PME-1 interacts with PP2A-B’ holoenzymes and induces a more open holoenzyme conformation
Alignment of high-resolution crystal structures of PP2A holoenzymes and the PP2A core enzyme-PME-1 complex shows that PP2A regulatory subunits in holoenzymes exert a huge steric hindrance that would exclude PME-1 binding (Fig. 1a, PP2A-B’g1 holoenzyme shown). Contradictory to this structural prediction, PME-1 co-migrates readily well with the PP2A-B’e holoenzyme (Fig. 1b), demonstrating a stable PME-1-PP2A holoenzyme interaction and the necessity of large conformational changes. We probed the holoenzyme conformation using an A-subunit FRET (fluorescence resonance energy transfer) sensor30, in which TC-FLASH produced by tetracysteine peptide (TC) fused to the C-terminus serves as an acceptor for the CFP (cyan fluorescent protein) fused to the N-terminus. Using the core enzyme and PP2A-B’g1 holoenzyme containing this FRET sensor, we showed that the energy transfer efficiency in the holoenzyme is much higher than the core enzyme, but significantly reduced upon the addition of PME-1 (Fig. 1c). This data demonstrates a much tighter holoenzyme conformation, corroborating the previous observation30, and indicates that PME-1 can interact with the holoenzyme and induce a more open holoenzyme conformation.
We next validated the interactions between PME-1 and PP2A-B’ holoenzymes in mammalian cells. We co-expressed PME-1-mRuby fusion protein and recombinant B’ regulatory subunits harboring HA-tag and assessed their interactions by co-immunoprecipitation (co-IP). PME-1-mRuby readily interacts with PP2Ac and multiple HA-tagged regulatory subunits in the B’ family, B’b, B’g1, B’g3, and B’d (Fig. 1d), suggesting that PME-1 might interact with the conserved common core in B’ subunits (Extended Data Fig. 1a). This common core is also crucial for substrate recognition 10-13. Compared to other B’ family members, B’d interacts with a much higher ratio of PP2Ac over PME-1 (Fig. 1d), suggesting that the interaction between PME-1 and the PP2A-B’d holoenzyme might be reduced by other structural elements unique in B’d (Extended Data Fig. 1a). Consistently, a much lower stoichiometric amount of PME-1 co-migrates with the PP2A- B’d holoenzyme over gel filtration chromatography (Extended Data Fig. 1b). These data support that PME-1 interacts with different B’ subunits in cells, and unique structural elements outside the common core might modulate this interaction.
PME-1 catalyzes direct demethylation of three families of PP2A holoenzymes
PME-1 activation relies on its binding to the PP2A active site23. The holoenzyme conformational changes induced by PME-1 might alleviate steric hindrance and allow PME-1 to interact with the PP2A active site, leading to methylesterase activation toward holoenzymes. To test this hypothesis, we assembled the core enzyme and three representative PP2A holoenzymes from three families using Bα, B’γ1 and PR70 with higher than 90% in vitro methylation. After incubation with PME-1, the methylation level of all three holoenzymes decreases significantly, comparable to that of the core enzyme (Fig. 1e). Our results suggest that PME-1-PP2A holoenzyme interactions enable all sequential events needed for PME-1 activation, allowing PME-1 to demethylate PP2A holoenzymes directly.
Mapping of B’-binding motifs in PME-1
The ability of PME-1 to induce conformational changes in PP2A holoenzyme led us to search for additional contacts made by PME-1 prior to its interaction with the PP2A active site. PME-1 has two ~40-residue disordered regions at the N-terminus and internal loop (Fig. 2a). The latter harbors a SLiM (251VEGII256E) highly similar to B’ substrates that interact with the conserved B’ common core via a signature motif “LxxIxE”10-13. Consistent with this observation, full-length PME-1 (PME-1 FL) interacts readily well with B’g1 as demonstrated using isothermal titration calorimetry (ITC) (Fig. 2b). The deletion of the internal loop harboring the B’-SLiM (ΔIL) abolishes their interaction completely (Fig. 2b). Intriguingly, deletion of N-terminal 18 residues (ΔN18) reduces the binding affinity by ~8 fold (Fig. 2b). To demonstrate that these PME-1 disordered regions contribute to holoenzyme interactions, we showed that, while PME-1 FL co-migrates readily well with the PP2A-B’g1 holoenzyme over gel filtration chromatography (Fig. 2c, left), both PME-1 truncations, ΔIL and ΔN18, drastically decreased PME-1 interaction with the PP2A-B’g1 holoenzyme (Fig. 2c, middle and right).
