Structure and Function of the Acetylpolyamine Amidohydrolase from

Article Cite This: Biochemistry XXXX, XXX, XXX−XXX

pubs.acs.org/biochemistry

Structure and Function of the Acetylpolyamine Amidohydrolase from the Deep Earth Halophile Marinobacter subterrani Jeremy D. Osko, Benjamin W. Roose, Stephen A. Shinsky,† and David W. Christianson* Roy and Diana Vagelos Laboratories, Department of Chemistry, University of Pennsylvania, 231 South 34th Street, Philadelphia, Pennsylvania 19104-6323, United States

Downloaded via MACQUARIE UNIV on August 29, 2019 at 16:22:44 (UTC). See https://pubs.acs.org/sharingguidelines for options on how to legitimately share published articles.

S Supporting Information *

ABSTRACT: Polyamines are small organic cations that are essential for cellular function in all kingdoms of life. Polyamine metabolism is regulated by enzyme-catalyzed acetylation− deacetylation cycles in a fashion similar to the epigenetic regulation of histone function in eukaryotes. Bacterial polyamine deacetylases are particularly intriguing, because these enzymes share the fold and function of eukaryotic histone deacetylases. Recently, acetylpolyamine amidohydrolase from the deep earth halophile Marinobacter subterrani (msAPAH) was described. This Zn2+-dependent deacetylase shares 53% amino acid sequence identity with the acetylpolyamine amidohydrolase from Mycoplana ramosa (mrAPAH) and 22% amino acid sequence identity with the catalytic domain of histone deacetylase 10 from Danio rerio (zebrafish; zHDAC10), the eukaryotic polyamine deacetylase. The X-ray crystal structure of msAPAH, determined in complexes with seven different inhibitors as well as the acetate coproduct, shows how the chemical strategy of Zn2+-dependent amide hydrolysis and the catalytic specificity for cationic polyamine substrates is conserved in a subterranean halophile. Structural comparisons with mrAPAH reveal that an array of aspartate and glutamate residues unique to msAPAH enable the binding of one or more Mg2+ ions in the active site and elsewhere on the protein surface. Notwithstanding these differences, activity assays with a panel of acetylpolyamine and acetyllysine substrates confirm that msAPAH is a broad-specificity polyamine deacetylase, much like mrAPAH. The broad substrate specificity contrasts with the narrow substrate specificity of zHDAC10, which is highly specific for N8-acetylspermidine hydrolysis. Notably, quaternary structural features govern the substrate specificity of msAPAH and mrAPAH, whereas tertiary structural features govern the substrate specificity of zHDAC10.

T

for function,22 whereas deacetylases require a single Zn2+ ion (or possibly a single Fe2+ ion) for function.23 The Zn2+ binding site in the deacetylases is conserved as the Mn2+B binding site of the arginases.19−21 Among the range of hydrolytic functions represented in the arginase−deacetylase phylogenetic tree, enzymes of polyamine biosynthesis are particularly intriguing. Arginase itself functions in polyamine biosynthesis to generate product L-ornithine, which undergoes decarboxylation to yield putrescine, a building block in the biosynthesis of spermidine and spermine.24−27 Generally present at millimolar concentrations in the cell, polyamines serve myriad biological functions such as the stabilization of nucleic acid structure28−30 and the regulation of transcription and translation.31−33 Notably, polyamines undergo reversible acetylation in cellular trafficking and function, and enzymes that catalyze polyamine deacetylation are structurally and functionally related to HDACs.17,18

he arginase−deacetylase superfamily of metalloenzymes has been extensively studied in recent years, but much of the phylogenetic tree remains uncharacterized in terms of structure and function.1−3 Rat arginase I was the first member of this superfamily to yield an X-ray crystal structure,4 and the subsequent crystal structure determination of the structurally homologous histone deacetylase-like protein from Aquifex aeolicus5 led to the designation of their common α/βmetallohydrolase topology as the arginase−deacetylase fold. Additional crystal structures of arginase-related metalloenzymes have since been reported, including proclavaminic acid amidino hydrolase,6 agmatinase,7 and formiminoglutamase.8 Among the deacetylases, crystal structures of class I and class II histone deacetylases (HDACs),3,9−16 as well as acetylpolyamine amidohydrolase from Mycoplana ramosa (mrAPAH),17,18 have been described. Structural comparisons indicate that the arginases and deacetylases divergently evolved from a common ancestral α/β-metallohydrolase despite sharing low amino acid sequence identities generally ranging from 10 to 15%.19−21 The stoichiometry of and selectivity for catalytic metal ion(s) have also diverged. Arginases generally require two Mn2+ ions © XXXX American Chemical Society

Received: July 8, 2019 Revised: August 19, 2019 Published: August 22, 2019 A

DOI: 10.1021/acs.biochem.9b00582 Biochemistry XXXX, XXX, XXX−XXX

Article

Biochemistry

Figure 1. (a) Polyamine deacetylase reaction catalyzed by msAPAH illustrated for substrate acetylputrescine. (b) Inhibitors studied in complex with msAPAH: 1, 5-[(3-aminopropyl)amino]pentylboronic acid; 2, 7-[(3-aminopropyl)amino]-1,1,1-trifluoroheptan-2-one; 3, 5-[(3aminopropyl)amino]pentane-1-thiol; 4, 6-amino-N-hydroxyhexanamide; 5, 8-amino-N-hydroxyoctanamide; 6, 6-[(3-aminopropyl)amino]-Nhydroxyhexanamide; 7, 4-(dimethylamino)-N-[7-(hydroxyamino)-7-oxoheptyl]benzamide (also known as M344). Inhibitor syntheses have been described previously.18,41

used without further purification. Inhibitory polyamine analogues 1−6 were synthesized as previously described.18,41 Inhibitor 7, 4-(dimethylamino)-N-[7-(hydroxyamino)-7oxoheptyl]benzamide (also known as M344), was purchased from Cayman Chemical. Protein Preparation. Full-length msAPAH was recombinantly expressed using a pGEX-6P-1 vector by Genscript. This construct utilizes a BamHI/XhoI cloning site and contains ampicillin bacterial resistance. An N-terminal GST tag is attached, followed by a PreScission protease cleavage site prior to the start of the protein sequence. Protein was expressed using Escherichia coli One Shot BL21(DE3) cells (Invitrogen) and grown in 2xYT medium in the presence of 100 μg/mL ampicillin. Cells were grown at 37 °C and 250 rpm in an Innova 40 incubator shaker until the OD600 reached approximately 0.80. The temperature was then decreased to 18 °C until the OD600 reached 1.0. At this point, cells were supplemented with 200 μM isopropyl β-L-1thiogalactopyranoside (IPTG) (Gold Biotechnology) and grown for an additional 18 h at 250 rpm. Cells were then centrifuged for 20 min at 5000 rpm using a Sorvall LYNX 6000 centrifuge. Cell pellets were stored at −80 °C until they were needed. The cell pellet was thawed and resuspended in 100 mL of buffer A [50 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) sodium salt (pH 7.5), 250 mM NaCl, 1 mM tris(2-carboxyethyl)phosphine (TCEP), 10% glycerol, and 10 μM ZnCl2]. Additionally, two protease inhibitor tablets, 0.1 mg/mL lysozyme, and 50 μg/mL DNase were added to the solution. The cells were then lysed by sonication. The cell lysate was centrifuged for 1 h at 15000 rpm using a Sorval LYNX 6000 centrifuge. The cell lysate was then applied to a 5 mL pre-equilibrated GSTrap HP column.

