FADD: Essential for Embryo Development and Signaling from Some, But Not All, Inducers of Apoptosis

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Science  20 Mar 1998:
Vol. 279, Issue 5358, pp. 1954-1958
DOI: 10.1126/science.279.5358.1954


FADD (also known as Mort-1) is a signal transducer downstream of cell death receptor CD95 (also called Fas). CD95, tumor necrosis factor receptor type 1 (TNFR-1), and death receptor 3 (DR3) did not induce apoptosis in FADD-deficient embryonic fibroblasts, whereas DR4, oncogenes E1A and c-myc, and chemotherapeutic agent adriamycin did. Mice with a deletion in the FADD gene did not survive beyond day 11.5 of embryogenesis; these mice showed signs of cardiac failure and abdominal hemorrhage. Chimeric embryos showing a high contribution of FADD null mutant cells to the heart reproduce the phenotype of FADD-deficient mutants. Thus, not only death receptors, but also receptors that couple to developmental programs, may use FADD for signaling.

CD95 (the Fas antigen) is a death domain–containing receptor of the TNFR family that signals apoptosis to eliminate unwanted, autoreactive lymphocytes (1). Activation of CD95 by either CD95 ligand (CD95L or FasL) or treatment with an agonistic antibody results in receptor aggregation and the rapid recruitment of FADD (Fas-associated death domain protein), a 26-kD cytoplasmic protein with a death domain (2). The interaction of FADD and Fas through their COOH-terminal death domains unmasks the NH2-terminal death effector domain (DED) of FADD, allowing it to recruit caspase-8 to the Fas signaling complex (3) and thereby activating a cysteine protease cascade leading to cell death.

FADD and its downstream caspase cascade may also participate in signaling by other members of the TNFR family. TRADD (TNFR-1 associated death domain protein), an adapter protein that binds directly to the death domain of TNFR-1, can transduce signals for apoptosis (4). In overexpression systems, FADD is recruited by TRADD to TNFR-1 and DR3 (also called Wsl-1) signaling complexes, and FADD mutants lacking the DED are dominant-negative inhibitors of TNFR-1– and DR3-induced cell death (5, 6).

Outside of cell death signaling and the immune system, the physiological roles of FADD remain to be characterized. We have now inactivated the FADD gene in mice by gene targeting in embryonic stem (ES) cells (Fig.1, A and B) (7). Out of the first 57 live pups genotyped from heterozygous FADD matings, no homozygous mutants were observed. Therefore, timed breedings followed by genotyping were done. Up to 9.5 days of gestation (E9.5), viable homozygous FADD mutants were present in the expected Mendelian ratio (25.2%). Thereafter, viable FADD mutant embryos were obtained at a decreased ratio from E10.5 (18.8%) to E11.5 (4.1%), until all were dead by E12.5. At E11.5, FADD mutant embryos were underdeveloped and showed signs of abdominal hemorrhage (Fig. 1C). FADD mRNA was expressed in wild-type (WT) ES cells and throughout development (Fig. 1D). Protein expression, however, was decreased in the day E9.5 heterozygous fibroblasts and completely absent in homozygous mutants (Fig. 1E), indicating that theFADD mutation was null.

Figure 1

Targeting of the FADDgene. (A) Design of a FADD targeting construct. The endogenous FADD locus contains two exons (shown in boxes) and an intervening intron. The targeting construct was designed to replace the entire coding region with a PGK-Neogene cassette in the reverse orientation. Extra Eco RI sites (E) introduced by the PGK-Neo cassette and a probe (flanking probe) in the flanking region outside the construct were used for the diagnosis of homologous recombination. (B) Southern blot analysis of DNA derived from WT,FADD +/−, and FADD −/−animals. DNA was digested with Eco RI, fractionated on a 0.8% agarose gel, blotted to a nylon membrane (Hybond N, Amersham), and probed with the radiolabeled flanking probe (A). MT, mutant. (C) E11.5 WT (left) and FADD mutant (right) embryos. The reduced size of the mutant embryo is apparent. The arrowhead points to the hemorrhagic abdominal region. Bar, 500 μm. (D) Northern blot analysis of FADD gene expression during embryo development. Total RNA (10 μg) from each of the successive stages of embryo development was fractionated on a 1% denatured agarose gel, blotted to a nylon membrane, hybridized with a FADD full-length cDNA probe, and then reprobed with the β-actin cDNA. ES, embryonic stem cells; E7.5, embryonic day 7.5 after fertilization; NB, newborn. (E) Protein immunoblot analysis of FADD protein expression. Equal amounts of total protein lysates from WT, FADD +/−, andFADD −/− mice were fractionated on a 15% denatured polyacrylamide gel, and the protein immunoblot was probed with a FADD-specific polyclonal antibody. CM, cross-reacting material.

