Spinophilin directs protein phosphatase 1 specificity by blocking substrate binding sites

Journal name:
Nature Structural & Molecular Biology
Year published:
Published online


The serine/threonine protein phosphatase 1 (PP1) dephosphorylates hundreds of key biological targets. PP1 associates with ≥200 regulatory proteins to form highly specific holoenzymes. These regulatory proteins target PP1 to its point of action within the cell and prime its enzymatic specificity for particular substrates. However, how they direct PP1's specificity is not understood. Here we show that spinophilin, a neuronal PP1 regulator, is entirely unstructured in its unbound form, and it binds PP1 through a folding-upon-binding mechanism in an elongated fashion, blocking one of PP1's three putative substrate binding sites without altering its active site. This mode of binding is sufficient for spinophilin to restrict PP1's activity toward a model substrate in vitro without affecting its ability to dephosphorylate its neuronal substrate, glutamate receptor 1 (GluR1). Thus, our work provides the molecular basis for the ability of spinophilin to dictate PP1 substrate specificity.

At a glance


  1. The unbound spinophilin PP1 binding domain.
    Figure 1: The unbound spinophilin PP1 binding domain.

    (a) Domain-structure map of spinophilin with the conserved RVXF motif highlighted in green, the PP1 binding domain in magenta and the PDZ domain in purple (color codes are constant throughout all figures). (b) 2D [1H,15N] HSQC spectrum of spinophilin417–494. The blue bar highlights the lack of dispersion for the chemical shifts in the 1HN dimension. (c) An overlay of the 2D [1H,15N] HSQC spectra of spinophilin417–602 (purple) and spinophilin417–494 (magenta). The blue bar highlights the larger dispersion for the chemical shifts in the 1HN dimension resulting from the well-ordered PDZ domain, whereas the chemical shifts of spinophilin417–494 remain unchanged. Red asterisks indicate side chain NH2 groups of asparagine and glutamine. (d,e) secondary-structure propensity scores (d) and 15N[1H]-NOE (hetNOE) (e) data are plotted against spinophilin residue numbers. These data indicate that there are no regions of transient structure nor reduced backbone motions in spinophilin417–494. Also, no areas of considerably populated transient secondary structure were detected. Regions of considerably populated transient structure were considered five residues or more with secondary-structure propensity score >0.2, indicating a transient α-helix, or <−0.2, indicating a region with extended structure (not considerably populated regions are indicted by a gray box).

  2. The spinophilin417-583-PP1[alpha]7-330 complex.
    Figure 2: The spinophilin417–583–PP1α7–330 complex.

    (a) Isothermal titration calorimetry of purified spinophilin417–583 and PP1α7–330, confirming that the intrinsically unstructured spinophilin PP1 binding domain is active. (b) Cartoon representation of the spinophilin417–583 PP1 binding (magenta) and PDZ domains (purple) and PP1α7–330 (gray, surface representation) complex; centered on the active site of PP1α7–330. Two Mn2+ ions (pink spheres) mark the active site of PP1. The newly formed secondary-structure elements of spinophilin417–494 are labeled β1, β2 and α1. The PDZ domain is C-terminal to α1. Spinophilin residues interacting with the RVXF binding pocket are represented as sticks in green and those interacting with the PP1 C-terminal groove are represented as sticks in cyan. (c) The spinophilin PP1 binding domain before (unstructured model, left) and after binding to PP1 (crystal structure without PP1, right), illustrating the unfolded-to-folded transition. The four unique regions that characterize the interaction of spinophilin (magenta) with PP1 are numbered; RVXF, green; C-terminal groove binding motif, cyan.

  3. Spinophilin's interaction with the PP1 RVXF binding pocket.
    Figure 3: Spinophilin's interaction with the PP1 RVXF binding pocket.

    (a) 90° rotation of Figure 2b to highlight the interaction of Ile449 and Phe451 in the RVXF binding pocket. (b) Spinophilin residues (RVXF sequence, green) and PP1 (gray, surface and stick representation for interacting residues) are shown with oxygen atoms in orange and nitrogen atoms in dark blue. Hydrogen bonds are represented as black dashes. The spinophilin RVXF sequence (447RKIHF451) is preceded by a long loop that folds back on itself (Fig. 2b,c) to form a strong hydrogen bond network. Some residues in the loop have been omitted for clarity.