Indeed, HDAC10 was recently discovered to be a highly specific N8-acetylspermidine deacetylase.3,34 Although this eukaryotic cytosolic activity was discovered more than 40 years ago,35,36 the responsible enzyme was not identified at the time. In recent years, Marinobacter species have been identified in harsh marine and non-marine environments, including the deep subsurface based on their isolation in hydraulic fracturing effluent.37−39 These species are well adapted to the high salinity and Fe2+ concentrations found in mines, wells, and other deep subsurface locations. The halophile Marinobacter subterrani was recently identified 714 m below the surface in the Soudan iron mine in Minnesota,40 and genomic analysis reveals an acetylpolyamine amidohydrolase (msAPAH, UniProt A0A0J7JFD7) similar to mrAPAH (53% amino acid sequence identity). Both msAPAH and mrAPAH retain approximately 22% amino acid sequence identity with the polyamine deacetylase domain of HDAC10 from Danio rerio (zebrafish; zHDAC10). Here, we report the X-ray crystal structure of msAPAH and show that msAPAH is a broad-specificity polyamine deacetylase (Figure 1a). To illustrate the structural basis of substrate specificity and the catalytic mechanism, crystal structures of msAPAH complexed with the acetate product as well as seven inhibitors (Figure 1b) are presented. These studies provide important insight regarding the relationship between eukaryotic histone deacetylases and prokaryotic polyamine deacetylases.

■

MATERIALS AND METHODS Reagents. In general, all chemicals were purchased from Fisher Scientific, Millipore Sigma, or Hampton Research and B


Article

Biochemistry

mixture containing the msAPAH protein, acetylcadaverine substrate, and inhibitor. The reaction mixture was analyzed using a Tecan Infinite M1000Pro plate reader as described above. Assays were run at 25 °C, with the absorbance being measured at 450 nm for a total of 100 min. The IC50 values were calculated from raw data using Prism 7. Data were first transformed using the function x = log(x). Data were then normalized by setting the 0% identity definition to be the smallest mean in each data set, while the 100% identity definition was set to be the largest mean in each data set. Finally, the data were fit to a nonlinear regression curve of log(inhibitor) versus the normalized response. All measurements were run in duplicate. Goodness of fit statistics ranged from 88% to 97%. IC50 measurements are recorded in Figure S1. Isothermal Titration Calorimetry (ITC). Given the somewhat modest inhibitory potencies for some of the inhibitors studied, we additionally performed direct measurements of enzyme−inhibitor complexation using ITC. Dissociation constants (Kd) for inhibitors 1−7 with msAPAH were measured using a MicroCal iTC 200 isothermal titration calorimeter (GE Healthcare). For each measurement, 300 μM inhibitor was titrated into 30 μM msAPAH in 50 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM TCEP-HCl, and 10% (v/v) glycerol at 25 °C. In the case of inhibitor 6, 600 μM inhibitor was titrated into 60 μM msAPAH. For inhibitor 7, both inhibitor and msAPAH solutions included 5% (v/v) DMSO. Nineteen 2 μL injections were made over 60 min. Enthalpogram data were visualized and analyzed using Origin software (OriginLab, Northampton, MA). Enthalpograms for inhibitors 1, 2, 4, and 7 could not be fit to binding curves. Regardless, the direct calorimetric measurement of enzyme−inhibitor affinity for inhibitors 5 and 6 yielded micromolar dissociation constants, while inhibitor 3 yielded a nanomolar dissociation constant (Figure S2). Crystallization. All msAPAH−inhibitor complexes were crystallized using the sitting-drop vapor diffusion method. For cocrystallization of all msAPAH complexes, a 350 nL drop of the protein solution [10 mg/mL msAPAH in buffer C and 2 mM inhibitor in phosphate-buffered saline (PBS)] was added to a 350 nL drop of the precipitant solution as outlined below and equilibrated against 80 μL of the precipitant solution in the well reservoir. All crystals appeared within approximately 2 days at 4 °C, except those of the msAPAH−3 complex, which grew at 21 °C. Ethylene glycol [15% (v/v)] was added to the mother liquor as a cryoprotectant prior to the crystal being flash-cooled for X-ray diffraction data collection. For cocrystallization of the msAPAH−acetate complex, the precipitant consisted of 0.2 M calcium acetate hydrate and 20% (w/v) PEG 3350, which yielded thick cubic crystals. For cocrystallization of the msAPAH−1 complex, the precipitant consisted of 0.2 M magnesium chloride hexahydrate, 0.1 M HEPES (pH 7.5), and 25% (w/v) PEG 3350, which yielded long rod-like crystals. For cocrystallization of the msAPAH−2 complex, the precipitant consisted of 0.2 M magnesium acetate tetrahydrate and 20% (w/v) PEG 3350, which yielded thick plate-like crystals. For cocrystallization of the msAPAH−3 complex, the precipitant consisted of 20% (v/v) 2-propanol, 0.1 M MES monohydrate (pH 6.0), and 20% (w/v) polyethylene glycol monomethyl ether 2000, which yielded long rod-like crystals. For cocrystallization of the msAPAH−4 complex, the precipitant consisted of 0.2 M magnesium acetate tetrahydrate and 20% (w/v) PEG 3350, which yielded thick

The GST-msAPAH fusion protein bound to the GSTrap HP column and was eluted using buffer B [50 mM HEPES sodium salt (pH 8.0), 250 mM NaCl, 1 mM TCEP, 10% glycerol, 10 μM ZnCl2, and 10 mM reduced glutathione]. Proteincontaining fractions were collected and digested overnight using 5 mg/mL recombinant PreScission protease in 4 L of buffer A through dialysis (RC dialysis tubing, 6−8 kDa molecular weight cutoff). After protease cleavage for 12 h, the protein digest was applied to the 5 mL GSTrap HP column. The GST tag remained bound to the column, and the msAPAH protein flowed through the column. The msAPAH-containing fractions were concentrated to 5 mL using a 15 mL centrifugal filter unit with a molecular weight cutoff of 10 kDa. The protein was then filtered using a 0.22 μM Millex-GV filter unit prior to being loaded onto a HiLoad Superdex 26/600 200 pg column. The column was pre-equilibrated with 360 mL of buffer C [50 mM HEPES sodium salt (pH 7.5), 200 mM KCl, 1 mM TCEP, and 5% glycerol]. The 5 mL sample of msAPAH was injected at a rate of 1 mL/min, and 5 mL fractions were collected. Pure msAPAH protein was confirmed by sodium dodecyl sulfate− polyacrylamide gel electrophoresis, concentrated to approximately 10 mg/mL, and stored at −80 °C. Enzyme Kinetics. To measure the rate of acetylpolyamine hydrolysis catalyzed by msAPAH, the generation of the acetate coproduct was measured using a colorimetric assay kit (SigmaAldrich, catalog no. MAK086). Briefly, the reaction mixture (50 μL) was prepared as described in the kit protocol: 37 μL of Acetate Assay Buffer, 2 μL of Acetate Enzyme Mix, 2 μL of ATP, 2 μL of Acetate Substrate Mix, 2 μL of Probe, and 5 μL of 10 μM msAPAH. Separately, 50 μL substrate solutions were prepared at concentrations ranging from 0 to 5000 μM in Acetate Assay Buffer. To initiate the reaction, 50 μL of the reaction mixture was combined with 50 μL of the substrate solution in a Greiner 96well flat-bottom transparent plate, which was immediately placed in a Tecan Infinite M1000Pro plate reader and monitored at 450 nm. The pH of the assay solution was measured to be 7.75. Measurements were taken every 10 s for a total of 40 min, with shaking every 5 s. The plate was covered at all times due to the photosensitivity of the probe. To calculate the msAPAH activity, absorbance measurements at 450 nm were converted into product formation using an acetate standard curve according to the kit protocol. Data were analyzed using GraphPad Prism version 7.00 for MAC OS X (GraphPad software, La Jolla CA, www.graphpad.com). Assays were performed in triplicate at 25 °C. Nonlinear regression fits to the Michaelis−Menten equation were used to determine steady-state parameters. Inhibitory Activity Measurements. Inhibitory potencies of compounds 1−7 against msAPAH were determined using the colorimetric acetate assay described above. Briefly, the reaction mixture (50 μL) was prepared as described above, except that the protein was omitted: 42 μL of Acetate Assay Buffer, 2 μL of Acetate Enzyme Mix, 2 μL of ATP, 2 μL of Acetate Substrate Mix, and 2 μL of Probe. Separately, 20 μL of 2 μM msAPAH protein was prepared with 10 μL of inhibitor (prepared in buffer C) with final inhibitor concentrations ranging from 0 to 2000 μM. The inhibitor was incubated with enzyme for 15 min prior to the addition of substrate. The reaction was initiated by the addition of 20 μL of 4000 μM acetylcadaverine substrate. Immediately after substrate addition, 50 μL of reaction mixture was added to the 50 μL of C


