Biochimica et Biophysica Acta (BBA) - Proteins and Proteomics
A dynamic loop provides dual control over the catalytic and membrane binding activity of a bacterial serine hydrolase
Graphical abstract
Introduction
Serine hydrolases (EC: 3.1) are a large class of ubiquitous and highly conserved cellular enzymes with broad substrate specificity and biological functions [1,2]. Acyl protein thioesterases (APTs) are one subgroup of serine hydrolases with well-defined biological functions [3,4]. In humans, two highly conserved APT homologues (APT1 and APT2) with overlapping enzyme activity are present with 66% sequence similarity [4,5]. Originally labeled as lysophospholipases and given the gene names LYPLA1 and LYPLA2, APT1 and APT2 have significantly higher catalytic efficiencies as depalmitoylases than as lysophospholipases [6,7]. In their role as depalmitoylases, APT1 and APT2 control reversible cycles of S-acylation with diverse signaling proteins, including various G-proteins, Ras isoforms, and ion channels [6,8,9]. Their depalmitoylase activity is regulated extracellularly through growth factor signaling [10], but how this extracellular signal is transduced into regulating the catalytic and membrane binding activities of APTs is still being investigated.
Although reversible cycles of palmitoylation are unknown in bacteria, structural and sequence homologues of human APTs exist across gram negative bacteria [[11], [12], [13]]. Previously, we determined the structure of FTT258, a bacterial APT homologue from F. tularensis, in comparison to the structure of APT1 [11]. The biological function of FTT258 is unknown, but bacterial pathogens, including Salmonella enterica, Legionella pneumophila, and Bacillus anthracis, take advantage of the mammalian palmitoylation machinery to rapidly target their own bacterial effectors to the host plasma membrane [[14], [15], [16]]. The overall three-dimensional structure of FTT258 showed that it had a highly similar fold to APT1 with a root mean square difference for the Cα atoms of 1.53 Å with similar patterns of membrane binding and catalytic activity [11,17,18]. Serendipitously, the structure of FTT258 also captured this enzyme in two different states, where the enzyme rearranged into closed and open conformations based on the movement of a dynamic loop, the β3-loop (Fig. 1A).
The structural rearrangement of this flexible β3-loop drastically reconfigured the structure of FTT258 and this rearrangement was predicted to control the biological functions of FTT258 and potentially other APT enzymes with similar structures and conserved β3-loops [11]. In addition to displacing the β3-loop from blocking the hydrophobic binding pocket of FTT258, the transition between the closed and open states also rearranged the catalytic His-Asp diad, removed an inhibitory aspartate (Asp25) from the binding site, and flipped out two proposed membrane binding residues (Trp66 and Tyr67) on the β3-loop. As a preliminary investigation of the biological importance of this structural rearrangement, these two proposed membrane binding residues were exchanged with alanine and changes in the catalytic and membrane binding activity of these FTT258 variants was measured [11]. Surprisingly, conversion of Trp66 or Tyr67 to alanine significantly decreased the catalytic activity of FTT258, but the membrane binding activity of FTT258 remained unchanged even with combinatorial substitution of both hydrophobic amino acids [11]. Similar dynamic loops with identical, central hydrophobic moieties are essential to the membrane binding activity of multiple human and bacterial proteins [[19], [20], [21], [22]].
Building on the surprising role for these two β3-loop residues in controlling the catalytic activity of FTT258, we have now broadly investigated the role of this dynamic loop in controlling the catalytic and membrane binding activities of FTT258. Using comprehensive mutagenesis across the β3-loop in combination with differential analysis of catalytic and membrane binding activity, we characterize distinct regions within the β3-loop responsible for each disparate function. We then used focused mutational analysis to investigate the molecular basis for β3-loop control over the catalytic and membrane binding activity of FTT258. Overall, we find that the β3-loop dually controls these two biological functions of FTT258 but through distinct subsections of the loop. These dual roles within a single dynamic loop provide a simple model for the interconnected regulation of these two essential biological functions in APTs.
Section snippets
Overexpression and purification of FTT258
FTT258 was purified similarly to a previous procedure [11]. Briefly, a bacterial plasmid (pDEST17-FTT258) was transformed into E. coli BL21 (DE3) RIPL cells (Agilent, La Jolla, CA). A saturated overnight culture of E. coli BL21 (DE3) RIPL (pDEST17-FTT258) in LB media containing ampicillin (200 μg/mL) and chloramphenicol (30 μg/mL) was used to inoculate LB-media (250 mL to 1.0 L depending on the FTT258 protein variant) containing ampicillin (200 μg/mL) and chloramphenicol (30 μg/mL) and the
Structural rearrangement of the β3-loop in FTT258
The x-ray crystal structure of FTT258 had eight subunits per unit cell and was crystallized in the presence of a weak, reversible, covalent cyclobutanone inhibitor [11,28]. In this heterogeneous mixture, four subunits of FTT258 were present in a closed conformation (Fig. 1A; orange) and four subunits in an open conformation (Fig. 1A; green) with one open subunit containing partial density for a cyclobutanone ligand [11]. The heterogeneity in the structural composition of FTT258 was proposed to
Conclusions
Dynamic protein loops provide a straightforward mechanism for the spatiotemporal control over interconnected protein activities [43,45,46]. For the bacterial serine hydrolase FTT258, a dynamic loop, previously identified serendipitously by X-ray crystallography [11], provides dual interconnected regulation over two biological functions. Based on movement of its β3-loop, FTT258 transitions from a closed substrate binding cleft with inactive catalytic arrangement to an open conformation with
Acknowledgements
We thank CH463 students from Butler University for assistance with preliminary enzyme purification and enzymatic analysis. We thank G. Hoops and M. Macbeth for providing constructive feedback on the manuscript. R.J.J. and M.A.S. were supported by a grant from the National Science Foundation (DUE-1140526). R.J.J. was also supported by a Senior research grant from the Indiana Academy of Sciences and by a Holcomb faculty research grant from Butler University.
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