The procaspase-8 dimer interface
The procaspase-8 dimer interface contains long-range interactions between β-strands 8 and 8' across the interface. The backbone hydrogen bonds are weak since the distances are between 3.3 Å and 3.9 Å [Fig. 1(B)]. The interface is strengthened by ring-stacking interactions between F468 from one monomer and P466' from the second monomer, as well as the symmetry-related interactions between P466 and F468'. The presence of P466 in the interface as well as reliance upon P466-F468 ring-stacking interactions introduces negative design elements into the interface. In order to examine the role of these elements in the interface of caspase-8, we replaced P466 and F468 with amino acids found in caspase-3, isoleucine, and serine, respectively. The rationale for the mutations was that the procaspase-3 homodimer has a more regular antiparallel β-sheet in the interface, with several hydrogen bonds below 3 Å distance [Fig. 1(D)]. In addition, the heterodimer of caspase-8 with FLIP shows that removal of the proline from one strand improves the inter-strand hydrogen bonds [Fig. 1(E)]. The structural model of caspase-8 lacking the negative design elements demonstrates an apparent improvement of H-bond formation in the interface [Fig. 1(C)].
As shown in Figure 2(A), the mutations were made in the context of full-length as well as ΔDED-caspase-8. In this case, the DEDs (death effector domains) are known to cause insolubility when the protein is expressed in E. coli, so the in vitro assays utilized the ΔDED constructs. The phenylalanine at 468 also was replaced with valine, rather than serine, in order to increase the hydrophobicity of the interface. In addition, two surface residues on helix 5, T441, and K473, were replaced with histidine and glutamate, respectively, in order to improve charge-charge interactions as observed in procaspase-3 [Fig. 1(F)]. We note however, that the ΔDED-casp8(T441H,K473E) variant was insoluble when expressed in E. coli, so it was not examined in the studies presented here. Finally, in order to determine whether chain cleavage of the IL is required to further improve dimer stability in the mutants, the mutations were made in the context of cleavable or uncleavable ILs (D374A,D384A) [Fig. 2(A)].
Figure 2. A: Procaspase-8 variants used in these studies. The interface mutants were placed in the contexts of fully uncleavable procaspase-8 (called D5A: D210A,D216A,D223A,D374A,D384A), cleavable intersubunit linker (called D3A - D210A,D216A,D223A), pro-less procaspase-8 (called ΔDED caspase-8), and with uncleavable linker (called D2A - D374A,D384A). The C360S mutation removes the catalytic cysteine to produce an enzymatically inactive variant. B: Activation profiles for procaspase-8 variants. Proteins were incubated in sodium citrate-containing buffer for 4 h, as described in Methods, and activity was measured at several substrate concentrations in order to determine kcat and KM values at each concentration of citrate. C: Maximum activity for each variant. For panels B and C, the numbers correspond to the mutations shown in panel A. Here, “wild-type casp-8” refers to the cleaved two-chain protein (that is, the mature heterotetramer of caspase-8), which shows the maximum activity expected.
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Procaspase-8 variants with mutated dimer interface demonstrate increased activation
The direct relationship between dimerization and activation of caspase-8 has been well-established,[11, 16, 28] so the enzyme activity of procaspase-8 is proportional to the population of dimer in the sample. Salvesen and coworkers also have shown that procaspase-8 dimerization is facilitated by sodium citrate, a kosmotrope, resulting in activation.[11, 16] To examine the effects of the interface mutations on dimerization, we measured the enzyme activity of each mutant after incubation in assay buffer that contained sodium citrate between 0M and 1M [Fig. 2(B)]. The data show that the uncleavable ΔDEDcaspase-8(D2A) [open diamonds, mutant “2” in Fig. 2(A–C)] achieves maximum activity in 0.8–0.9M citrate of ∼1.7 × 105 M−1 s−1 [Fig. 2(C)]. For the ΔDEDcaspase-8(D2A) with wild-type interface, no activity was observed below 0.6M citrate, demonstrating that the protein is monomeric below 0.6M citrate. The activity profile was similar for the P466I, F468S double mutant [open triangles, mutant “3” in Fig. 2(B,C)], the P466I,F468V double mutant [closed diamonds, mutant “4” in Fig. 2(B,C)], the single mutant T441 [open circles, Fig. 2(B), mutant “5” in Fig. 2(C)], and the triple mutant P466I, F468S, T441H [closed squares, Fig. 2(B), mutant “6” in Fig. 2(C)]. These four mutants demonstrated maximum activities in ∼0.8–0.9M citrate, although we note that the activity of the P466I, F468S was somewhat higher than that of uncleavable ΔDEDcaspase-8(D2A) (∼3.7 × 105 M−1 s−1 versus ∼1.7 × 105 M−1 s−1) [Fig. 2(C)].