The spatial distribution of FADD between days E9.5 and E12.5 of development was analyzed by in situ hybridization (8). At E10, FADD mRNA was expressed widely, consistent with its involvement in developmental processes in the embryo (Fig.2A). Mutant embryos did not show anyFADD signal (Fig. 2B), consistent with the protein immunoblot results. At E11.5, FADD expression was concentrated in the brain (Fig. 2, C and D), myocardium (Fig. 2, C and E), liver (Fig. 2, C and F), and the developing vertebrae (Fig.2, C and G). Histological analyses of these tissues between E9.5 and E11.5 revealed that at E10.5 (Fig. 3, A and C), in ∼80% of the FADD mutant embryos, the ventricular myocardium was thinner than in their WT littermates (Fig.3, B and D). In addition, the inner trabeculation was poorly developed (Fig. 3D) (9). In contrast, the endocardial cushions appeared normal (Fig. 3D). Chorioallantoic fusion occurred normally inFADD mutant embryos (10); thus, their developmental delay was not associated with abnormal placental development.

Figure 2

Widespread FADDexpression during embryonic development. (A) Whole mount in situ hybridization in a WT E10 embryo.FADD expression (purple) is detected in all tissues, particularly in the brain, developing somites, and facial mesenchyme. No signal is found in FADD mutant embryos (B). (C) Radioactive in situ hybridization. General bright-field view of an E11.5 WT embryo. (D) to (G) correspond to the insets shown in higher magnification as dark-field views. (D toG) FADD expression in the brain (D, arrowhead), myocardium (E, arrowhead), liver (F, arrowhead), and developing vertebrae (G, arrowhead). The bright cells in the atrial cavity in (E) are unlabeled, refracting red blood cells. Bar, 200 μm (A and B), 1.3 mm (C), and 250 μm (D to G).

Figure 3

Histological and chimeric analysis of FADD mutant embryos. H&E stainings of longitudinal sections from E10.5 WT (A and B) and FADD mutant embryos (C and D). (A) Wild-type embryo. General view. (B) Detail showing the ventricular myocardium with its developing trabeculae (arrowhead). General view (C) and detail (D) of a FADD mutant embryo, showing the thin ventricular myocardium, the primitive trabeculae (arrowhead), and the normal appearance of the endocardial cushion (arrow). (Eand F) Anti-BrdU staining revealing generalized proliferation in E10.5 WT (E) and FADDmutant embryos (F). (G and H) β-Gal staining showing normal flk1 expression in the vascular endothelium of E11.5 WT (G, arrowhead) and FADDmutant embryos (H, arrowhead). (I) β-Gal staining revealing a highly chimericFADD −/− :ROSA 26 lacZ embryo that reproduces the phenotype observed in FADD −/−embryos obtained from heterozygous breedings. The arrowhead points to WT (blue) cells in the base of the yolk sac. Bar, 500 μm (A, C, and E to I); 200 μm (B and D).