D

5332 44 354 11 21 14

0.007 0.9

97.0 3.00 0.00 6PHZ

10580 69 598

12 15 15

0.007 0.9

96.0 4.00 0.00 6PHT

89.79−2.00 (2.07−2.00) 41498 (4163) 0.161/0.212 (0.158/0.227)

47.4, 82.4, 90.7 90, 98, 90 0.180 (0.597) 0.180 (0.595) 0.952 (0.525) 3.4 (3.5) 90.2 (91.4) 10.2 (6.1)

65.6, 163.5, 65.7 90, 94, 90 0.161 (0.456) 0.102 (0.312) 0.958 (0.706) 3.6 (3.6) 98.9 (99.7) 4.7 (2.8)

65.44−1.65 (1.68−1.65) 125734 (12518) 0.193/0.225 (0.216/0.264)

P21

P21

msAPAH−2

96.0 4.00 0.00 6PHR

0.008 0.9

15 19 19

5325 54 294

61.77−1.65 (1.71−1.65) 79658 (7965) 0.179/0.208 (0.254/0.284)

45.9, 120.7, 65.4 90, 109, 90 0.163 (1.018) 0.106 (0.675) 0.991 (0.657) 6.2 (6.1) 99.1 (99.2) 6.7 (2.1)

P21

msAPAH−3

97.0 3.00 0.00 6PIC

0.004 0.7

10 8 9

21126 112 648

66.3, 67.0, 167.5 90, 90, 93 0.205 (0.474) 0.158 (0.403) 0.830 (0.588) 1.9 (1.9) 83.2 (86.5) 4.4 (2.2) Refinement 66.97−2.03 (2.10−2.03) 154543 (15944) 0.220/0.247 (0.278/0.315)

P1

msAPAH−4 Data Collection

97.0 3.00 0.00 6PID

0.006 0.8

12 16 14

5343 33 254

59.53−1.54 (1.60−1.55) 95821 (7699) 0.244/0.280 (0.415/0.415)

52.3, 119.1, 66.2 90, 109, 90 0.172 (0.742) 0.172 (0.742) 0.969 (0.537) 3.4 (3.4) 86.7 (88.7) 5.6 (2.1)

P21

msAPAH−5

96.0 4.00 0.00 6PIA

0.006 0.8

17 20 21

10689 85 774

46.27−1.75 (1.81−1.75) 131376 (12851) 0.175/0.212 (0.242/0.293)

65.7, 163.2, 65.7 90, 94, 90 0.092 (0.531) 0.085 (0.470) 0.992 (0.657) 3.2 (3.0) 95.0 (95.6) 7.3 (1.9)

P21

msAPAH−6

96.0 4.00 0.00 6PI1

0.006 0.8

13 24 20

10673 105 953

60.71−1.70 (1.76−1.70) 136205 (13035) 0.162/0.199 (0.202/0.247)

65.6, 163.0, 65.6 90, 95, 90 0.105 (0.370) 0.100 (0.353) 0.986 (0.811) 3.4 (3.1) 90.8 (81.8) 6.8 (2.5)

P21

msAPAH−7

97.0 3.00 0.00 6PI8

0.006 0.9

19 19 26

5386 25 502

27.85−1.64 (1.69−1.64) 95961 (9560) 0.168/0.194 (0.277/0.333)

52.4, 121.1, 66.6 90, 109, 90 0.104 (0.826) 0.066 (0.531) 0.997 (0.703) 6.7 (6.6) 99.9 (99.7) 10.2 (2.1)

P21

msAPAH−acetate

a Values in parentheses refer to the highest-resolution shell indicated. bRmerge = ∑hkl∑i|Ii,hkl − ⟨I⟩hkl|/∑hkl∑iIi,hkl, where ⟨I⟩hkl is the average intensity calculated for reflection hkl from replicate measurements. cRpim = {∑hkl[1/(N − 1)]1/2∑i|Ii,hkl − ⟨I⟩hkl|}/∑hkl∑iIi,hkl, where ⟨I⟩hkl is the average intensity calculated for reflection hkl from replicate measurements and N is the number of reflections. d Pearson correlation coefficient between random half-data sets. eRwork = ∑||Fo| − |Fc||/∑|Fo| for reflections contained in the working set. |Fo| and |Fc| are the observed and calculated structure factor amplitudes, respectively. Rfree is calculated using the same expression for reflections contained in the test set held aside during refinement. fPer asymmetric unit. gCalculated with PROCHECK.

no. of atomsf protein ligand solvent average B factor (Å2) protein ligand solvent root-mean-square deviation bond lengths (Å) bond angles (deg) Ramachandran plotg (%) favored allowed outliers PDB entry

no. of reflections Rwork/Rfreee

resolution (Å)

space group unit cell dimensionsa a, b, c (Å) α, β, γ (deg) Rmergeb Rpimc CC1/2d redundancy completeness (%) I/σ

msAPAH−1

Table 1. Crystallographic Data Collection and Refinement Statistics

Biochemistry Article


Article

Biochemistry

unable to determine steady-state kinetic parameters in higher concentrations of KCl. No significant catalytic activity was observed for the hydrolysis of Aly-NH2 or the MSH2-based peptide substrates Ac-Ala-Aly, Ac-Gly-Ala-Aly, Ac-Gly-Ala-AlyAsn-Leu-Gln-NH2, and Ac-Aly-Asn-Leu-NH2 (Aly = acetyllysine) using the liquid chromatography−mass spectrometry assay employed for the assay of HDAC6 activity.16 Thus, while msAPAH is not a particularly efficient polyamine deacetylase, it is clearly not a lysine deacetylase. Structure of the msAPAH−1 Complex. The 1.65 Å resolution crystal structure of msAPAH complexed with the boronic acid analogue of N8-acetylspermidine (1; IC50 = 390 μM) contains four monomers in the asymmetric unit. The structure of msAPAH reveals the arginase−deacetylase fold, as previously observed for the broad-specificity polyamine deacetylase mrAPAH.17 Like mrAPAH, msAPAH is a dimer with an identical quaternary structure (Figure 3). There are no major conformational changes between the monomers or dimers of msAPAH and mrAPAH (PDB entry 4ZUM). The root-mean-square deviation (rmsd) is 0.53 Å for 268 Cα atoms (monomer A) and 0.75 Å for 566 Cα atoms for the dimer. Electron density corresponding to the boronic acid moiety of 1 clearly indicates trigonal planar geometry (Figure 4), with the hydroxyl groups coordinated to the catalytic Zn2+ ion with asymmetric bidentate coordination geometry (average Zn2+− O1 distance of 2.1 Å and Zn2+−O2 distance of 2.4 Å). Notably, inhibitor binding displaces the Zn2+-bound solvent molecule expected for the unliganded enzyme. Surprisingly, the electrophilic boronic acid does not undergo nucleophilic attack to form a tetrahedral boronate anion, which would mimic the tetrahedral transition state for acetylpolyamine hydrolysis. Such chemistry is typically observed, for example, in the binding of boronic acids in the arginase active site,49−52 but not exclusively so.53 Boronic acid hydroxyl group O1 hydrogen bonds with Y323, and hydroxyl group O2 hydrogen bonds with H158 and H159. These catalytic residues are unique to metal-dependent deacetylases. In catalysis, Y323 assists the Zn2+ ion in polarizing the amide carbonyl group of the substrate for nucleophilic attack by a Zn2+-bound solvent molecule, and tandem histidine residues H158 and H159 serve general base and general acid functions. Additional hydrogen bond interactions stabilize the binding of 1 in the msAPAH active site. The primary amino group of 1 forms water-mediated hydrogen bonds with the side chain of E17, the backbone carbonyl of L18, and the side chain of D19. The binding conformation of 1 is slightly different in monomer B, such that there is a direct hydrogen bond with the side chain of Y168. The secondary amino group of 1 forms watermediated hydrogen bonds with the side chains of D117 and Y168 and the primary amino group of 1. Although the quaternary structure of msAPAH is identical to that of mrAPAH, there are several key differences in residues that define the active site cleft at the dimer interface of msAPAH. Residue Y19 in mrAPAH corresponds to D19 in msAPAH, a residue that is critical for accepting hydrogen bonds from the polyamine substrates. Residue F27 in mrAPAH corresponds to H27 in msAPAH and is located at the surface of the active site pocket. The most noticeable difference at the dimer interface is the substitution of residues Y83, S91, and E106 in mrAPAH with K83, F91 and D106, respectively, in msAPAH. This difference enables the binding of a Mg2+ ion at the dimer interface of msAPAH. This Mg2+ ion is observed in all msAPAH crystal structures described herein and is