Combining the interface mutations, P466I and F468V, with the single surface mutation, T441H, resulted in a significant increase in activity as well as a shift in the activity profile to lower citrate concentrations. As shown in Figure 2(B), the P466I, F468V, T441H triple mutant (closed triangles, mutant “8”) demonstrated a maximum activity at ∼0.7M citrate, with about 5-fold increase in activity compared with the uncleavable ΔDEDcaspase-8(D2A) [Fig. 2(C)].
We observed robust increases in activity upon combining the mutations for improved interface (P466I, F468S, or F468V) with the mutations for improved helix 5 surface interactions (T441H and K473E) to generate the quadruple mutants. Both quad mutants demonstrated maximum activities in ∼0.8M citrate [Fig. 2(B), open squares and closed circles, mutants “7” and “9”] at ∼1.2 × 106 M−1 s−1 [Fig. 2(C)]. Both quad mutants and the P466I, F468V, T441H triple mutant retained approximately half their maximum activity in 0.6M citrate and had significant activity in 0.4M citrate. Indeed, the maximum activities for the two quad mutants were similar to the wild-type caspase-8 with a cleavable IL. The data further support the notion that dimerization is sufficient for activity in procaspase-8.[15, 28] Stabilizing the dimer, as observed for the two quad mutants, appears to be as efficient in forming the active procaspase as cleavage of the IL in the wild-type caspase-8 [Fig. 2(C)], because the IL is not cleaved in these mutants.
The redesigned dimer interface accelerates the dimerization of procaspase-8
The data shown in Figure 2 represent the steady-state formation of the caspase-8 dimer resulting from incubation in citrate-containing buffer and the changes in the steady-state profile as a result of the interface mutations. To gain an understanding of how the mutations affect the dimerization of procaspase-8 monomers, we also examined the kinetics of dimerization for three of the interface mutants by monitoring gain of enzyme activity upon incubation in citrate (Fig. 3). It was shown previously that the dimer of caspase-8 is stabilized upon cleavage of the IL.[15, 16, 28] In order to eliminate the influence of IL cleavage on the kinetics assay, we performed the studies on the D374A, D384A uncleavable mutants [called ΔDED,D2A, Fig. 2(A)], which removes the cleavage sites in the linker. As the activity of procaspase-8 can only be detected in the dimer, the gain of activity over time represents dimerization of the monomers.
Scheme 1. Proposed model for assembly of the procaspase-8 dimer. “M” represents procaspase-8 monomer, “I” represents an enzymatically inactive dimeric intermediate, “D” represents the active dimer, and “A” represents aggregation of the monomer.
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Figure 3. Activation kinetics for procaspase-8 variants. Proteins were added to 1M sodium citrate-containing buffer to start the dimerization reaction, and aliquots were measured over time to determine activity, as described in Methods. The protein concentration assayed for enzyme activity was 20 nM, and the initial protein concentration for each experiment is shown to the right of each curve (26–333 nM). Wild-type procaspase-8 (Panel A), the double mutants, P466I,F468S (Panel B) and P466I,F468V (Panel C), and the quad mutant (P466I, F468V, T441H, K473E) were monitored for ∼250 min. In this experiment, all mutations were in the context of the pro-less (ΔDED) and IL-uncleavable (D2A) procaspase-8. Solid lines represent simulations of the data using Kinsim and the model in Scheme 1, as described in the text. Apparent rate constants determined from the simulations are provided in Table 1.