Together with dual interactions of the PME-1 core to the PP2A active site and PP2Ac-tail23, these results indicate that PME-1 makes multi-partite interactions with PP2A-B’ holoenzymes. The direct interactions between PME-1 disordered motifs and PP2A regulatory subunit are essential for stable interactions with the PP2A holoenzyme.
PME-1 inhibits substrate recognition by PP2A holoenzyme
Since the B’-binding motif in the PME-1 internal loop is similar to those in substrates (Fig. 2a), we examined the potential role of PME-1 in blocking substrate recognition by PP2A holoenzymes. Previously we elucidated the SLiM-based interactions that target PP2A-B’ to its substrates using proteomic peptide phage display (ProP-PD)10. In parallel, we also performed ProP-PD for the holoenzyme in complex with PME-1. The presence of PME-1 blocked the binding of all peptide motifs recognized by the PP2A-B’g1 holoenzyme, and reduced the counts for all B’-binding motifs to zero in phage selection (Extended Data Fig. 2a). SYT16 peptide was the strongest hits of B’ substrates identified by ProP-PD10. By competition pull-down assay, we showed that the interaction between GST-SYT16 and PP2A-B’γ1 was blocked by increasing concentrations of PME-1 (Fig. 3b). No competitions were detected between PME-1 ΔIL or PME-1 ΔN18 and the PP2A-B’γ1 substrate (Fig 3b). Our data demonstrated that PME-1 is capable of blocking substrate recognition of the holoenzyme, which relies on PME-1-holoenzyme interactions and the substrate-mimicking SLiM in PME-1 (Extended Data Fig. 2b).
Overall structure of the PP2A-B’g1-PME-1 complex
To define the structural and molecular basis that enables PME-1 to interact with and demethylate PP2A holoenzymes and dissect its multifaceted activities, we determined a three-dimensional structure of the PP2A-B’g1-PME-1 heterotetrameric complex using single-particle cryoelectron microscopy (cryo-EM). To trap the enzymatic intermediate, we assembled the complex using the fully methylated PP2A-B’g1 holoenzyme and an inactive PME-1 mutant, S156A23, followed by covalent crosslink by glutaraldehyde. After extensive 2D and 3D classifications and careful separation of the tetrameric complex particles from the unbound holoenzyme, the structure was finally determined at an overall resolution of 3.4 Å (Extended Data Fig. 3, Table 1). The PP2A-B’g1-PME-1 complex adopts a pentagram architecture with a size of 100 x 100 x 90 Å (Fig. 3a). The structure reveals multiple B’-PME-1 interfaces, large conformational changes in both the holoenzyme and PME-1, and orchestrated mechanisms for PME-1’s multifaceted activities.
The PME-1-bound holoenzyme has a similar overall architecture and maintains most intersubunit interfaces but exhibits three major structural changes compared to the unbound holoenzyme. Overlaying structures by PP2Ac showed that the last five huntingtin-elongation-A-subunit-TOR (HEAT) repeats of the A-subunit remains mostly unchanged. The N-terminal ten HEAT repeats shift significantly by increasing distances of 4 to 12 Å from HEAT repeat ten to one, resulting in a 12 Å shift in B’ that alleviates the steric hindrance for PME-1 binding (Fig. 3b). This structural observation is consistent with the sigal changes in the holoenzyme FRET sensor in response to PME-1 binding (Fig. 1c). Finally, the methylated PP2Ac tail has a drastically reduced occupancy to the A-B’ interface compared to the holoenzyme as detailed later.