plate-like crystals. For cocrystallization of the msAPAH−5 complex, the precipitant consisted of 0.2 M magnesium acetate tetrahydrate and 20% (w/v) PEG 3350, which yielded thick plate-like crystals. For cocrystallization of the msAPAH−6 complex, the precipitant consisted of 0.2 M calcium chloride dihydrate and 20% (w/v) PEG 3350, which yielded thick plate-like crystals. For cocrystallization of the msAPAH−7 complex, the precipitant consisted of 0.2 M calcium chloride dihydrate and 20% (w/v) PEG 3350, which yielded thick plate-like crystals. Data Collection and Structure Determination. X-ray diffraction data were collected at Northeastern Collaborative Access Team beamline 24-ID-E, Advanced Photon Source, Argonne National Laboratory, from crystals of msAPAH complexed with 1 and 3. X-ray diffraction data were collected at Northeastern Collaborative Access Team beamline 24-ID-C, Advanced Photon Source, Argonne National Laboratory, from crystals of msAPAH complexed with 2, 5, and 7. X-ray diffraction data were collected at beamline 12-2, Stanford Synchrotron Radiation Laboratory, Stanford University, from crystals of msAPAH complexed with 4 and 6. X-ray diffraction data were collected at Frontier Macromolecular Crystallography beamline 17-ID-2 (FMX), National Synchrotron Light Source II, Brookhaven National Laboratory, from crystals of msAPAH complexed with acetate. Data were indexed and integrated using iMosflm42 and scaled using Aimless in the CCP4 program suite.43 Data reduction statistics are listed in Table 1. Molecular replacement using the atomic coordinates of mrAPAH [Protein Data Bank (PDB) entry 4ZUM]18 was used as a search model to phase the initial electron density map of the first msAPAH structure. Subsequent msAPAH structures were determined in a similar fashion using either mrAPAH or msAPAH as a search probe for molecular replacement. Autobuild44 was used to register the msAPAH sequence in the initial electron density map. The graphics program COOT45 was used to manually adjust residue conformations, and structural refinement was performed using PHENIX.46 The quality of each structure was assessed using MolProbity47 and PROCHECK.48 Refinement statistics are listed in Table 1.

■

RESULTS Catalytic Activity. Recombinant msAPAH exhibits broad specificity for short and long acetylpolyamine substrates, with a moderate preference for putrescine or N8-acetylspermidine (depending on the concentration of KCl) based on catalytic efficiency (kcat/KM) (Figure 2 and Table 2). Even so, catalytic efficiency is modest; at best, kcat/KM = 470 M−1 s−1 for N8acetylspermidine in the presence of 10 mM KCl. We were

Figure 2. Michaelis−Menten plot showing the steady-state kinetics of msAPAH-catalyzed acetylpolyamine hydrolysis. E


Article

Biochemistry Table 2. Steady-State Kinetic Parameters for Acetylpolyamine Hydrolysis by msAPAHa

All measurements represent means ± the average of absolute deviations from the mean in a given set of data. All measurements were taken in triplicate at 25 °C.

a

absent in monomer A, where Y323 adopts the “out” conformation. The gem-diol O2 group hydrogen bonds with H158 and H159. The binding of trifluoroketone inhibitors as gem-diols has similarly been observed in the active sites of other deacetylases.3,16,18 Each of the C−F bonds of the trifluoro moiety of 2 engages in hydrogen bond interactions with protein atoms. One fluorine atom accepts a hydrogen bond from Y323, and a second fluorine atom accepts a hydrogen bond from the backbone NH group of G321. A third fluorine atom accepts a hydrogen bond from C169; this fluorine atom also makes a van der Waals contact with the backbone carbonyl of G167. The primary amino group of 2 donates hydrogen bonds to the side chain of E17, the side chain of D19, and the backbone carbonyl oxygen of L18. The secondary amino group makes water-mediated hydrogen bond interactions with the side chains of D117 (monomer B) or Y168 (monomer A). In addition to the Mg2+ ion bound in the active site, magnesium hexahydrate [Mg2+(OH2)6] binds near D275 in monomer A, and the side chain carboxylate of D275 makes hydrogen bond interactions with metal-bound water molecules. In monomer B, E288 coordinates to a Mg2+ ion along with five water molecules, yielding an octahedral metal coordination geometry. Structure of the msAPAH−Acetate Complex. The 1.64 Å resolution crystal structure of msAPAH complexed with the acetate coproduct contains two monomers in the asymmetric

coordinated by the carboxylate side chains of D104 and D106; the Mg2+ coordination polyhedron is completed by water molecules. Additional residues that are important for dimer assembly of mrAPAH include F92, which engages in offset π stacking interactions with F92 in the adjacent monomer. In msAPAH, this interaction is maintained by the W92−W92 link at the dimer interface. In addition to the Mg2+ ion bound in the active site at the dimer interface, an additional Mg2+ ion is coordinated by E288 in monomer A. The coordination sphere of this Mg2+ ion is completed by four water molecules, yielding a pentacoordinate metal ion. This Mg2+ ion is absent in monomers B−D. Structure of the msAPAH−2 Complex. The 2.00 Å resolution crystal structure of msAPAH complexed with the trifluoroketone analogue of N8-acetylspermidine (2; IC50 = 350 μM) contains two monomers in the asymmetric unit. The trifluoroketone moiety exists predominantly as the gem-diol in solution;54 accordingly, it binds as the gem-diol or perhaps the ionized gem-diolate in the active site of msAPAH (Figure 5). This binding mode mimics the binding of the tetrahedral intermediate and its flanking transition states in the hydrolysis of an acetylpolyamine substrate. The gem-diol(ate) coordinates to the catalytic Zn2+ ion with nearly symmetric bidentate coordination geometry, with average Zn2+−O1 and Zn2+−O2 distances of 2.2 and 2.3 Å, respectively. The gemdiol O1 group also hydrogen bonds with Y323, which adopts the “in” conformation in monomer B. This hydrogen bond is F


Article

Biochemistry

Figure 5. Polder omit map of the msAPAH−2 complex, monomer B (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH monomer B), dark gray (msAPAH monomer A), or wheat (inhibitor), F in magenta, S in yellow, N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

Figure 3. Crystal structure of the msAPAH−1 complex (top) illustrating the dimer interface between monomer A (blue) and monomer B (red), looking down the 2-fold symmetry axis. There are no major conformational differences between monomers, and leastsquares superposition yields an rmsd of 0.15 Å for 300 Cα atoms. The cutaway view of the active site (bottom) shows how the dimer interface constricts the approach to the catalytic Zn2+ ions (gray spheres). Only long and slender acetylpolyamine substrates, and not bulky acetyllysine-containing peptide substrates, can bind in this constricted active site.