Because ∼50% of the mutant embryos surviving at E11.5 were hemorrhagic, we examined the development of the vascular endothelium by breeding into the FADD mutant background a mutation for the receptor tyrosine kinase Flk1, an early endothelial marker (11). In generating the targeted mutation of theflk1 gene, the lacZ gene was knocked-in, allowing the assessment of flk1 expression inFADD −/− and WT embryos by β-galactosidase (β-Gal) staining (12). The vascular endothelium was clearly delineated in both theFADD +/+;flk1 −/+ andFADD −/−;flk1 −/+embryos (Fig. 3, G and H). Therefore, the hemorrhage observed in mutant embryos was unlikely to be due to the abnormal development of blood vessels. Because ∼80% of FADD mutant embryos exhibited a delayed and underdeveloped phenotype, we investigated whether there was an excess of apoptosis or reduced cellular proliferation. Apoptosis as measured by terminal deoxynucleotidyl transferase–mediated dUTP nick-end labeling (TUNEL) staining was not increased in FADD null mutant embryos (10). Likewise, cellular proliferation as indicated by bromodeoxyuridine (BrdU) incorporation was not drastically reduced in FADDnull mutant embryos (Fig. 3, E and F) (13).

To determine if FADD is required in specific embryonic structures, we generated chimeric embryos by injection ofFADD −/− ES cells into ROSA26 lacZblastocysts (14). These cells carry a ubiquitously expressedlacZ transgene that does not affect the viability of the embryo (12). Upon β-Gal staining, tissues generated from the recipient blastocysts stain blue, and tissues derived fromFADD −/− ES cells remain white, allowing discrimination between the contributions of mutant and WT embryonic cells. Embryos with a low contribution from the mutant ES cells showed a normal phenotype at E11.5 (10). Conversely, embryos with high chimerism (mostly white), all with high contributions ofFADD −/− cells in the heart (Fig. 3I), showed a phenotype similar to that of FADD null mutant embryos derived from heterozygous breedings. Thus, although FADD is widely expressed in the embryo, its function may be particularly important for normal cardiac development.

To determine whether FADD is required for Fas-mediated apoptosis, we transfected a Fas expression vector intoFADD −/− and FADD +/+embryonic fibroblast (EF) cells. After 15 hours, essentially all of the Fas-transfected FADD-deficient EF cells were alive, whereas only half of the Fas-transfected WT cells survived. Treatment of the transfected cells with an agonistic antibody to Fas further enhanced killing of theFADD +/+ EF cells (Fig.4A). Expression vectors for TNFR-1, DR3 (6), and DR4 (15) were transfected into the EF lines to determine whether FADD is involved in signaling apoptosis by other death domain receptors. Overexpression of either TNFR-1 (in the presence or absence of TNF treatment) or DR3 induced more apoptosis in WT cells than in FADD −/− cell lines. However, DR4 overexpression killed FADD-expressing and FADD-deficient cells equally well (Fig. 4A). Thus, FADD is required for Fas-, TNFR-1–, and DR3-induced apoptosis pathways, whereas DR4-mediated apoptosis can occur independently of FADD.

Figure 4

Defective death signaling in FADD-deficient fibroblasts. (A) Wild-type andFADD −/− embryonic fibroblasts were transiently transfected with 2 μg of each death receptor expression vector (TNFR-1, Fas, DR3, or DR4) plus 0.25 μg of pCMV–β-Gal. Fifteen hours after transfection, cells were left untreated or treated with TNF (100 ng/ml) or with an agonistic antibody to Fas (10 μg/ml) for 8 hours. Cells were then fixed and stained with X-Gal. The number of surviving blue cells were counted in 10 randomly selected high-power fields under a microscope and expressed as a percentage relative to the control vector transfection. (B) Similar experiments were performed by overexpressing cytoplasmic proteins TRADD, FADD, or caspase-8.

Furthermore, consistent with other reports (3, 5,6), overexpression of TRADD did not cause apoptosis in FADD-deficient EF cells, whereas overexpression of either FADD or caspase-8 induced apoptosis in the absence of FADD (Fig. 4B).

