1295
Development 130, 1295-1305 © 2003 The Company of Biologists Ltd doi:10.1242/dev.00355
Dp1 is required for extra-embryonic development Matthew J. Kohn1, Roderick T. Bronson2, Ed Harlow3, Nicholas J. Dyson3 and Lili Yamasaki1,* 1Department of Biological Sciences, Columbia University, New York, NY 10027 USA 2Tufts University School of Veterinary Medicine, North Grafton, MA 01536 USA 3Laboratory of Molecular Oncology, MGH Cancer Center, Charlestown, MA 02129, USA
*Author for correspondence (e-mail:
[email protected])
Accepted 20 December 2002
SUMMARY Release of E2F1/DP1 heterodimers from repression mediated by the retinoblastoma tumor suppressor (pRB) triggers cell cycle entry into S phase, suggesting that E2F1 and DP1 proteins must act in unison, either to facilitate or to suppress cell-cycle progression. In stark contrast to the milder phenotypes that result from inactivation of E2Fs, we report that loss of Dp1 leads to death in utero because of the failure of extra-embryonic development. Loss of Dp1 compromises the trophectoderm-derived tissues –
specifically, the expansion of the ectoplacental cone and chorion, and endoreduplication in trophoblast giant cells. Inactivation of p53 is unable to rescue the Dp1-deficient embryonic lethality. Thus, DP1 is absolutely required for extra-embryonic development and consequently embryonic survival, consistent with E2F/DP1 normally acting to promote growth in vivo.
INTRODUCTION
p130). By contrast, only two DP family members (DP1 and DP2) have been cloned, which interact with all E2F family members equally well, and subsequently are found in trimeric complexes with all three pRB family members. Although mutations in genes encoding pRB or upstream regulators of pRB are frequently found in human tumors, intragenic mutations in the genes encoding the E2F and DP transcription factor families have not been isolated. This may be due to the dual nature of E2F/DP heterodimers, which act as bifunctional transcriptional switches to drive cell cycle-dependent target gene activation when free, and otherwise repression of target genes when complexed to pRB family members (Dyson, 1998). Moreover, free E2F/DP heterodimers stimulate S-phase entry and either subsequent proliferation or apoptosis, depending on environmental signals (Yamasaki, 1999). Given the paradoxical functions of E2F/DP heterodimers, it has been predicted that mutations in E2F or DP genes simply may be too pleiotropic for organismal survival. Inactivation of E2F family members results in a range of diverse phenotypes. We and others have reported that E2f1deficient mice display tissue-specific atrophy (e.g. testes, thyroid) and tumor predisposition (e.g. lymphoma, lung adenocarcinoma, uterine sarcoma) (Field et al., 1996; Yamasaki et al., 1996). However, loss of E2f1 can also reduce the pituitary and thyroid tumorigenesis that develops in Rb+/– mice (Yamasaki et al., 1998), and reduce the nervous system and erythropoietic defects seen in the Rb–/– embryos (Tsai et al., 1998). Taken together, it is clear that E2f1 acts as a tissuespecific growth regulator, ranging from an oncogene to a tumor suppressor. Inactivation of E2f2 results in viable adults that, when crossed to E2f1-deficient mice, are highly tumor prone (Zhu et al., 2001). Loss of E2f3 in mice results in strain-
In humans and mice, the retinoblastoma tumor suppressor (pRB) is a critical inhibitor of tumorigenesis. Human tumors that bear mutations in the RB gene or in genes encoding upstream regulators of pRB form mutually exclusive classes and collectively account for nearly all tumors, demonstrating the importance of the pRB tumor suppressor pathway (Palmero and Peters, 1996; Sherr, 1996). Rb+/– mutant mice develop neuroendocrine tumors in which loss of the wild-type allele of Rb occurs, recapitulating the loss-of-heterozygosity at the RB locus seen in human tumors (Harrison et al., 1995; Hu et al., 1994; Jacks et al., 1992). Meanwhile, Rb-deficient mice die in utero between 13.5 and 15.5 days of gestation, displaying erythropoietic defects and central nervous system abnormalities that result from excessive apoptosis, ectopic Sphase entry and failed differentiation (Clarke et al., 1992; Jacks et al., 1992; Lee et al., 1992). By contrast, mice that lack the pRB homologs p107 or p130 in mixed genetic backgrounds are viable and tumor free at 2 years of age, whereas mice that lack both p107 and p130 die perinatally (Cobrinik et al., 1996; Lee et al., 1996). Combined loss of pRB family members immortalizes mouse embryo fibroblasts and abolishes G1 arrest after γ-irradiation, demonstrating that these pRB family members have critical yet overlapping functions in vivo (Dannenberg et al., 2000; Peeper et al., 2001; Sage et al., 2000). Members of the pRB family restrict cell cycle progression and apoptosis, to a large part, through the direct repression of the E2F/DP transcription factor family (Dyson, 1998; Trimarchi and Lees, 2002). Six E2F family members have been cloned, five of which form active repressor complexes with pRB family members (E2F1-3 with pRB, E2F4-5 with p107 or
Key words: E2F transcription, Trophectoderm, Mouse
1296 M. J. Kohn and others dependent embryonic lethality and congestive heart failure in those surviving E2f3-deficient adults without obvious tumor predisposition (Cloud et al., 2002; Humbert et al., 2000b). Loss of E2f3 lessens the nervous system and erythropoietic defects in Rb-deficient embryos (Ziebold et al., 2001), while a combination of E2f3-deficiency with E2f1-deficiency accentuates the phenotype of either single mutant (Cloud et al., 2002). MEFs that are triply deficient for E2f1-E3f3 are unable to proliferate, demonstrating the importance of those E2Fs with high affinity to pRB for cell cycle progression (Wu et al., 2001). Loss of E2f4 leads to neonatal death with abnormal hematopoiesis and intestinal defects (Humbert et al., 2000a; Rempel et al., 2000), while loss of E2f5 in mice leads to juvenile hydrocephaly because of a defect in the production of cerebral spinal fluid by the choroid plexus (Lindeman et al., 1998). Finally, the simultaneous inactivation of E2F4 and E2F5 in mice results in neonatal lethality (Gaubatz et al., 2000). Clearly, E2F family members have unique roles in vivo, and loss of any single member still allows at least part of each mutant population to survive until birth. Much less is known about the roles of the DP1 and DP2 in vivo. DP1 is ubiquitously expressed at high levels in tissues and in cell lines (Gopalkrishnan et al., 1996; Wu et al., 1995). By contrast, DP2 is expressed at low levels with alternative splicing in a restricted set of tissues and cell lines (Ormondroyd et al., 1995; Rogers et al., 1996; Wu et al., 1995; Zhang and Chellappan, 1995). Despite their distinct patterns of expression, DP1 and DP2 function indistinguishably in in vitro assays, such as those for heterodimerization, DNA binding and transactivation, when overexpressed with various E2F partners and pRB family members. Overexpression of DP1 or DP2 cooperates with activated Ras to transform fibroblasts (Jooss et al., 1995) and a dominant-negative DP1 mutant arrests cells in G1 (Wu et al., 1996). Importantly, E2Fs require prior dimerization with a DP member for all functions except pRB family interaction. To delineate a role for DP1 in vivo, we inactivated the Dp1 locus in mice by homologous recombination. This paper reports that Dp1 deficiency results in embryonic lethality prior to 12.5 days of gestation, owing to a failure of extra-embryonic tissues to develop properly.