Figure 6. Polder omit map of the msAPAH−acetate complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH monomer A), dark gray (msAPAH monomer B), wheat (inhibitor), or orange (ethylene glycol), N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

In addition to the Mg2+ ion bound in the active site, an additional Mg2+ ion is coordinated by E288, D324, D326, and three water molecules in monomer A. However, this Mg2+ ion is absent in monomer B. Structure of the msAPAH−3 Complex. The 1.65 Å resolution crystal structure of msAPAH complexed with the thiol analogue of N8-acetylspermidine (3; IC50 = 160 μM; Kd = 0.2 μM) contains two monomers in the asymmetric unit. The thiol group is presumably ionized to the negatively charged thiolate anion as it coordinates to the active site Zn2+ ion, with an average Zn2+−S distance of 2.4 Å (Figure 7). With Zn2+··· S−C angles of 116° and 100° and Zn2+···S−C−C dihedral angles of −7° in monomer A and 59° in monomer B, thiolate− metal coordination parameters deviate somewhat from ideal values for thiolate−metal interactions first outlined for the side chain of cysteine (Zn2+···S−C angle of 90° and Zn2+···S−C−C dihedral angle of ±90° or ±180°).55 The phenolic hydroxyl group of Y323 additionally donates a hydrogen bond to the Zn2+-bound thiolate. As observed in the msAPAH−2 complex, the primary amino group of 3 hydrogen bonds to E17, D19, and the backbone carbonyl of L18. Also similar are the hydrogen bonds between the secondary amino group of 3 and D117 and Y168.

Figure 4. Polder omit map of the msAPAH−1 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH monomer D), dark gray (msAPAH monomer B), or wheat (inhibitor), B in light green, N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

unit. Absent the polyamine coproduct bound in the active site, this structure represents a partially dissociated product complex. Acetate coordinates to the active site Zn2+ ion with nearly symmetric bidentate geometry (average Zn2+−O1 and Zn2+−O2 distances of 2.2 and 2.4 Å, respectively) (Figure 6). The carboxylate O1 atom additionally accepts a hydrogen bond from Y323, and the carboxylate O2 atom accepts hydrogen bonds from H158 and H159. G


Article

Biochemistry

(Figure 7). The primary amino group of 4 also engages in water-mediated hydrogen bonds with E17, D19, and the backbone carbonyl of L18. Structure of the msAPAH−5 Complex. The 1.55 Å resolution crystal structure of msAPAH complexed with hydroxamate inhibitor 5 (IC50 = 380 μM; Kd = 29 μM) contains two monomers in the asymmetric unit. Hydroxamate−metal coordination interactions and hydrogen bond interactions (Figure 9) are identical to those observed for the

Figure 7. Polder omit map of the msAPAH−3 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH, monomer B), dark gray (msAPAH, monomer A), or wheat (inhibitor and MES), S in yellow, N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

Unique to this structure is the binding of a buffer molecule, 2-(N-morpholino)ethanesulfonic acid (MES), in the active site tunnel (Figure 7). The sulfonate group of MES accepts a hydrogen bond from the backbone NH group of F225 and a Mg2+-bound water molecule. It also makes a hydrogen bond with H197. Structure of the msAPAH−4 Complex. The 2.03 Å resolution crystal structure of msAPAH complexed with hydroxamate inhibitor 4 (IC50 = 410 μM) contains eight monomers in the asymmetric unit. Inhibitor 4 is a hydroxamate analogue of N-acetylputrescine. Intermolecular interactions of the hydroxamate group provide useful structural inferences about catalysis (Figure 8). Specifically, just as the

Figure 9. Polder omit map of the msAPAH−5 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH, monomer A), dark gray (msAPAH, monomer B), or wheat (inhibitor), N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

binding of hydroxamate inhibitor 4. While inhibitor 5 is not isosteric with any naturally occurring polyamine substrate, its intermolecular interactions reveal that alternative hydrogen bond interactions can be achieved for the primary amino group of a long, slender polyamine analogue in the active site of msAPAH. Inhibitor 5 is two methylene groups longer than inhibitor 4 and accordingly extends farther out of the active site tunnel. The additional length allows for the primary amino group to engage in direct hydrogen bonds rather than watermediated hydrogen bonds with E17, D19, and the backbone carbonyl of L18. The D19 hydrogen bond occurs only in monomer A, while a hydrogen bond to Y168 occurs in monomer B. In addition to the Mg2+ ion bound in the active site, an additional Mg2+ ion is coordinated by E288, D324, D326, and three water molecules in monomer A. However, this Mg2+ ion is absent in monomer B. Structure of the msAPAH−6 Complex. The 1.75 Å resolution crystal structure of the complex between msAPAH and hydroxamate inhibitor 6 (IC50 = 470 μM; Kd = 11 μM) contains four monomers in the asymmetric unit. Hydroxamate−metal coordination interactions and hydrogen bond interactions (Figure 10) are identical to those observed for other hydroxamate inhibitors described above. Inhibitor 6 is the hydroxamate analogue of the polyamine substrate N8acetylspermidine. The primary amino group of 6 donates hydrogen bonds to E17, D19, and the backbone carbonyl of L18. The secondary amino group of 6 donates hydrogen bonds to D117 in monomers A and B and Y168 in all four monomers. In addition to the Mg2+ ion bound in the active site, an additional Mg2+ ion is coordinated by E288 and five water molecules in monomer B; D324 and D326 form hydrogen bonds with metal-bound water molecules. However, this Mg2+ ion is absent in monomers A, C, and D.

Figure 8. Polder omit map of the msAPAH−4 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH, monomer A), dark gray (msAPAH, monomer C, symmetry mate), or wheat (inhibitor), N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

hydroxamate carbonyl group coordinates to Zn2+ and accepts a hydrogen bond from Y323, so too would the scissile carbonyl group of N-acetylputrescine in the precatalytic enzyme− substrate complex. Both Zn2+ coordination and Y323 hydrogen bond interactions are required to activate the amide carbonyl of N-acetylputrescine for nucleophilic attack by a Zn2+-bound solvent molecule in catalysis. The primary amino group of inhibitor 4 donates hydrogen bonds to D117 and Y168 (Figure 8), in a manner similar to the interactions of the secondary amino group of inhibitor 3 H


