Adenomatous polyposis coli protein nucleates actin assembly and synergizes with the formin mDia1

Abstract

The tumor suppressor protein adenomatous polyposis coli (APC) regulates cell protrusion and cell migration, processes that require the coordinated regulation of actin and microtubule dynamics. APC localizes in vivo to microtubule plus ends and actin-rich cortical protrusions, and has well-documented direct effects on microtubule dynamics. However, its potential effects on actin dynamics have remained elusive. Here, we show that the C-terminal "basic" domain of APC (APC-B) potently nucleates the formation of actin filaments in vitro and stimulates actin assembly in cells. Nucleation is achieved by a mechanism involving APC-B dimerization and recruitment of multiple actin monomers. Further, APC-B nucleation activity is synergistic with its in vivo binding partner, the formin mDia1. Together, APC-B and mDia1 overcome a dual cellular barrier to actin assembly imposed by profilin and capping protein. These observations define a new function for APC and support an emerging view of collaboration between distinct actin assembly-promoting factors with complementary activities.

Figure 1.

APC directly nucleates actin assembly. (A) Schematic of APC, and Coomassie-stained gel of purified APC polypeptides. (B) GFP fluorescence and rhodamine-phalloidin staining of serum-starved NIH3T3 cells. Arrowheads, cells microinjected with EGFP-APC-B plasmid. Arrow, F-actin accumulation at cell–cell junction. Bar, 10 µm. Right panel shows linescan quantification of F-actin at cell–cell junctions in the absence (b, red line) or presence (d, red line) of GFP-APC-B. (C) Quantification of F-actin levels in cells (n > 50 cells). (D) Assembly of actin (2 µM; 5% pyrene labeled) induced by 0–200 nM APC-B. (E) Electron micrographs of actin polymerized for 30 s in the presence or absence of 50 nM APC-C. Bar, 100 nm. (F) TIRF microscopy of actin (1 µM; 30% labeled) assembled in the presence and absence of 2 nM APC-B, visualized 2–3 min after initiation of polymerization. Bar, 5 µm. (G) Severing and depolymerization of 2 µM preformed actin filaments was induced by Cof1 but not APC-B upon addition of 3 µM Vitamin-D binding protein (a monomer sequestering factor) to induce depolymerization. (H) 100 nM cytochalasin D blocks the assembly of actin (2 µM) stimulated by 20 nM APC-B. (I) APC-B does not protect filament barbed ends against 100 nM CapZ (CP) in seeded filament elongation assay.

Figure 2.

Properties of APC as an actin assembly–promoting factor. (A) Comparison of nucleation effects for 20 nM APC-B and 20 nM C-mDia1 at different concentrations of actin monomers (5% pyrene labeled). (B and C) Effects of 2 nM C-mDia1 or 20 nM APC-B on the assembly of actin (2 µM; 5% pyrene labeled) in the presence or absence of 3 µM profilin. (D) Increase in barbed end elongation rate of individual actin filaments by C-mDia1 and profilin but not by APC-B. Rates of elongation were measured in real time by TIRF microscopy and averaged for >10 filaments. Error bars, standard deviation. (E) Native PAGE assay for APC-B binding to G-actin. Reactions were loaded on gels and run toward either the cathode (top) or anode (bottom), then gels were Coomassie stained. (F) Fluorescence-based assay for concentration-dependent binding of APC-B (0–75 nM) to G-actin (100 nM; 100% pyrene labeled). Dashed line indicates binding saturation at a 1:2 molar ratio of APC-B to actin. Error bars, standard deviation (n = 3). (G) 5 µM profilin does not affect the ability of 50 nM APC-B to increase the fluorescence of pyrene–G-actin (0.1 µM, 100% labeled). Error bars, standard deviation (n = 3).

Figure 3.

Dissection of APC-B actin nucleation activity and mechanism. (A) Schematic of APC-B truncated polypeptides analyzed. The actin nucleation sequences identified (ANS1 and ANS2) are shaded blue. MTBS, microtubule binding site; N/A, not applicable; N/D, not determined. (B) Deletion of ANS1 diminishes the actin nucleation activity of APC-B in cells. Serum-starved NIH3T3 cells microinjected with EGFP-APC-B, EGFP-APC-B-ΔN4, or EGFP-APC-B-ΔN5 plasmids (arrows) were fixed and imaged for GFP fluorescence and rhodamine-phalloidin staining. Bar, 10 µm. (C) Quantification of total cellular F-actin levels (n > 50 cells). (D) Dimerization of APC-C polypeptide. Stokes radius and sedimentation coefficient were determined for MBP–APC-C and used to calculate its native molecular weight (MW). Predicted MW of monomer and dimer for MBP–APC-C are listed for comparison. (E) Dominant-negative effects of truncated APC-B polypeptides on wild-type APC-B–induced actin assembly. Reactions contain actin (2 µM; 5% pyrene labeled) and 50 nM wild-type APC-B, with or without 50 nM APC-B–ΔN4 (top) or 50 nM APC-B–ΔC2 (bottom).

Figure 4.

mDia1 and APC-B synergize to promote actin assembly in the combined presence of profilin and capping protein. (A) Reactions contain 2 µM actin monomers (5% pyrene labeled), 3 µM profilin, 2 nM CapZ, with or without 20 nM APC-B and/or 2 nM C-mDia1. (B) Quantification of elongation rates from the slopes of curves as in A. Rates are averages for reactions performed in triplicate in each of three separate experiments, where error bars represent standard error (n = 3). (C) Rhodamine-phalloidin staining of GAPDH or mDia1-depleted cells microinjected with EGFP-APC-B plasmid. Arrowheads point to injected cells. Insets show GFP signal for the same area. Bar, 10 µm. Right panel is an immunoblot showing expression levels 72 h after siRNA transfection for GAPDH and mDia1, with vinculin as a loading control. (D) Quantification of F-actin levels induced by EGFP-APC-B in GAPDH or mDia1-depleted cells (n > 40). (E) Model for synergy between APC and mDia1 in promoting actin assembly, where APC efficiently seeds polymer formation by recruiting actin monomers from a pool of profilin-actin to form a prenucleation complex. The barbed end of the seed is captured by the FH2 domain of mDia1, which processively moves with the growing barbed end, protecting it from capping proteins while accelerating elongation through FH1–profilin–actin interactions.