MATERIALS AND METHODS Dp1 targeting vector, selection of Dp1+/– ES cells and chimaera production Two overlapping phages were identified by screening 129Sv mouse genomic libraries and then mapped to position the Dp1 exons along their lengths. A targeting vector was constructed using 6 kb of DNA from the Dp1 genomic locus inserted into pPNT, which contains the neomycin resistance gene (neoR) and thymidine kinase gene (TK), allowing subsequent positive-negative selection. Deletion of part of exon 4 through part of exon 9 in this targeting vector removes sequences encoding amino acid residues 66-330, and inserts an inframe stop codon in exon 4. After electroporation into D3 mouse embryonic stem cells, transformants underwent positive selection with G418 (300 µg/ml) and negative selection with gancyclovir (2 µM). After 8 days, doubly resistant clones were picked, expanded and then either frozen or used to prepare genomic DNA. By Southern analysis using a 5′ external probe and a 3′ internal probe with an EcoNI digest, we identified three out of 225 doubly resistant clones
screened that had undergone homologous recombination at the Dp1 locus. Three Dp1+/– ES clones were injected into C57BL/6 blastocysts, reimplanted into pseudopregnant CD-1 recipients and one of these produced a high percentage chimaera that gave germline transmission of the Dp1-mutant allele. Production of F1 and F2 animals and genomic PCR genotyping F1 Dp1+/– animals were genotyped by Southern analysis and genomic PCR. These Dp1+/– animals were then intermated to produce the F2 offspring. F2 Dp1+/– animals were used to maintain the mutant line, with the inclusion of one backcross to C57BL/6 females during the course of this study. To genotype the animals by PCR, tail DNA was added to PCR cocktails containing the common L48 intron primer (5′-GACTCATCACAAGACTAGCGTGACC-3′) and either the Dp1 exon-specific L43 primer (5′-GACATTGAGGTGCTCAAGCGCATGG-3′) to amplify the wild-type allele (~350 bp) or the neoRspecific L28 primer (5′-CTACCCGGTAGAATTGACCTGCA-3′) to amplify the mutant allele (~300 bp). Reactions were run at 64°C annealing temperature for 39 cycles and electrophoresed on a 1.6% TAE-agarose gel to visualize products with ethidium bromide. For small amounts of DNA from manually dissected or laser-captured embryos, a different set of primers (L75, L78 and L79) was used in a combined reaction. For manually dissected embryos, yolk sacs were removed to genotype the embryo, using PCR cocktails with the common L75 intron primer (5′-GACACGTCTGATTGTTGTGAAT3′), the Dp1 exon-specific L78 primer (5′-ACTGGGGTGGGGGTGCTCACCC-3′) to amplify the wild-type allele (~180 bp) and the neoR-specific L79 primer (5′-CCTCTGTTCCACATACACTTCAT-3′) to amplify the mutant allele (~150 bp). Reactions were run at 58°C annealing temperature for 29 to 39 cycles and electrophoresed on a 2% TAE-agarose gel to visualize products. For genotyping of the p53;Dp1-double mutant animals, PCR reactions for the p53 status were performed as previously described (Jacks et al., 1994). Western blotting of embryo lysates Manually dissected embryos (E9.5) were genotyped from yolk sacs and lysed in 2× Laemmli Buffer (~10 volumes) with repeated rounds of sonication and boiling until completely solubilized. Equal amounts of total proteins (as judged by Coomassie staining) were separated on 10% SDS-PAGE and semi-dry transferred to Immobilon P for immunoblotting. The immunoreactivity of the monoclonal antibodies used were first verified as specific for each DP family member using recombinant mouse DP1 protein or human DP2 protein in western blotting experiments (data not shown). Monoclonal antibodies against DP1 (1DP06, Labvision), DP2 (G-12, Santa Cruz) and PCNA (PC10, Zymed) were used at 1 µg/ml, and visualized with an HRP-sheep antimouse IgG secondary antibody (Amersham). A rabbit polyclonal antibody against actin (RBI/Sigma) was used at a 1:250 dilution with an HRP-donkey anti-rabbit IgG secondary antibody (Amersham). Western blots were developed with an ECL kit (Amersham) and exposed to autoradiographic film. In situ embryo analysis Pregnant females were injected i.p. with BrdU/FdU mixtures (100 µg 5-bromo-2′ deoxyuridine and 6 µg 5-fluoro-2′ deoxyuridine per gram of body weight) 30 or 120 minutes before sacrifice. Decidua with embryos were isolated from the uterine horn, fixed briefly in buffered formalin and then serially sectioned. Sections were then used for Hematoxylin/Eosin staining and immunohistochemical detection of trophectoderm-derived tissues with TROMA1 (Developmental Studies Hybridoma Bank, NICHD and University of Iowa), using a biotin-goat anti-rat IgG secondary antibody and streptavidinperoxidase for amplification. Immunohistochemical detection of DP1 was performed using a monoclonal against DP1 (1DP06 from LabVision) or mouse IgG at 1 µg/ml for E8.5 and 3 µg/ml for E7.5 and developed with a mouse-on-mouse detection kit (LabVision). For
Dp1-deficient mutant mice 1297 detection of proliferation in utero, sections were measured for BrdU incorporation using a biotin-anti-BrdU antibody kit (Zymed). Slides were counterstained briefly in Hematoxylin following development with 3,3′-diaminobenzidine (DAB, Vector Labs). Laser capture microdissection Sections were stained with Hematoxylin, and dehydrated finally with HistoClear II (National Diagnostics). Microdissection was performed on an Arcturus Pixcell station on sections of decidua containing target conceptuses. Genomic DNA was recovered from the microdissected conceptuses using 0.04% proteinase K digestion in 10 mM Tris, 1 mM EDTA and 0.5% Tween 20 for 4 hours at 50°C. After inactivation of proteinase K, genomic PCR was performed as described above.