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The activation profile of wild-type procaspase-8 (that is, ΔDED,D2A) displays a protein-concentration dependence with at least two phases, the first of which is dependent on the protein concentration [Fig. 3(A)]. At higher protein concentrations, there is a rapid increase in activity in the first ∼10 min followed by a slower increase in activity over the course of the experiment (∼4 h).
The activation profile for ΔDEDcaspase-8(D2A,P466I,F468S) is similar to that for wild-type procaspase-8. The data show a rapid increase in activity within the first ∼5 min [Fig. 3(B)]. In contrast, for ΔDEDcaspase-8(D2A,P466I,F468V), the initial activity is higher than that of procaspase-8(P466I, F468S) [compare starting values in Fig. 3(C) with those in Fig. 3(B)], which we interpret as the formation of a population of dimerized protein within the dead-time of mixing, ∼30 s. This result supports the data in Figure 2 which show that the mutations improve dimerization. Following the initial activity, the data demonstrate a lag of ∼20 min followed by a protein-concentration dependent increase in activity. The maximum activity was obtained at a protein concentration of 67 nM, and the activity decreased at higher protein concentration. This trend is further amplified in procaspase-8 in the presence of four mutations (P466I,F468V,T441H,K473E) [Fig. 3(D)]. In this case, maximum activity was achieved at a protein concentration of 50 nM, and the activity decreased in higher protein concentrations.
We attempted to define a minimal model for association to explain all of the data for the four proteins shown in Figure 3. The data show that activation of procaspase-8 does not result from simple association of two monomers to yield the fully active dimer (M <__> D). Rather, the activation data show that there are multiple components that participate in activation. We propose the model shown in Scheme 1 as the simplest explanation for the activation data shown in Figure 3. We assume that there is an intermediate dimeric form that exists during activation (I), which does not contain the mature active site. Following the initial second-order reaction of dimer formation, procaspase-8 undergoes structural rearrangements to form the active protein (D) in a first-order reaction. This simple two-step model explains the protein concentration dependence in the dimerization process followed by the slower increase in activity. The rearrangement of the dimer to yield active protein is supported by previous data that shows citrate not only facilitates dimerization but also assists in active site rearrangement after dimerization. Finally, the two interface mutants containing P446 to Ile and F468 to Val, demonstrate lower activity at higher protein concentrations, indicating that the protein aggregates under these conditions. In the model shown in Scheme 1, the apparent aggregation of procaspase-8 is represented by formation of an enzymatically inactive species (A). We note two caveats to this scheme. First, rather than forming aggregates, it is possible that the procaspase-8 mutants form dimers that do not directly reflect activity (misfolded dimer). Under the conditions of these experiments, we cannot distinguish between procaspase-8 that forms soluble aggregates versus formation of dimeric species that are enzymatically inactive. In either case, the species would result in lower activity in the solution. Second, we cannot rule out more complicated assembly schemes for the procaspase-8 monomers. For example, as is evident from the simulations of the data shown in Figure 3(C), the simple model does not explain the lag phase observed in the P466I,F468V variant [Fig. 3(B)]. These results suggest a pre-existing equilibrium in the monomer (M* <__> M), where only one species is competent to dimerize. At present, however, there is insufficient data to support anything more than the simple model presented in Scheme 1. As such, we present the model as a way to compare differences in the wild-type and mutant proteins. According to this scheme for dimer assembly, k1 and k−1 represent the association and dissociation rates in dimerization, respectively (M <__> I), k2 and k−2 represent the forward and reverse rates in forming activate procaspase-8 (I <__> M), and k3 and k−3 represent the apparent aggregation step which forms an inactive dimer (M <__> A).