We next investigated the sensitivity of FADD-deficient fibroblasts to various apoptotic stimuli (16). As above, WT EF cells became sensitive to TNF in the presence of increasing concentrations of cycloheximide, whereas FADD −/− cells remained resistant to TNF (Fig. 5A). Evidence suggests that oncogenes or chemotherapeutic agents can induce apoptosis by way of the Fas signaling pathway (17). We investigated the direct apoptotic effect of c-myc by infecting EF cells with a c-myc–expressing adenovirus or a controllacZ-expressing adenovirus (Fig. 5, B and C) (18). However, we observed no significant difference in c-myc–induced apoptosis between FADD-expressing and FADD-deficient cells. Similarly, adenovirus E1A protein can induce apoptosis equally well in FADD −/− cells and in WT cells (Fig. 5D) (18). Finally, we examined the requirement of FADD for programmed cell death of oncogenically transformed embryonic fibroblasts. Oncogenic transformation byE1A and ras oncogenes renders mouse EF cells extremely sensitive to the induction of apoptosis by chemotherapeutic drugs (19). FADD-deficient cells transformed byE1A or Ras were as sensitive to various concentrations of adriamycin as their WT counterparts (Fig. 5E) (20). These results suggest that FADD is dispensable for oncogene- or drug-induced apoptosis.

Figure 5

FADD is required for TNF signaling but is dispensable for both oncogene- and adriamycin-induced apoptosis. For all parts in this figure,FADD +/+ was represented by circles andFADD / by squares (at least two separate mutant lines are represented). (A) FADD +/+ andFADD / embryonic fibroblasts were left untreated or incubated with recombinant murine TNF (10 ng/ml, Genzyme) plus various concentrations of cycloheximide as indicated. After 24 hours, viable cells were determined by negative stains of trypan blue and 7-amino-actinomycin D (26). The latter results were presented as a percentage of untreated controls. (B and C) PrimaryFADD +/+ or FADD −/− EF cells were infected with adenoviruses expressing eitherlacZ (dotted lines) or c-myc (solid lines). Trypan blue exclusion, used to determine cell viability, was performed at various time points after infection with 25 multiplicities of infection (MOI) (B) or 48 hours after infection with different MOI (C). (D) Wild-type and FADD-deficient EF cells were infected with an E1B 19K-deficient adenovirus, which induces apoptosis in a manner dependent on E1A (27). Forty-eight hours after infection with different MOI, cell viability was determined. (E) To determine the requirement forFADD in chemotherapy-induced apoptosis, we treatedFADD +/+ andFADD / E1A- and Ras-transformed EF cells with varying concentrations of adriamycin, and measured cell viability 24 hours later by trypan blue exclusion. In all cases, dead cells displayed the condensed chromatin pattern characteristic of apoptotic cell death (29). Data are the means ± SD of four separateFADD −/− EF lines and three separate WT lines.

We have shown here that apoptosis-inducing signaling by Fas, TNFR-1, and DR3 (but not by DR4) is impaired in FADD-deficient cells. This supports a report (15) that DR4 is a death-domain receptor that kills cells by a FADD-independent mechanism. In our system, FADD was also dispensable for oncogene- or drug- induced apoptosis. Together, these findings are consistent with the hypothesis that, in the absence of FADD, apoptosis induced by oncogenes and chemotherapeutic drugs may utilize DR4, related death receptors, or other pathways. Further investigations are needed to address this issue.

FADD was found to be essential for the viability of the mouse embryo and is specifically required for the development of the ventricular myocardium. The lethality of FADD null mutant mice contrasts with the viability of Fas- and TNF-R1–deficient animals (21). It is intriguing that in FADD-deficient embryos the heart is the primary site affected, despite widespread embryonic expression of the gene. This finding is similar to that in mice defective in N-myc proto-oncogene (22). Another finding that remains to be explained is the developmental retardation of FADD-deficient embryos. Little is known about the embryonic expression pattern of the characterized “death” signaling complexes. It is possible that FADD is engaged in the signaling of an as yet unidentified embryonic death receptor. Alternatively, FADD may transduce signals during embryogenesis from other unknown receptors that function primarily for embryo proliferation and survival. Further investigation of these issues is needed to clarify the roles of FADD during embryogenesis.

  • * To whom correspondence should be addressed. E-mail: t.mak{at}


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