RESULTS Inactivation of the Dp1 locus To understand the role of DP1 in vivo, we have inactivated the mouse Dp1 locus by homologous recombination in embryonic stem cells. Using 129Sv genomic phage clones, we constructed a targeting vector, which deleted part or all of exons 4-9 (encoding amino acid residues 66-330), while inserting the neomycin resistance gene in the opposite orientation (Fig. 1A). After electroporation into D3 embryonic stem (ES) cells and positive-negative selection, Dp1+/– ES clones were isolated that had homologously recombined the altered regions into the Dp1 locus, as judged by Southern analysis using 5′ external and 3′ internal probes. Germline transmission from a chimera constructed with one of these Dp1+/– ES clones resulted in the establishment of an F1 generation of Dp1+/– animals. We observed no abnormalities or marked increase in tumor predisposition in Dp1+/– mice by 9 and 12 months of age over
Table 1. Timing of Dp1-deficient embryonic lethality Age
Total
Dp1+/+
Dp1+/–
Dp1–/–
Empty
Adults E15.5 E12.5 E11.5 E10.5 E9.5 E8.5 E7.5
798 9 9 19 32 154 76 43
279 2 4 4 8 34 11 10
519 2 4 9 17 78 38 21
0 0 1 3 4 33 21 9
5 0 3 3 9 6 3
Male and female Dp1+/– were interbred for timed pregnancies. Embryos were manually dissected and yolk sacs were used for genotyping.
that seen in wild-type mice. Interbreeding of these F1 Dp1+/– animals produced F2 animals, which upon genotyping by Southern analysis (Fig. 1B) were either wild type or Dp1+/– offspring. Out of 798 F2 offspring (including animals generated with one additional backcross to the C57BL/6 strain), we never recovered a viable Dp1–/– animal (Table 1). In addition, there has been no evidence for perinatal lethality of Dp1–/– pups. This experiment clearly demonstrates that Dp1 is an essential gene in mice. Timing of Dp1-deficient embryonic lethality To identify the developmental window in which the Dp1–/– embryos die, we intercrossed Dp1+/– animals and harvested embryos from pregnant females from E15.5 to E7.5. After dissection and removal of all maternal tissues, the embryos were inspected morphologically and the yolk sacs were genotyped using PCR primers specific for the wild-type or mutant allele (Fig. 1C; Table 1). Dp1–/– embryos die by E12.5, and are only rarely viable (1 out of 3) and petite at E11.5. Although non-Mendelian numbers of Dp1–/– embryos were recovered from manual dissections at E15.5 to E10.5, the expected number of Dp1–/– embryos was found by E9.5, when they appeared viable. None of the recovered Dp1–/– embryos were normal in size or appearance at E10.5 or E9.5 compared with the wild-type and Dp1+/– embryos (Fig. 2A,B). Dp1–/– embryos are smaller and developmentally delayed with regard to Fig. 1. Inactivation of the Dp1 locus. (A)The Dp1 locus is shown with the indicated coding exons and restriction enzyme sites (E, EcoRI; X, Xba; N, EcoNI). A star denotes a polymorphic 129Sv EcoNI site not seen in the C57BL/6 strain. The targeting vector and mutant Dp1 allele are shown, in which part or all of exons 4-9 (hatched boxes) is deleted with the insertion of the neoR gene in the opposite reading frame. A 5′ external probe and a 3′ internal probe (below the wild-type allele) used for Southern analysis and six primers (arrowheads) used for genomic PCR are shown. (B) Southern analysis with the 5′ external probe and EcoNI digestion was performed on F2 animals to visualize an 8.5 kb wildtype C57BL/6 band and a 6.6 kb mutant 129Sv band. Only wild-type and Dp1+/– F2 animals are recovered. (C) Genomic PCR on laser-captured embryos from microdissected decidua is shown.