Overall, the simple model shown in Scheme 1 is adequate to explain all of the data in Figure 3, but it is clear that assembly of the procaspase-8 dimer is more complex than can be explained by following the gain of enzyme activity. Using the model in Scheme 1 and KinSim, we simulated the data, assuming that the reverse reactions (k−1, k−2, k−3) were zero (that is, the reactions were irreversible) because the activation assays shown in Figure 3 provide no information on the dissociation process. The solid lines in Figure 3, Panels A–D, show the results of the simulations, and the apparent rates obtained from the simulations are shown in Table 1. For wild-type procaspase-8, dimerization occurs with an apparent rate of 5.8 × 103 M−1 s−1, which is consistent with that determined previously for procaspase-8 in the presence of 1.0M citrate, 5.03 × 103 M−1 s−1 (Ref. ) and is about 100-fold faster than dimerization of procaspase-3 in the absence of citrate (70 M−1 s−1). Each of the mutants demonstrate increases in the apparent second order rate of association (M <__> I, Scheme 1) from 7.5 × 103 M−1 s−1 for the P466I,F468S double mutant [Table 1, Fig. 3(B)], to 1.7 × 104 M−1 s−1 for the P466I,F468V double mutant [Table 1, Fig. 3(C)] to 1 × 105 M−1 s−1 for the quad mutant (P466I,F468S,T441H,K473E) [Table 1, Fig. 3(D)]. There was little to no change in the apparent rate of isomerization of the dimer (I <__> D, Scheme 1), where the apparent rates were 1.7 × 104 s−1 to 3.3 × 104 s−1 regardless of the mutation (Table 1). The increase in the rate of dimerization is also consistent with the lower dependence of dimerization on sodium citrate from the steady-state measurements [Fig. 2(C)].
Table 1. Apparent rate constants from simulations of assembly kinetics for wild-type and interface mutants of procaspase-8
|Rate||Wild type Procaspase-8||Procaspase-8 (P466I,F468S)||Procaspase-8 (P466I,F468V)||Procaspase-8 (P466I,F468V T441H,K473E)|
|k1 (M−1 s−1)||5.8 × 103||7.5 × 103||1.7 × 104||1.0 × 105|
|k2 (s−1)||1.7 × 104||1.7 × 104||3.3 × 104||3.3 × 104|
|k3 (M−1 s−1)||1.7 × 104||8.3 × 104||2.5 × 105||5.0 × 105|
The simulations also showed a competing reaction of aggregation (M <__> A, Scheme 1). For simplicity, the aggregation is modeled as formation of an inactive dimer from the monomeric state, although we note that the data in Figure 3 provide no information on the oligomeric state of the aggregate, only that aggregation competes with formation of an enzymatically active dimer. Thus, the apparent rates for aggregation shown in Table 1 should not be viewed as rate constants for aggregation of the monomer, but rather the simulations show that the mutations increase the aggregation propensity of the procaspase-8 variants. In this regard, while aggregation increases somewhat for the P466I,F468S double mutant compared to wild-type procaspase-8, the P466I,F468V variants (double and quad mutants) show dramatic increases in aggregation. So, while removing the proline and phenylalanine from the dimer interface leads to increased rates of dimerization, the mutations also increase the competing aggregation reactions.
Increased dimerization in vitro is not sufficient to trigger apoptosis in cell culture
Although the in vitro activity studies show that the two quad mutants, (P466I,F468S/V,T441H,K473E) have lower dependence on sodium citrate, the activity in 0.6M citrate is ∼30–40% of that in 0.8M citrate, and below 0.3M citrate the activity is nearly zero. The cellular environment cannot provide the high kosmotrope conditions of the in vitro studies, although molecular crowding in the cytoplasm may facilitate dimerization. Thus, it was not clear that increased dimerization in vitro, resulting in a lower requirement for citrate, would manifest as constitutive activation in cellulo.
To test the effect of the mutated procaspase-8 proteins in human cells, the procaspase-8 genes were cloned into the eukaryotic expression vector pcDNA 3.1(-) to which we added a FLAG tag. The full-length procaspase-8 and the ΔDED procaspase-8 variants were examined in the context of a cleavable versus an uncleavable IL. This resulted in procaspase-8(D5A), where all five cleavage sites were replaced with alanine: D210, D216, and D223, between the DED regions and the large subunit, and D374 and D384 in the IL [Fig. 2(A)]. In procaspase-8(D3A), the three sites between the DED regions and the large subunit were mutated, D210A, D216A, and D223A, while the IL remained cleavable [Fig. 2(A)]. Finally, in ΔDED procaspase-8(D2A) the two sites in the IL, D374, and D384, were mutated to alanine [Fig. 2(A)]. The rationale for using the three backgrounds in the contexts of the interface mutants is to examine whether cleavage of the IL and/or interactions with death receptors were required for efficient apoptosis.