Article

Biochemistry

polyamine deacetylase. Common to all metal-dependent deacetylases in the arginase−deacetylase superfamily, catalytically important residues in the msAPAH active site include tandem histidine residues H158 and H159. We speculate that H158 serves as a general base in catalysis by deprotonating a Zn2+-bound water molecule for nucleophilic attack at the scissile amide carbonyl of the substrate, which in turn is polarized by coordination to Zn2+ and also by a hydrogen bond with Y323. This mechanistic proposal is consistent with intermolecular interactions observed for the binding of trifluoroketone transition-state analogue 2 (Figure 5) as well as the acetate coproduct (Figure 6). On the basis of the observation of two conformations for Y323 in the msAPAH−2 complex (the “in” conformation oriented into the active site and the “out” conformation oriented toward the bulk solvent), it appears likely that Y323 undergoes a substrate-induced conformational change in catalysis. Conformational flexibility of the catalytic tyrosine has also been observed in crystal structures of mrAPAH,17,18 as well as molecular dynamics simulations of HDAC3 and HDAC8.56−58 This flexibility appears to be enhanced by the location of Y323 in a glycinerich loop.58 Molecular recognition of cationic polyamine substrates by msAPAH and mrAPAH is facilitated by substantial negative electrostatic potential on the protein surface surrounding the mouth of the active site (Figure 12). The negatively charged protein surface provides electrostatic attraction to facilitate the binding of positively charged polyamine substrates. Once bound in the active site of msAPAH, the backbone carbonyl of L18 and the side chains of E17, D19, D117, and Y168 provide direct or water-mediated hydrogen bond interactions to the positively charged amino groups of the substrate, as observed in the array of crystal structures presented in Figures 4−11. Unique to msAPAH is the binding of a Mg2+ ion in the substrate binding cleft in the structures of all enzyme−inhibitor complexes. This metal ion is coordinated by D104 and D106 of the alternate subunit of the dimer, and water molecules usually complete an octahedral coordination complex. The binding of this Mg2+ ion further constricts the active site and likely contributes to the catalytic preference for acetylpolyamine substrates. The Mg2+ ligand D106 is not conserved in mrAPAH, and Mg2+ binding has not been observed in this related bacterial deacetylase. The binding of additional Mg2+ ions as observed in all msAPAH structures may reflect the functional adaptation of this deacetylase as M. subterrani evolved in the high-salt subterranean environment. A characteristic feature of halophilic proteins is an increase in the number of acidic residues on the protein surface.59−62 Accordingly, msAPAH has increased aspartate and glutamate content (15% total) compared with mrAPAH (12% total). Increased negative charge accounts for the lower pI of msAPAH compared with that of mrAPAH. On the basis of pI values calculated from their amino acid sequences, the pI of msAPAH is 5.0 and the pI of mrAPAH is 5.3. Additional aspartate residues present in msAPAH also account for additional Mg2+ binding sites observed in the crystal structures of complexes with different inhibitors. The structural features described above likely contribute to the substrate specificity of mrAPAH and msAPAH. Our previous steady-state kinetic studies utilizing a liquid chromatography−mass spectrometry assay for the direct measurement of dansylated polyamine products3 show that mrAPAH exhibits broad substrate specificity and catalyzes the

Figure 10. Polder omit map of the msAPAH−6 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH, monomer A), dark gray (msAPAH, monomer B), or wheat (inhibitor), N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

Structure of the msAPAH−7 Complex. The 1.70 Å resolution crystal structure of msAPAH complexed with inhibitor 7 (IC50 = 160 μM) contains four monomers in the asymmetric unit. Inhibitor 7 is also known as M344 and is characterized by a m-dimethylaminobenzamide “capping group”. Hydroxamate−metal coordination interactions and hydrogen bond interactions (Figure 11) are identical to those

Figure 11. Polder omit map of the msAPAH−7 complex (contoured at 3.0σ). Atoms are color-coded as follows: C in light blue (msAPAH, monomer B), dark gray (msAPAH, monomer A), or wheat (inhibitor), N in blue, O in red, Zn2+ as a gray sphere, Mg2+ as a large, dark red sphere, and solvent as small red spheres. Metal coordination and hydrogen bond interactions are indicated by solid and dashed black lines, respectively.

observed for other hydroxamate inhibitors described above. The aromatic capping group is nestled in the dimer interface and is oriented away from the Mg2+ ion. The amide group forms water-mediated hydrogen bonds with H197, the backbone amide of F225, the backbone carbonyl of I291, and a Mg2+-bound water molecule. The amide group also forms water-mediated hydrogen bonds with E17, D117, and Y168. In addition to the Mg2+ ion bound in the active site, an additional Mg2+ ion is coordinated by E288 and five water molecules in monomer B. However, this Mg2+ ion is absent in monomers A, C, and D.

■

DISCUSSION Crystal structures of msAPAH−inhibitor complexes reveal key insights into substrate recognition and catalysis by a bacterial I


Article

Biochemistry

first observed in partially purified cell extracts more than 40 years ago, the responsible enzyme had not been identified.35,36 It is interesting to contrast the structural basis of the catalytic specificity in the bacterial polyamine deacetylases msAPAH and mrAPAH with that of HDAC10.34 This specificity is rooted in structural and electrostatic features in each enzyme active site. First, each enzyme active site is sterically constricted to favor the binding of long, slender polyamine substrates over bulky, acetyllysine-containing peptide substrates. However, the active sites of the bacterial enzymes are constricted by quaternary structure (Figure 3), whereas the active site of the eukaryotic enzyme is constricted by tertiary structure: a 310 helix unique to HDAC10 and no other HDAC isozymes.3,34 Second, each enzyme is characterized by significant negative electrostatic potential on the protein surface flanking the active site (Figure 12), as well as specific aspartate and/or glutamate residues in the active site cleft that engage in electrostatic interactions with the positively charged amino groups of polyamine substrates. As noted by Hai and colleagues,3 polyamine substrate specificity in HDAC10 and APAH enzymes appears to have evolved convergently. In closing, it is interesting to consider the biology and ecology of the halophile M. subterrani from which msAPAH derives. As noted in the introductory section, these species are well adapted to the high Fe2+ concentrations found in deep subsurface locations.40 Given that HDAC enzymes can utilize Fe2+ for catalytic activity slightly exceeding that measured with Zn2+, and the implication that metal-dependent deacetylases could utilize Fe2+, Zn2+, or perhaps a mixture of both in vivo,23 it is interesting to speculate that msAPAH could potentially function as an Fe2+-dependent polyamine deacetylase in the iron-rich environment of the deep terrestrial biosphere. Future structure−function studies of msAPAH will shed further light on this possibility.

■

ASSOCIATED CONTENT

S Supporting Information *

Figure 12. Electrostatic surfaces of msAPAH, mrAPAH, and zHDAC10. Arrows indicate the mouth of each enzyme active site.

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biochem.9b00582.

hydrolysis of N-acetylcadaverine, N-acetylputrescine, N1acetylspermidine, N8-acetylspermidine, and N-acetylspermine with turnover numbers ranging from 0.20 to 2.5 s−1 and catalytic efficiencies ranging from 2.5 × 103 to 6.8 × 104 M−1 s−1. These results are consistent with previous observations of broad substrate specificity based on specific activity measurements using native and recombinant mrAPAH.63,64 Similarly, msAPAH exhibits broad substrate specificity but lower catalytic activity using an assay that quantitates the generation of coproduct acetate. Turnover numbers range from 0.0038 to 0.026 s−1, and catalytic efficiencies range from 5 to 470 M−1 s−1. We cannot rule out the possibility that msAPAH assay conditions do not fully reflect the cytosolic conditions of the deep earth halophile from which msAPAH derives, nor can we rule out the possibility that the biological substrate of msAPAH in vivo is an as yet unidentified acetylated small molecule. The biological substrate is unlikely to be acetyllysine, because a peptide or protein substrate would not fit in the constricted approach to the active site (Figure 3). Recently, eukaryotic HDAC10 was discovered to be a highly specific cytosolic polyamine deacetylase that utilizes N8acetylspermidine as a substrate.3 Although this activity was

IC50 plots (Figure S1) and ITC enthalpograms (Figure S2) (PDF) Accession Codes

The atomic coordinates and crystallographic structure factors of msAPAH complexes with the inhibitors shown in Figure 1b have been deposited in the Protein Data Bank (www.rcsb.org): 1, 6PHT; 2, 6PHZ; 3, 6PHR; 4, 6PIC; 5, 6PID; 6, 6PIA; 7, 6PI1; acetate, 6PI8.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