1298 M. J. Kohn and others Fig. 2. Dp1-deficient embryos die despite expressing DP2. (A) Dp1+/+ and Dp1–/– embryos at E10.5 are shown for comparison. Note the small size and decreased number of somites in this Dp1–/– embryo, the most-developed of the E10.5 mutants recovered. (B) Dp1+/– and Dp1–/– embryos at E9.5 are shown for comparison. Again, note the small size and developmental delay (absence of turning, unfused neural folds) of the Dp1–/– embryo. Scales bars to the right of A and B: 1 mm. (C) Western blots of manually dissected E9.5 embryo lysates are shown with an antiDP1 monoclonal antibody (top panel), an anti-DP2 monoclonal antibody (second panel), an anti-PCNA antibody (third panel) and an anti-actin antibody (bottom panel). Lanes marked A-F are E9.5 embryos recovered from a single Dp1+/– pregnant female. A positive control E9.5 wild-type embryo is shown in the lane marked 9.5. Genotyping of yolk sacs established the genotypes shown at the bottom of the panels. (D) A developmental timecourse is shown of DP1 and DP2 protein expression in manually dissected embryos from E8.5 to E12.5 by western blotting.
morphological staging criteria (e.g. turning, somite number), and substantial variability in the degree of growth retardation was observed at E11.5-E9.5. This miniaturization occurred in the extra-embryonic tissues, as well as the embryonic tissues of the Dp1–/– conceptuses recovered. Upon manual dissection, we have always observed an abnormally small yolk sac associated with a petite Dp1–/– embryo, suggesting that the requirement for Dp1 affects the development of both extraembryonic and embryonic compartments. Expression of DP1 and DP2 in utero To verify the loss of expression of DP1 protein in Dp1–/– embryos and to assess the levels of DP2 in these mutant embryos, we genotyped E9.5 embryos obtained from intercrossing Dp1+/– animals, and then subjected embryo lysates to immunoblotting analysis with specific monoclonal antibodies. DP1 protein is not expressed in the Dp1–/– embryo extracts, but is clearly present in the wild-type embryo extracts (Fig. 2C, top panel). By contrast, all of the embryos expressed DP2 protein (Fig. 2C, second panel). Thus, the embryonic lethality, miniaturization and developmental delay of the Dp1deficient embryos occur despite the expression of DP2, demonstrating that DP1 and DP2 proteins fulfill distinct roles during development. These experiments revealed that Dp1–/–
embryos did not express more DP2 than that expressed in wild-type embryos, using actin as a loading control (Fig. 2C, bottom panel). In all of the embryo extracts, high levels of PCNA were detected (Fig. 2C, third panel), suggesting that embryonic lethality occurred despite obvious embryonic proliferation. Using western blotting, we can also detect the expression of both DP1 and DP2 proteins in lysates of manually dissected wild-type embryos from E8.5 to E12.5 (Fig. 2D). Thus, the developmental window in which Dp1–/– embryos are clearly compromised, is a period in which both DP family members are expressed. Additionally, we detected DP1 protein expression in wild-type conceptuses immunohistochemically at E7.5 and E8.5 in situ using a monoclonal antibody specific for DP1 (Fig. 3). DP1 is expressed in both the extra-embryonic and embryonic tissues at E7.5 (compare Fig. 3A with 3B). Expression of DP1 also occurs in the E8.5 embryo (Fig. 3C,F), and within extraembryonic tissues (Fig. 3C,E,F) such as the chorion, ectoplacental cone and trophoblast giant cells. At E8.5, there is a strong background of cytoplasmic staining with mouse IgG in the trophoblast giant cells (Fig. 3D,G,H); however, specific nuclear staining is evident in these trophoblast giant cells with the DP1 monoclonal antibody. Furthermore, DP1 mRNA expression using in situ hybridization has previously been documented in embryonic (E8.5-E18.5) as well as extraembryonic tissues (E8.5), including trophoblast giant cells (Gopalkrishnan et al., 1996).
Dp1-deficient extra-embryonic defects at E8.5 To examine how loss of Dp1 affected the development of earlier embryos, we analyzed in situ serially sectioned decidua containing E8.5 embryos from Dp1+/– intermatings. To genotype the embryos in situ and identify Dp1–/– embryos, we used laser-capture microdissection followed by genomic PCR (Fig. 1C). Using this method, we recovered 31 microdissected E8.5 embryos with the following genotypes: eight Dp1–/–, 18
Dp1-deficient mutant mice 1299 Fig. 3. Expression of DP1 in extra-embryonic and embryonic tissues. Conceptuses were stained immunohistochemically with an anti-DP1 monoclonal antibody (A,C,E,F) or mouse IgG (B,D,G,H). (A,B) Serial sections of a wild-type embryo at E7.5. (C-H) Serial sections of a wildtype embryo at E8.5. The regions in C,D indicated by a star and triangle are shown at higher magnification in E-H. Expression of DP1 occurs in the embryo (e), in the ectoplacental cone (epc) and in the chorion (ch). Additionally, despite the background seen with mouse IgG, clear nuclear expression of DP1 is visible in trophoblast giant cells (tg).