Transfection of HEK293A cells with the ΔDED variants did not result in significant apoptosis regardless of whether the IL was cleavable, where ∼10–18% apoptosis was observed compared with ∼10% for the empty vector [Fig. 4(A,B)]. These levels of apoptosis likely represent basal levels since removal of the catalytic cysteine, C360S, resulted in apoptosis in the range of ∼10–15%. We have shown previously that the constitutively active procaspase-3 variant, V266E, results in robust cell death. Under the conditions used here, the constitutively active procaspase-3 results in apoptosis levels of ∼30–35%. Transfection of cells with the quad mutant—P466I,F468V,T441H,K473E—in the context of D5A (uncleavable DED, uncleavable IL), D3A (uncleavable DED, cleavable IL), or D2A (ΔDED, uncleavable IL) resulted in no appreciable increase in apoptosis unless the DED motifs were included (that is, full-length proteins). The full-length uncleavable procaspase-8 with a wild-type interface demonstrated robust apoptosis of ∼30%, while the quad mutant (P466I,F468V,T441H,K473E) was somewhat lower at ∼20% [Fig. 4(A)]. While cleavage of the IL made no difference in efficiency for the wild-type procaspase-8 [that is, procaspase-8(D3A)], cleavage of the IL in the quad mutant (procaspase-8-D3A,P466I,F468V,T441H,K473E) resulted in an increase in apoptosis to levels observed for wild-type procaspase-8 and the constitutively active procaspase-3 [compare Fig. 4(A,B)]. In both backgrounds, D5A and D3A, removal of the catalytic cysteine in the quad mutant resulted in return to basal levels of apoptosis, demonstrating that the increase in apoptosis was due to the activity of the quad mutant. Western blots of the full-length proteins showed robust protein expression [Fig. 4(C)].
Figure 4. Transfection assays of procaspase-8 variants. Uncleavable (Panels A and D) or cleavable (Panels B and E) procaspase-8 variants were transfected into HEK293A (Panels A–C) or NB7 cells (Panels D–F) and levels of apoptosis were measured as described in Methods. Control experiments were performed with empty vector, pcDNA(-)FLAG, and a constitutively active procaspase-3 variant [procaspase-3(D3A,V266E)]. Western blots (Panels C and F) using antibodies to procaspase-8 or to the FLAG tag show levels of protein production. Error bars represent the standard error of three independent experiments.
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To examine whether the presence of endogenous procaspase-8 affected the activation of the mutants, we performed transfection studies in NB7 cells, a neuroblastoma cell line that lacks caspase-8. The results of transfection of NB7 cells with the procaspase-8 interface mutants were similar to those observed for the HEK293A cells [Fig. 4(D,E)]. Basal levels of apoptosis were ∼15%, and there was little difference in apoptosis for the interface mutants regardless of whether the IL could be cleaved. As in the case of HEK293A cells, robust cell death was observed in the full-length procaspase-8. While we observed an increase in apoptosis in the case of wild-type caspase-8 upon IL cleavage (∼30% to ∼40%), there was no significant change in the quad mutant with a cleavable IL. Western blots showed low levels of protein translation for the uncleavable quad mutant [Fig. 4(F)], although the mutant provides robust apoptosis. This feature was observed previously for the constitutively active procaspase-3(V266E) and was interpreted as a toxicity effect of an active caspase. Consistent with this interpretation, protein levels increased when the catalytic cysteine was removed. Overall, the transfection studies show that the interface mutations do not support increased apoptosis in either HEK293A or NB7 cells. The highest levels of apoptosis are observed when procaspase-8 retains the DED motifs, and a small increase in apoptosis is observed when the IL is cleaved.