oct4-EGFP ES cells stained with an anti-GFP antibody reveals pockets of undifferentiated Oct4-EGFP expressing cells. (B–D) Sections of liver, skeletal muscle, the lateral ventricle of the brain, and intestinal epithelium, respectively, from Oct4-EGFP mice stained with an anti-GFP antibody revealing no positive cells. (E) Intestinal epithelium from wildtype mouse stained with and anti-GFP antibody acts as a negative control.
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Acknowledgments
The authors would like to thank Jessica Dausman for assistance with animal husbandry and surgery, Dongdong Fu for histological processing, Jonathan Johnnidis for assistance with flow cytometric analyses, and the Jaenisch lab for critical comments on the manuscript. This work is supported by NIH grant RO1-CA087869, R37-CA084198, RO1-HD0445022, and a grant from Phillip Morris International. CJL is supported by an NIH Ruth L. Kirschstein NRSA F32-CA119647.
 
Footnotes
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>References
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