Dp1+/– and five wild-type embryos. Histological inspection showed that the Dp1–/– conceptuses at E8.5 were thinner and less developed than the wild-type conceptuses, appearing as the E7.5-E8.0 conceptuses would (compare Fig. 4A with 4B,C). At the distal pole of these conceptuses, Dp1–/– embryos were smaller and less developed than were the wild-type E8.5 embryos. To determine whether extra-embryonic tissues were formed correctly in the absence of Dp1, we examined the trophectoderm-derived tissues surrounding each conceptus. The ectoplacental cone (EPC) and both the primary and secondary trophoblastic giant cells arise from the trophectoderm of the implanted blastocyst. While the EPC proliferates markedly at E8.5, the primary and secondary trophoblast giant cells instead undergo successive rounds of endoreduplication reaching up to 500 times the normal content of genomic DNA (Hogan et al., 1994). All of these trophectoderm derivatives can be visualized using the trophoblast marker TROMA1 (Brulet and Jacob, 1982). Immunohistochemical detection of the EPC and trophoblast giant cells using the TROMA1 antibody revealed the abnormal formation of these structures surrounding the Dp1–/– embryos (compare Fig. 4D with 4E,F). Although present, the EPC and trophoblast giant cells were vastly under-represented in the
Dp1–/– mutant conceptuses. Normally, it is the EPC and trophoblast giant cells that must establish the implantation site in the uterine crypt, forming the essential connection to the maternal circulation through the placenta and the parietal yolk sac, which ultimately are responsible for nutrient acquisition and filtering of fetal wastes. To assess whether loss of Dp1 influenced the level of proliferation occurring in these E8.5 conceptuses in situ, we injected 5-bromo-2′ deoxyuridine (BrdU) into these pregnant Dp1+/– females (30 minutes before sacrifice) and measured BrdU incorporation immunohistochemically in serial sections (Fig. 4G-L). Remarkably, this analysis demonstrated that all of the extra-embryonic, trophectoderm-derived tissues of the Dp1–/– conceptuses display dramatic DNA replication defects. The EPC and both the primary and secondary trophoblast giant cells surrounding the implantation site incorporated little if any BrdU in the Dp1–/– conceptuses (Fig. 4H-I), yet incorporated high levels of BrdU in the wild-type conceptuses (Fig. 4G). At higher magnification, the primary trophoblast giant cells are much smaller and incorporate very little BrdU in the Dp1–/– conceptuses compared with those in the wild-type conceptuses (compare Fig. 4K-L with 4J, arrowheads). Surprisingly, at E8.5 no obvious differences in BrdU incorporation were seen in the embryonic compartment of the Dp1–/– conceptuses (Fig. 4J versus 4K-L; serpentine-shaped embryos are present).
Dp1-deficient extra-embryonic defects at E7.5 and E6.5 To further characterize the failure of the Dp1-deficient extraembryonic tissue observed at E8.5 (Fig. 4), we expanded our analysis to include Dp1-deficient embryos resulting from Dp1+/– intercrosses at E7.5 (Fig. 5) and E6.5 (Fig. 6). By examining earlier timepoints, we attempted to distinguish between two possibilities of how trophectoderm-derived tissues are compromised by the loss of Dp1. The first possibility is that loss of Dp1 limits DNA replication and thereby proliferation of the diploid chorion and EPC (from which secondary trophoblast giant cells arise), and additionally limits endoreduplication of the primary and secondary trophoblast giant cells. The second possibility is that loss of
1300 M. J. Kohn and others
Fig. 4. In situ analysis of Dp1deficient conceptuses at E8.5. Decidua containing a wild-type (A,D,G,J) and two Dp1-deficient (B-C,E-F,H-I,K-L) conceptuses were analyzed after serial sectioning. The genotypes were established for these embryos using laser-capture microdissection and then genomic PCR. (A-C) Hematoxylin and Eosin staining. (D-F) Immunohistochemical staining with TROMA1, which detects abundant trophectoderm-derived tissue in the wild-type conceptuses. (G-L) BrdU incorporation after a 30 minute in vivo labeling. (G-I) Low magnification; (J-L) high magnification. The embryo is the serpentine-like structure at the distal pole of each conceptus. The star denotes the position of the ectoplacental cone (EPC) and secondary trophoblastic giant (TG) cells at the proximal pole of the conceptus. The arrowheads denote the positions of trophoblast giant cells at the distal pole of the conceptus. Note the intensely BrdUpositive EPC and trophoblast giant cells surrounding the wild-type, but not the mutant embryos.
Dp1 results in a deficit in the number of trophoblast precursors, which give rise to trophoblast giant cells, the EPC and the chorion. This possibility could involve the premature differentiation of trophoblast precursors. Both possibilities would affect lineages derived from both the mural and polar trophectoderm. As the majority of secondary trophoblast giant cells arise from the EPC prior to E8.5, scoring BrdU incorporation at earlier timepoints can help visualize defects specifically in the EPC and secondary trophoblast giant-cell population. Normal morphology and BrdU incorporation of the EPC at earlier timepoints would suggest that no defect in the trophoblast precursor population exists. Normal morphology but abnormal BrdU incorporation in the EPC would suggest that only the proliferation of trophoblast precursors is defective. Alternatively, normal BrdU incorporation in the EPC with abnormal morphology would suggest that the trophoblast precursor population is deficient, perhaps prematurely differentiated. Finally, abnormal morphology and abnormal BrdU incorporation at earlier timepoints would prevent the assignment of the defect to only one of these two possibilities. In contrast to our analysis of embryos at E8.5, we were not able to assign genotypes to embryos in situ at E7.5 or E6.5 by laser-capture microdissection. Although we could use immunohistochemical detection of DP1 to classify expressing versus non-expressing embryos at E8.5 (data not shown), we were unable to use this methodology at E7.5 and E6.5, either due to lower levels of DP1 expression or the possible
persistence of maternally encoded DP1. Instead we relied on our morphological analysis of serial sections at E7.5 and E6.5 to find the small, developmentally delayed and abnormal embryos, which comprised approximately one-quarter of embryos analyzed and were the presumptive Dp1-deficient mutants (Fig. 5, embryos shown in B-D are presumptive mutants at E7.5; Fig. 6, embryos in B,C are presumptive mutants at E6.5). At least for E7.5, the abnormal morphology of these presumptive Dp1-deficient mutants agreed with the abnormal morphology we observed for sets of manually dissected Dp1-deficient embryos, which we genotyped unequivocally by genomic PCR. At both E7.5 and E6.5, we continued to observe impaired BrdU incorporation in the EPC and trophoblast giant cells of presumptive Dp1 mutant embryos compared with that in normal conceptuses (Fig. 5I-L; Fig. 6G-I). In addition, the EPC appears abnormally small in presumptive Dp1 mutant conceptuses compared with normal conceptuses, as judged by histological staining (Fig. 5A-D for E7.5; Fig. 6A-C for E6.5) and by TROMA1 staining at E7.5 and E6.5 (Fig. 5E-H for E7.5; Fig. 6D-F for E6.5). These qualitative results do not allow us to distinguish easily between the two possibilities outlined above (defective trophoblast proliferation or decreased numbers of trophoblast precursors). As we always observe presumptive mutants with small EPCs, which incorporate BrdU very poorly, loss of Dp1 appears to affect the number of trophoblast precursors (possibly by differentiation), as well as their ability to replicate DNA appropriately.