PME-1-binding largely suppresses the holoenzyme activities. In addition to the direct binding and blocking of the phosphatase active site, PME-1 also occupies the B’ protein groove for recruiting substrates. PME-1 undergoes a significant angular movement away from B’ up to 6 Å pivoted at the PME-1 helix contacting the phosphatase active site (Fig. 3c). The later remains the same compared to the core enzyme-PME-1 complex (Fig. 3c). The PME-1 angular shift further accommodates B’-PME-1 interactions. This changes is different from the global allosteric changes essential for methylesterase activation, underscoring the ability of PME-1 to undergo different modes of dynamic changes.
B’-PME-1 interfaces
Three separate interfaces involving both PME-1 structured core and disordered regions govern the interactions with B’g1 (Fig. 4a). Consistent with the mapping and sequence analysis earlier (Fig. 2), the B’-docking SLiM in PME-1 occupies the substrate-binding groove similar to the substrate peptide from BubR1 (Fig. 4a). Five residues (V251, E252, I254, I255, and E256) in this SLiM form sidechain interactions with the B’g1 groove (Fig. 4b, interface I). Next to this B’ groove features a network of salt-bridge and H-bond interactions between the PME-1 core (residues N192, Q195, N196, and R199) and B’g1 (residues D180, K183, E226, and Q266), centered at the R199-E226 salt bridge interaction (Fig. 4c, interface II). Interface III is two HEAT repeats away from interface I and involves both the PME-1 structured core and the N-terminal disordered region (1-40) (Figs. 4a&d). This interface harobrs three widely separated salt bridge interactions between PME-1 residues K217, R39, and R37, and B’g1 residues D313, E353, and E399 (Fig. 4d). The rest of the N-terminal disordered region (1-36) is invisible in the electron density map, but our earlier mapping of PME-1 disordered regions suggested that PME-1 N-terminal residues 1-18 also contribute to B’g1 interaction (Fig. 2), indicating the presence of a fourth interface. The mode of PME-1 binding is likely common for all B’ subunits as the PME-1-B’g1 interface involves predominantly the B’ common core (Extended Data Fig. 1a).
To assess the function of the B’-PME-1 interfaces, we introduced mutations to PME-1 and B’g1 residues at the above three interfaces. Using in vitro pulldown assay, we showed that all single or combined interface mutations in either PME-1 or B’g1 weakened their interactions (Figs. 4e-f), underlying that all three interfaces are essential. PME-1 mutations from each individual interface all reduced PME-1 binding to the PP2A-B’ɛ holoenzyme, another B’ family member, and the combined mutations from three interfaces (3MU) completely disrupted this binding (Fig. 4g). These results demonstrate that B’-PME-1 interfaces are crucial for PME-1 interaction with all PP2A-B’ holoenzymes, consistent with the earlier data that detects interactions between the recombinant PME-1 and multiple B’ subunits in mammalian cells (Fig. 1d).
Effects of PME-1 mutations and inhibitor on different PME-1 activities
We reason that B’-PME-1 interfaces specifically dictate PME-1’s activity toward PP2A-B’ holoenzymes, but not the core enzyme. Consistent with this notion, PME-1 bearing 3MU above exhibits a drastically reduced methylesterase activity toward the PP2A-B’g1 holoenezyme, but an unaltered activity toward the core enzyme (Fig. 5a). In constrast, ABL127, a compound that blocks PME-1’s activity by producing an enzyme-inhibitor adduct at the PME-1 active site35, is expected to block PME-1’s activity toward all PP2A complexes. Consistent with this notion, ABL127 reduced PME-1 binding to both the core enzyme and the PP2A-B’ɛ holoenzyme (Fig. 5b). This result also indicates that methylesterase activation and the entry of PP2Ac tail into the PME-1 active site is essential for PME-1-holoenzyme interactions. Consistently, the cryo-EM electron density map for the PP2Ac tail is barely retained at the A-B’g1 interface, sharply different from the holoenzyme (Fig. 5c).