David W. Christianson: 0000-0002-0194-5212 Present Address †

S.A.S.: Department of Biology, The College of New Jersey, 2000 Pennington Rd., Ewing, NJ 08618. Funding

The authors thank the National Institutes of Health for Grants GM49758 (D.W.C.) and F32GM125141 (S.A.S.). J


Article

Biochemistry Notes

W. (2004) Structural snapshots of human HDAC8 provide insights into the class I histone deacetylases. Structure 12, 1325−1334. (10) Vannini, A., Volpari, C., Filocamo, G., Casavola, E. C., Brunetti, M., Renzoni, D., Chakravarty, P., Paolini, C., De Francesco, R., Gallinari, P., Steinkuhler, C., and Di Marco, S. (2004) Crystal structure of a eukaryotic zinc-dependent histone deacetylase, human HDAC8, complexed with a hydroxamic acid inhibitor. Proc. Natl. Acad. Sci. U. S. A. 101, 15064−15069. (11) Bottomley, M. J., Lo Surdo, P., Di Giovine, P., Cirillo, A., Scarpelli, R., Ferrigno, F., Jones, P., Neddermann, P., De Francesco, R., Steinkuhler, C., Gallinari, P., and Carfi, A. (2008) Structural and functional analysis of the human Hdac4 catalytic domain reveals a regulatory zinc-binding domain. J. Biol. Chem. 283, 26694−26704. (12) Schuetz, A., Min, J., Allali-Hassani, A., Schapira, M., Shuen, M., Loppnau, P., Mazitschek, R., Kwiatkowski, N. P., Lewis, T. A., Maglathin, R. L., McLean, T. H., Bochkarev, A., Plotnikov, A. N., Vedadi, M., and Arrowsmith, C. H. (2008) Human HDAC7 harbors a class IIa histone deacetylase-specific zinc binding motif and cryptic deacetylase activity. J. Biol. Chem. 283, 11355−11363. (13) Bressi, J. C., Jennings, A. J., Skene, R., Wu, Y., Melkus, R., De Jong, R., O’Connell, S., Grimshaw, C. E., Navre, M., and Gangloff, A. R. (2010) Exploration of the HDAC2 foot pocket: Synthesis and SAR of substituted N-(2-aminophenyl)Benzamides. Bioorg. Med. Chem. Lett. 20, 3142−3145. (14) Watson, P. J., Fairall, L., Santos, G. M., and Schwabe, J. W. R. (2012) Structure of Hdac3 bound to co-repressor and inositol tetraphosphate. Nature 481, 335−340. (15) Millard, C. J., Watson, P. J., Celardo, I., Gordiyenko, Y., Cowley, S. M., Robinson, C. V., Fairall, L., and Schwabe, J. W. (2013) Class I HDACs share a common mechanism of regulation by inositol phosphates. Mol. Cell 51, 57−67. (16) Hai, Y., and Christianson, D. W. (2016) Histone deacetylase 6 structure and molecular basis of catalysis and inhibition. Nat. Chem. Biol. 12, 741−747. (17) Lombardi, P. M., Angell, H. D., Whittington, D. A., Flynn, E. F., Rajashankar, K. R., and Christianson, D. W. (2011) Structure of prokaryotic polyamine deacetylase reveals evolutionary functional relationships with eukaryotic histone deacetylases. Biochemistry 50, 1808−1817. (18) Decroos, C., and Christianson, D. W. (2015) Design, synthesis, and evaluation of polyamine deacetylase inhibitors, and highresolution crystal structures of their complexes with acetylpolyamine amidohydrolase. Biochemistry 54, 4692−4703. (19) Christianson, D. W. (2005) Arginase: structure, mechanism, and physiological role in male and female sexual arousal. Acc. Chem. Res. 38, 191−201. (20) Lombardi, P. M., Cole, K. E., Dowling, D. P., and Christianson, D. W. (2011) Structure, mechanism, and inhibition of histone deacetylases and related metalloenzymes. Curr. Opin. Struct. Biol. 21, 735−743. (21) Porter, N. J., and Christianson, D. W. (2019) Structure, mechanism, and inhibition of the zinc-dependent histone deacetylases. Curr. Opin. Struct. Biol. 59, 9−18. (22) Ash, D. E. (2004) Structure and function of arginases. J. Nutr. 134, 2760S−2764S. (23) Gantt, S. L., Gattis, S. G., and Fierke, C. A. (2006) Catalytic activity and inhibition of human histone deacetylase 8 is dependent on the identity of the active site metal ion. Biochemistry 45, 6170− 6178. (24) Tabor, C. W., and Tabor, H. (1984) Polyamines. Annu. Rev. Biochem. 53, 749−790. (25) Thomas, T., and Thomas, T. J. (2001) Polyamines in cell growth and cell death: molecular mechanisms and therapeutic applications. Cell. Mol. Life Sci. 58, 244−258. (26) Bachmann, A. S., and Geerts, D. (2018) Polyamine synthesis as a target of MYC oncogenes. J. Biol. Chem. 293, 18757−18769. (27) Casero, R. A., Jr., Murray Stewart, T., and Pegg, A. E. (2018) Polyamine metabolism and cancer: treatments, challenges and opportunities. Nat. Rev. Cancer 18, 681−695.

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors thank the Northeastern Collaborative Access Team (NE-CAT) beamlines, which are funded by the National Institute of General Medical Sciences from the National Institutes of Health (P30 GM124165). The Pilatus 6M detector on beamline 24-ID-C is funded by a NIH-ORIP HEI grant (S10 RR029205). This research used resources of the Advanced Photon Source, a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under Contract DE-AC02-06CH11357. Additionally, the authors thank the Stanford Synchrotron Radiation Lightsource (SSRL), SLAC National Accelerator Laboratory, which is supported by the DOE Office of Science, Office of Basic Energy Sciences, under Contract DE-AC02-76SF00515. The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research and by the NIH, National Institute of General Medical Sciences (NIGMS) (including Grant P41GM103393). Finally, the authors thank the FMX beamline at the National Synchrotron Light Source II, a DOE Office of Science User Facility operated for the DOE Office of Science by Brookhaven National Laboratory under Contract DE-SC0012704. The Life Science Biomedical Technology Research resource is primarily supported by the National Institutes of Health, NIGMS, through a Biomedical Technology Research Resource P41 grant (P41GM111244), and by the DOE Office of Biological and Environmental Research (KP1605010).

■

REFERENCES

(1) Ouzounis, C. A., and Kyrpides, N. C. (1994) On the evolution of arginases and related enzymes. J. Mol. Evol. 39, 101−104. (2) Leipe, D. D., and Landsman, D. (1997) Histone deacetylases, acetoin utilization proteins and acetylpolyamine amidohydrolases are members of an ancient protein superfamily. Nucleic Acids Res. 25, 3693−3697. (3) Hai, Y., Shinsky, S. A., Porter, N. J., and Christianson, D. W. (2017) Histone deacetylase 10 structure and molecular function as a polyamine deacetylase. Nat. Commun. 8, 15368. (4) Kanyo, Z. F., Scolnick, L. R., Ash, D. E., and Christianson, D. W. (1996) Structure of a unique binuclear manganese cluster in arginase. Nature 383, 554−557. (5) Finnin, M. S., Donigian, J. R., Cohen, A., Richon, V. M., Rifkind, R. A., Marks, P. A., Breslow, R., and Pavletich, N. P. (1999) Structures of a histone deacetylase homologue bound to the TSA and SAHA inhibitors. Nature 401, 188−193. (6) Elkins, J. M., Clifton, I. J., Hernández, H., Doan, L. X., Robinson, C. V., Schofield, C. J., and Hewitson, K. S. (2002) Oligomeric structure of proclavaminic acid amidino hydrolase: evolution of a hydrolytic enzyme in clavulanic acid biosynthesis. Biochem. J. 366, 423−434. (7) Ahn, H. J., Kim, K. H., Lee, J., Ha, J.-Y., Lee, H. H., Kim, D., Yoon, H.-J., Kwon, A.-R., and Suh, S. W. (2004) Crystal structure of agmatinase reveals structural conservation and inhibition mechanism of the ureohydrolase superfamily. J. Biol. Chem. 279, 50505−50513. (8) Hai, Y., Dugery, R. J., Healy, D., and Christianson, D. W. (2013) Formiminoglutamase from Trypanosoma cruzi is an arginase-like manganese metalloenzyme. Biochemistry 52, 9294−9309. (9) Somoza, J. R., Skene, R. J., Katz, B. A., Mol, C., Ho, J. D., Jennings, A. J., Luong, C., Arvai, A., Buggy, J. J., Chi, E., Tang, J., Sang, B. C., Verner, E., Wynands, R., Leahy, E. M., Dougan, D. R., Snell, G., Navre, M., Knuth, M. W., Swanson, R. V., McRee, D. E., and Tari, L. K