Dp1-deficient mutant mice 1301 Fig. 5. In situ analysis of Dp1-deficient conceptuses at E7.5. Decidua containing a wild-type (A,E,I) and three abnormal, presumptive Dp1 mutant (B-D,F-H,J-L) conceptuses were analyzed after serial sectioning. (A-D) Visualized with Hematoxylin and Eosin staining. (E-H) Immunohistochemical staining with TROMA1 to detect trophectoderm-derived tissue. (I-L) BrdU incorporation after a 30 minute in vivo labeling. The star indicates the position of the ectoplacental cone (EPC) and secondary trophoblastic giant (TG) cells at the proximal pole of the conceptus. The arrowhead indicates the position of a trophoblast giant cell at the distal pole of the wild-type conceptus.
To quantify the Dp1 deficiency in extra-embryonic tissues, we counted the total number of trophoblast giant cells or EPC cells, and scored the fraction of which were BrdU positive for embryos at E8.5 (those from Fig. 4 and two additional sets of embryos at E8.5) as well as for those at E7.5 (Fig. 5) and at E6.5 (Fig. 6). What is clear from this quantitation (Table 2) is that the total number of trophoblast giant cells and total number of EPC cells differ greatly between the wild-type and Dp1 mutant conceptuses for all timepoints, arguing that the number of trophoblast precursors contributing to the mural and polar trophectoderm derivatives is reduced substantially with the loss of Dp1. Interestingly, the percentage of trophoblast giant cells surrounding Dp1 mutant embryos that are BrdU positive at E6.5-E8.5 is moderately reduced or unchanged compared with that for wild-type embryos (Table 2). However, for all Dp1 mutant conceptuses, the extent of BrdU incorporation in trophoblast giant cells (as indicated by the intensity of the BrdU labeling per cell in Figs 4-6) is greatly diminished. This argues that the fraction of Dp1 mutant trophoblast giant cells
engaged in DNA replication is the same as that in wild-type conceptuses, but the rate of DNA replication (30 minutes of BrdU labeling for Figs 4, 5) is much slower in Dp1 mutant conceptuses. As trophoblast giant cells are exclusively engaged in successive rounds of DNA replication without mitosis (endoreduplication), the Dp1deficient defect results in fewer and smaller trophoblast giant cells with only minimal nuclear enlargement. In contrast to the situation in trophoblast giant cells, the percentage of EPC cells that are BrdU-positive in the wild-type conceptus is very different from that in the Dp1 mutant conceptus (Table 2). At E8.5, a 17- to 1.5-fold greater percentage of EPC cells are incorporating BrdU in wildtype versus mutant conceptuses. At E7.5, a 1.8-fold greater difference is seen in the percentage of EPC cells that are incorporating BrdU in wild-type conceptuses than in Dp1 mutant conceptuses. At E6.5, no difference is seen between wild-type and Dp1 mutant conceptuses; however, this could be due to the longer labeling period used (2 hours of BrdU labeling for Fig. 6) or the absence of a BrdU defect at E6.5. Nevertheless, the decreased percentages of EPC cells incorporating BrdU in the Dp1 mutant conceptuses at E8.5 and E7.5 argue that in addition to arising from reduced numbers of trophoblast precursors, Dp1 mutant EPC cells proliferate poorly by E7.5. To evaluate how the reduced BrdU incorporation in Dp1deficient conceptuses affected trophoblast giant cell maturation over time, we extended the BrdU analysis to serially sectioned extra-embryonic tissue from E8.5 and E9.5 conceptuses. The inability of Dp1-deficient trophoblast giant cells to replicate their DNA at E8.5 is even more apparent at E9.5 (data not shown), and resulted in many fewer trophoblast giant cells (in each serial section) with markedly smaller nuclei, presumably owing to fewer rounds of endoreduplication. Clearly, the malformation and defective proliferation of the EPC, combined
1302 M. J. Kohn and others Table 2. Reduced cell number and BrdU incorporation in Dp1-deficient extra-embryonic tissues Trophoblast giant cells Genotype*
Number of cells
E8.5
+/+ +/+ –/– –/–
257 337 81 48
E8.5
+/+ +/+ –/– –/–
E8.5
Ectoplacental cone cells
% BrdU+ cells
Number of cells
221 257 65 38
86.0 76.3 80.2 79.2
265 326 83 60
149 214 31 57
115 152 19 35
77.2 71.0 61.3 61.4
155 156 Not determined 60
+/+ –/– –/–
261 41 55
215 17 33
82.4 41.5 60.0
258 51 29
191 9 6
74.0 17.6 20.7
E7.5
w m m m
69 31 17 32
49 21 11 21
71.0 67.7 64.7 65.6
125 55 32 67
90 40 13 27
72.0 72.7 40.6 40.3
E6.5
w m m
45 22 12
35 19 10
77.8 86.4 83.3
141 55 48
100 43 35
70.9 78.2 72.9
Timepoint
Number of BrdU+ cells
Number of BrdU+ cells 224 276 7 3 118 116 Not determined 30
% BrdU+ cells 84.5 84.7 8.4 5.0 76.1 74.4 Not determined 50.0
*For E7.5-E6.5, w represents normal wild-type morphology and m represents abnormal mutant morphology.