  4. Spinophilin's PP1 interaction regions II, III and IV.
    Figure 4: Spinophilin's PP1 interaction regions II, III and IV.

    (a) Region II: spinophilin forms two novel β-strands upon binding PP1. Stick representation of spinophilin residues 428–435 (β1 includes residues 430–434) and 454–461 (β2 includes residues 456–460), which form an extended β-sheet with the strands β13 and β14 of PP1. (b) Region III: spinophilin forms a four-turn α-helix upon interaction with PP1. View of the helix highlighting electrostatic and hydrophobic interaction clusters between the helix and the surface of PP1. (c) Region IV: spinophilin residues 462–469 interact directly with the C-terminal groove of PP1. These residues protrude directly into the binding pocket, forming a complex hydrogen bonding network both with PP1 and within spinophilin. Spinophilin residues 465 and 468 have been omitted for clarity.

  5. Spinophilin creates a unique holoenzyme with novel substrate specificity.
    Figure 5: Spinophilin creates a unique holoenzyme with novel substrate specificity.

    (a) The binding of spinophilin WT and mutants (region I: I449A, H450A, F451A; region II: Q457A, F459D, T461A; region III: E482A, E486A; region IV: Y467A, R469A, R469D) to PP1α7–330 was determined using a capture assay. Elution fractions and all loaded spinophilin samples were subjected to SDS-PAGE and Coomassie blue staining. (b) Densitometry analysis of gels as shown in a, to determine the amount of spinophilin coeluted with PP1α7–330, normalized to WT. The experiment was repeated twice. Error bars, s.d. (c) Overlay of the active site of spinophilin417–583–PP1α7–330–nodularin-R (orange) with PP1α7–330–nodularin-R (cyan; PDB 3E7A). (d) PP1α7–330 dephosphorylation of substrates in vitro: pNPP (green) GST-GluR1809–889 (blue) and phosphorylase a (red). Error bars, s.d. (n = 4). (e) Electrostatic surface representation (red, negative charge; blue, positive charge; gray, hydrophobic) of PP1α7–330 and spinophilin417–583–PP1α7–330 centered on the active site. (f) Mutation of PP1 Asp71 inhibits the dephosphorylation of phosphorylase a. PP1α7–330 WT and D71N dephosphorylation of GST-GluR1809–889 (blue) and phosphorylase a (red) in vitro. Error bars, s.d. (n = 4). (g) The PP1 D71N mutant reduces spinophilin binding. WT spinophilin binding to WT PP1 (blue) and PP1 D71N mutant (red) was tested. Error bars, s.d. (n = 2).

  6. Spinophilin affects the PP1 substrate binding surface without changing the active site.
    Figure 6: Spinophilin affects the PP1 substrate binding surface without changing the active site.

    (a) Model for substrate recognition by apo PP1. (b) Model for substrate recognition by the spinophilin–PP1 holoenzyme. The three PP1 substrate binding grooves are colored yellow. Potential substrates are shown as green, purple and black lines. Spinophilin achieves substrate specificity by steric exclusions of substrate binding sites.

Accession codes

Primary accessions

Protein Data Bank

Referenced accessions


Protein Data Bank


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Author information


  1. Department of Molecular Pharmacology, Physiology and Biotechnology, Brown University, Providence, Rhode Island, USA.

    • Michael J Ragusa,
    • Barbara Dancheck &
    • Wolfgang Peti
  2. Department of Molecular Biology, Cell Biology and Biochemistry, Brown University, Providence, Rhode Island, USA.

    • Michael J Ragusa,
    • David A Critton &
    • Rebecca Page
  3. Department of Psychiatry, Yale University School of Medicine, New Haven, Connecticut, USA.

    • Angus C Nairn


M.J.R. performed ITC, dephosphorylation assays and crystallization of the spinophilin–PP1, spinophilin–PP1–nodularin-R and neurabin–PP1 complexes; W.P. purified unbound spinophilin for NMR studies; B.D. performed and analyzed NMR studies of unbound spinophilin; M.J.R., D.A.C. and R.P. collected, processed and refined X-ray data; M.J.R., A.C.N., R.P. and W.P. wrote the paper; all authors discussed the data and manuscript.

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