Our structural and biochemical observations collectively arrive at a “latch-to-induce-and-lock” model for PME-1 interaction with holoenzymes and methylesterase activation (Fig. 5d). Initial latching of PME-1 disordered regions triggers holoenzyme conformational changes, allowing PME-1 to make dual contacts to the PP2Ac active site and tail. These contacts lock a stable interaction and activate the methylesterase activity toward holoenzymes.
Uncovering and dissecting cellular PME-1 funtions in AKT-p53 signaling
The fact that PME-1 mutations at B’ interfaces selectively block its function toward the PP2A-B’ holoenzymes and PME-1 inhibitor blocks all PME-1 activities provides us an opportunity to dissect multi-faceted PME-1 functions in mammalian cells. PME-1 knockdown was shown to decrease PP2Ac methylation, reduce the phosphorylation level of AKT and ERK in cancer cells, and elevate PP2A-B’ holoenzyme activity36,37. Here we show that overexpression of PME-1-mRuby fusion protein elevates the level of AKT phosphorylation at Ser473, and ABL127 significantly suppresses this elevation (Figs. 6a-b). PME-1 with 3MU or DIL suppresses this elevation similar to ABL127 (Figs. 6a-b). In contrast, ABL127 reduces the cellular level of unmethylated PP2Ac much more than those PME-1 mutants (Figs 6a-b). These data indicate that the role of PME-1 on AKT phosphorylation is mediated by its activity toward PP2A-B’ holoenzymes, and B’-interface mutations selectively affect the demethylation of PP2A-B’ holoenzymes, unlike ABL127 that affects the demethylation of all PP2A complexes.
p53 is a tumor suppressor and functions by inducing cell cycle arrest and apoptosis in response to DNA damage. PP2A facilitates p53 activation by targeting pThr55, inhibitory phosphorylation added by TAF1 kinase that reduces p53 stabilization38,39. PP2A also regulates MDM2 phosphorylation and affects its E3 ligase activity toward p5340,41. Furthermore, AKT facilitates p53 degradation by elevating the activity of MDM241,42. The complex roles of PP2A holoenzymes in this intricate signaling network led us to dissect the versatile PME-1 functions in p53 signaling. Overexpression of PME-1-mRuby elevates p53 phosphorylation at Thr55, and ABL127 reduces this elevation (Figs. 6a-b). B’-interface mutations, 3MU and DIL, abolish this PME-1 activity comparable to or better than ABL127 (Figs. 6a-b). These data demonstrate a novel role of PME-1 in regulating p53 phosphorylation and pinpoint this cellular function to its activity toward PP2A-B’ holoenzymes. Consistently, a previous study showed that p53 pThr55 is a target site of PP2A-B’ holoenzymes38.
Next, we further demonstrated the role of PME-1 in p53 signaling during DNA damage response (DDR). Upon doxorubicin treatment to induce DDR, both total p53 protein and pThr55 were increased over time and accumulated to high levels after 24 in 293T cells (Figs. 6c-d). During DDR, the presence of ABL127 leads to a more rapid p53 accumulation, accompanied by an attenuated increase in pThr55 (Figs. 6c-d). PME-1 B’-interface mutations gave similar effects during DDR with PME-1-mRuby overexpression. PME-1 3MU leads to more rapid p53 accumulation and attenuated increase in pThr55 compared to WT PME-1 (Figs. 6e-f). These results underscore an inverse relationship between pThr55 and p53 stability and a role of PME-1 in suppressing p53 accumulation by enhancing pThr55 during DDR, and suggest that PME-1 activity toward PP2A-B’ holoenzymes might contribute to this cellular function.
In summary, we demonstrated coherent roles of PME-1 in stimulating oncogenic AKT signaling and inhibiting tumor suppressor p53 signaling by enhancing activation pS473 of AKT and inhibitory pThr55 of p53. Consistent with these observations, PME-1 amplification is found in many type of cancer and associated with poorer survival outcome (Extended Data Fig. 4). The B’-interface mutations allow us to pinpoint these PME-1 functions to its activity toward PP2A-B’ holoenzymes at both basal conditions and in reponse to DNA damage (Fig. 6g), suggesting a better strategy to target PME-1 than the active site inhibitor.