with the failure of the trophoblast giant cells to endoreduplicate in the Dp1–/– embryos, result in the petite size and subsequent failure of the Dp1-deficient mutant embryos between E8.5 and E11.5. Loss of p53 fails to rescue Dp1-deficient embryonic lethality During the course of these studies, we considered an alternative mechanism for embryonic lethality of Dp1-deficient embryos: that loss of Dp1 may have increased apoptosis in utero. Although we observed numerous TUNEL-positive cells in the EPC and at the distal tip of the implantation site on all conceptuses, we have seen few if any apoptotic cells in the embryonic region of any conceptus at E8.5. So, if increased apoptosis were responsible for Dp1-deficient embryonic lethality, it is more likely that it results from apoptosis in the extra-embryonic region of the conceptus or in the surrounding deciduum. Presumably, such extra-embryonic or decidual apoptosis would have to be distinct from the p53-mediated apoptosis induced by ionizing radiation, which is restricted to only the embryonic region of the fetus (Heyer et al., 2000). Importantly, it is well established that overexpression of E2F1/DP1 in tissue culture is associated with induction of apoptosis, which can be either p53 dependent or p53 independent (reviewed by Ginsberg, 2002; Melino et al., 2002), and which can be suppressed by co-expression of pRB. E2F1/DP1-mediated apoptosis can occur via the induction of specific target genes, such as Arf (Bates et al., 1998; DeGregori et al., 1997), which stabilizes p53 by inhibiting Mdm2-mediated degradation, and/or p73, the p53 homolog with apoptotic capabilities (Irwin et al., 2000; Stiewe and Putzer, 2000). Theoretically, loss of Dp1 could lead to increased apoptosis by preventing pRB-mediated repression of p19ARF or p73. To determine whether p53-dependent apoptosis (e.g. at the implantation site) played a role in the death of the Dp1deficient embryos, we crossed Dp1+/– animals to p53 mutant animals. If the lethality of the Dp1-deficient embryos were due
to excessive apoptosis in utero, then inactivation of p53 could reduce this apoptosis leading to improved viability, as has been shown previously for Mdm2-deficient and XRCC4-deficient embryos (Gao et al., 2000; Jones et al., 1995; Montes de Oca Luna et al., 1995). Inactivation of p53 failed to produce any adult animals that lacked Dp1 (Table 3). Analysis of five p53–/–;Dp1–/– or p53+/–;Dp1–/– embryos from double mutant crosses at E10.5 revealed that inactivation of p53 clearly did not rescue the Dp1-deficient embryonic lethality even partially in utero (data not shown). Furthermore, from intercrossing p53+/–;Dp1+/– double mutants, we also obtained p53–/–;Dp1+/– adult animals (Table 3). However, this genotype was represented at only half the expected frequency (observed=27, expected=52), a highly significant reduction (P=0.0005, χ2 test). The cohort of viable male and female p53–/–;Dp1+/– adults was recovered without obvious perinatal lethality, suggesting that lethality of approximately half of this cohort occurred sometime prior to birth. Analysis of three p53–/–;Dp1+/– embryos from a double mutant cross at E10.5 did not reveal any differences relative to p53+/–;Dp1+/– or p53+/–;Dp1+/+ embryos that might have contributed to this Table 3. Recovery of p53;Dp1 double mutant mice Genotypes of offspring from p53+/–;Dp1+/– intercrosses Genotypes p53–/–;Dp1–/– p53+/–;Dp1–/– p53+/+;Dp1–/– p53–/–;Dp1+/– p53+/–;Dp1+/– p53+/+;Dp1+/– p53–/–;Dp1+/+ p53+/–;Dp1+/+ p53+/+;Dp1+/+ Total recovered
Expected ratio
Observed number
1 2 1 2 4 2 1 2 1
0 0 0 27 91 56 18 49 29 270
M, F
Comments
0, 0 0, 0 0, 0 19, 8 42, 49 30, 26 15, 3 29, 20 19, 10
Lethal Lethal Lethal ~50% viable Viable Viable Viable Viable Viable
Dp1-deficient mutant mice 1303 Fig. 6. In situ analysis of Dp1-deficient conceptuses at E6.5. Decidua containing a wild-type (A,D,G) and two abnormal, presumptive Dp1 mutants (B,C,E,F,H,I) conceptuses were analyzed after serial sectioning. (A-C) Hematoxylin and Eosin staining. (D-F) Immunohistochemical staining with TROMA1 to detect trophectoderm-derived tissue. (G-I) BrdU incorporation after a 2 hour in vivo labeling. The star indicates the position of the ectoplacental cone (EPC) and secondary trophoblastic giant (TG) cells at the proximal pole of the conceptus. The arrowheads indicate the positions of a trophoblast giant cell.
reduced viability. These data indicate that the cause for this decreased recovery of p53–/–;Dp1+/– animals is distinct from the extra-embryonic mechanism responsible for the death of the Dp1–/– embryos. However, 3 (2 females and 1 male) out of 13 p53–/–;Dp1+/– embryos recovered at E13.5 displayed exencephaly, which has been described previously for a subset of the p53–/– females (Sah et al., 1995). In fact, we recovered fewer female versus male p53–/–;Dp1+/– adult animals (Table 3, 8 females versus 19 males). DISCUSSION Embryonic lethality of Dp1-deficient embryos The first critical role for Dp1 defined by this study is the absolute requirement of Dp1 for embryonic survival. The timing of Dp1-deficient embryonic lethality demonstrates that the in vivo temporal requirement for Dp1 clearly surpasses that of any other gene in the pRB, E2F and even the CKI (INK4 and CIP/KIP) families. Dp1-deficient embryos die prior to E12.5 because of obvious extra-embryonic defects in trophectodermderived compartments. After implantation of blastocysts, the expansion and differentiation of the trophectoderm absolutely require DP1. From our qualitative and quantitative analysis of
Dp1 mutant embryos between E6.5 and E8.5, it is clear that the number of trophoblast precursors is reduced with loss of Dp1, leading to substantially fewer cells in the EPC and chorion and fewer trophoblast giant cells. This could be due to premature differentiation of these trophoblast precursors in the absence of Dp1. Furthermore, the percentage of diploid EPC cells incorporating BrdU is drastically reduced in the Dp1 mutants, while the extent of BrdU incorporation is reduced in the endoreduplicating primary and secondary trophoblast giant cells. The failure of the EPC, the chorion and the trophoblast giant cells clearly compromises the growth and maturation of the Dp1deficient embryo. Proper development of extraembryonic tissues is crucial for the formation of the placenta, through which oxygenation, nourishment and waste removal occur, and upon which the embryo is totally dependent. Consistent with the well-documented role of E2F/DP heterodimers in stimulating S-phase entry by transactivating the ensemble of E2F target genes required for DNA replication (DeGregori et al., 1997; Muller et al., 2001), the greatly reduced BrdU incorporation in the Dp1-deficient EPC (percentage of cells) and Dp1-deficient trophoblast giant cells (extent per cell) strongly suggest that at least one major mechanism for embryonic lethality is the failure to replicate extra-embryonic DNA appropriately. Presumably, this is due to the inability of the trophectoderm-derived tissues of the Dp1–/– conceptuses to induce the large number of E2F target gene products necessary for S-phase entry and successive rounds of endoreduplication. Interestingly, the endoreduplication failure of the mouse trophoblast giant cells reported here is reminiscent of the inability of Drosophila E2F and DP mutants to amplify the chorion gene as part of the normal polytenization observed in ovarian follicle cells (Royzman et al., 1999). However, in that case, E2F appears to associate with ORC complexes suggesting a direct role may exist for RBF and E2F/DP heterodimers in endoreduplication (Bosco et al., 2001). Currently, we have no such evidence for the direct participation of mammalian E2F/DP1 heterodimers in the replicative machinery of the extra-embryonic genomes. The requirement for DP1 in the Dp1-deficient extraembryonic tissues correlates with growth retardation for the embryo. Developmental delay and runting is at least partly due to the progressive failure of the extra-embryonic tissues to form the placenta correctly, which must develop properly to sustain the growing embryo with nutrients and prevent hypoxia and
1304 M. J. Kohn and others waste accumulation. At this time, however, it is impossible to rule out the possibility that DP1 also has embryonic roles, which are masked in this study by the overwhelming failure of the extra-embryonic compartment. By E8.5, the expansion and massive endoreduplication seen in the extra-embryonic compartment (the EPC, chorion and the trophoblast giant cells) may exhaust the maternally encoded E2F target gene products for S-phase entry and DNA replication faster than the requirement for these same maternally encoded products in the embryonic compartment. Accordingly, it is likely that proliferation defects within the Dp1-deficient embryos would become apparent if the placental defect from Dp1-deficiency were rescued, revealing later roles for Dp1 in the embryo proper. The death of Dp1-deficient embryos in utero is in stark contrast to the much less severe adult and neonatal phenotypes seen with the loss of E2F family members. Thus, it must be concluded that the requirement for one half of the heterodimer, DP1, greatly exceeds the necessity for the E2F half of the complex during embryonic development. This is due in part to the fact that the DP family differs in many respects from the E2F family (Dyson, 1998). The E2F family is larger (E2F1-6) and has partial biochemical redundancy built into it (E2F1-3 bind pRB versus E2F4-5 bind p107 or p130). Inactivation of E2F1 compromises transcriptional activation through itself and transcriptional repression due to complex formation of E2F1 with pRB. By contrast, the DP family is small (DP1 and DP2), and biochemically redundant; yet, DP1 is the major family member expressed. Thus, loss of Dp1 potentially compromises transactivation as well as repression through all pRB family members, leading to a more pleiotropic phenotype. The death of the Dp1-deficient embryos underscores the vital importance of E2F/DP heterodimers for DNA replication during embryonic development. Effect of p53-deficiency on Dp1 mutants Loss of p53 failed to rescue Dp1-deficient embryonic lethality. Minimally, this means that p53-dependent apoptosis is not involved in the failure of the Dp1-deficient extra-embryonic tissues to develop. Formally, it is still possible that p73dependent apoptosis may be involved in the Dp1-deficient phenotype. A much more likely scenario is that loss of Dp1 compromises the number of trophoblast precursors, as well as their proliferation. Only half the expected number of p53–/–;Dp1+/– animals survived past birth. The absence of any obvious deformities or defects in p53–/–;Dp1+/– embryos at E10.5 when Dp1-deficient embryos are severely compromised, indicates that the mechanism for the death of p53–/–;Dp1+/– animals cannot be the same as the extra-embryonic failure responsible for the death of the Dp1-deficient embryos. Instead, these animals die of exencephaly similar to that reported previously for the p53deficient females (Sah et al., 1995). Conclusions This study defines a crucial role for DP1 in vivo, the proper development of the extra-embryonic lineages arising from the trophectoderm. The participation of DP1 in this process demonstrates its significant biological importance. Unexpectedly, the nature of the Dp1-deficient extra-embryonic phenotype demonstrates that the role of DP1 in vivo can be highly lineage specific. At this time, it is impossible to rule out
that DP1 plays additional later or more-subtle roles in the embryo. To some degree, the lineage specificity of the DP1 mutant phenotype reflects the existence and possible reliance of the mouse on DP2 for overlapping functions in most tissues. Assessment of the potentially essential nature of DP2 awaits its targeted inactivation. L. Y. especially thanks T. Jacks, John Mkwandawire and the Jacks Laboratory for their generosity, and members of the Laboratory of Molecular Oncology for help through the initial phase of this work. The authors thank the past and present members of the Yamasaki Laboratory for their helpful discussions. The authors also thank K. Manova at MSKCC for her support with the laser capture microdissection, A. Silva for his 129Sv mouse genomic library and C.-L. Wu for the GST-DP2 recombinant clone. L. Y. thanks M. Pagano and C. Prives for critical reading of this manuscript, and M. and I. Pagano for their continual support throughout this project. M. J. K. thanks J. Boyle for her support. This work was supported by grants from the NIH-NCI (#CA79646), the Pew Scholars Program (#P0229SC) in the Biomedical Sciences and the Special Fellows Program from the Leukemia Society of America (#3601-98).
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