Article

Biochemistry

for macromolecular crystallography. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 12−21. (48) Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) PROCHECK: A program to check the stereochemical quality of protein structures. J. Appl. Crystallogr. 26, 283−291. (49) Cox, J. D., Kim, N. N., Traish, A. M., and Christianson, D. W. (1999) Arginase-boronic acid complex highlights a physiological role in erectile function. Nat. Struct. Biol. 6, 1043−1047. (50) Ash, D. E., Cox, J. D., and Christianson, D. W. (1999) Arginase: a binuclear manganese metalloenzyme. In Manganese and Its Role in Biological Processes, Vol. 37 of Metal Ions in Biological Systems (Sigel, A., and Sigel, H., Eds.) pp 407−428, Marcel Dekker, New York. (51) Di Costanzo, L., Sabio, G., Mora, A., Rodriguez, P. C., Ochoa, A. C., Centeno, F., and Christianson, D. W. (2005) Crystal structure of human arginase I at 1.29-Å resolution and exploration of inhibition in the immune response. Proc. Natl. Acad. Sci. U. S. A. 102, 13058− 13063. (52) Hai, Y., Edwards, J. E., Van Zandt, M. C., Hoffmann, K. F., and Christianson, D. W. (2014) Crystal structure of Schistosoma mansoni arginase, a potential drug target for the treatment of schistosomiasis. Biochemistry 53, 4671−4684. (53) Ilies, M., Di Costanzo, L., Dowling, D. P., Thorn, K. J., and Christianson, D. W. (2011) Binding of α, α-disubstituted amino acids to arginase suggests new avenues for inhibitor design. J. Med. Chem. 54, 5432−5443. (54) Gelb, M. H., Svaren, J. P., and Abeles, R. H. (1985) Fluoro ketone inhibitors of hydrolytic enzymes. Biochemistry 24, 1813−1817. (55) Chakrabarti, P. (1989) Geometry of interaction of metal ions with sulfur-containing ligands in protein structures. Biochemistry 28, 6081−6085. (56) Arrar, M., Turnham, R., Pierce, L., de Oliveira, C. A. F., and McCammon, J. A. (2013) Structural insight into the separate roles of inositol tetraphosphate and deacetylase-activating domain in activation of histone deacetylase 3. Protein Sci. 22, 83−92. (57) Decroos, C., Christianson, N. H., Gullett, L. E., Bowman, C. M., Christianson, K. E., Deardorff, M. A., and Christianson, D. W. (2015) Biochemical and structural characterization of HDAC8 mutants associated with Cornelia de Lange Syndrome spectrum disorders. Biochemistry 54, 6501−6513. (58) Porter, N. J., Christianson, N. H., Decroos, C., and Christianson, D. W. (2016) Structural and functional influence of the glycine-rich loop G302GGGY on the catalytic tyrosine of histone deacetylase 8. Biochemistry 55, 6718−6729. (59) Lanyi, J. K. (1974) Salt-dependent properties of proteins from extremely halophilic bacteria. Bacteriol. Rev. 38, 272−290. (60) Dym, A., Mevaerech, M., and Sussman, J. L. (1995) Structural features that stabilize halophilic malate dehydrogenase from an archaebacterium. Science 267, 1344−1346. (61) Graziano, G., and Merlino, A. (2014) Molecular basis of protein halotolerance. Biochim. Biophys. Acta, Proteins Proteomics 1844, 850−858. (62) DasSarma, S., and DasSarma, P. (2015) Halophiles and their enzymes: negativity put to good use. Curr. Opin. Microbiol. 25, 120− 126. (63) Fujishiro, K., Ando, M., and Uwajima, T. (1988) Crystallization and some properties of acetylpolyamine amidohydrolase from Mycoplana bullata. Biochem. Biophys. Res. Commun. 157, 1169−1174. (64) Sakurada, K., Ohta, T., Fujishiro, K., Hasegawa, M., and Aisaka, K. (1996) Acetylpolyamine amidohydrolase from Mycoplana ramosa: gene cloning and characterization of the metal-substituted enzyme. J. Bacteriol. 178, 5781−5786.

(28) Bloomfield, V. A. (1997) DNA condensation by multivalent cations. Biopolymers 44, 269−282. (29) Thomas, T. J., and Thomas, T. (2018) Collapse of DNA in packaging and cellular transport. Int. J. Biol. Macromol. 109, 36−48. (30) Trachman, R. J., III, and Draper, D. E. (2017) Divalent ion competition reveals reorganization of an RNA ion atmosphere upon folding. Nucleic Acids Res. 45, 4733−4742. (31) Childs, A. C., Mehta, D. J., and Gerner, E. W. (2003) Polyamine-dependent gene expression. Cell. Mol. Life Sci. 60, 1394− 1406. (32) Ivanov, I. P., Atkins, J. F., and Michael, A. J. (2010) A profusion of upstream open reading frame mechanisms in polyamine-responsive translational regulation. Nucleic Acids Res. 38, 353−359. (33) Dever, T. E., and Ivanov, I. P. (2018) Roles of polyamines in translation. J. Biol. Chem. 293, 18719−18729. (34) Shinsky, S. A., and Christianson, D. W. (2018) Polyamine deacetylase structure and catalysis: prokaryotic acetylpolyamine amidohydrolase and eukaryotic HDAC10. Biochemistry 57, 3105− 3114. (35) Blankenship, J. (1978) Deacetylation of N8-acetylspermidine by subcellular fractions of rat tissue. Arch. Biochem. Biophys. 189, 20−27. (36) Libby, P. R. (1978) Properties of an acetylspermidine deacetylase from rat liver. Arch. Biochem. Biophys. 188, 360−363. (37) Handley, K. M., and Lloyd, J. R. (2013) Biogeochemical implications of the ubiquitous colonization of marine habitats and redox gradients by Marinobacter species. Front. Microbiol. 4, 136. (38) Fathepure, B. Z. (2014) Recent studies in microbial degradation of petroleum hydrocarbons in hypersaline environments. Front. Microbiol. 5, 173. (39) Cluff, M. A., Hartsock, A., MacRae, J. D., Carter, K., and Mouser, P. J. (2014) Temporal changes in microbial ecology and geochemistry in produced water from hydraulically fractured Marcellus shale gas wells. Environ. Sci. Technol. 48, 6508−6517. (40) Bonis, B. M., and Gralnick, J. A. (2015) Marinobacter subterrani, a genetically tractable neutrophilic Fe(II)-oxidizing strain isolated from the Soudan Iron Mine. Front. Microbiol. 6, 719. (41) Decroos, C., Bowman, C. M., and Christianson, D. W. (2013) Synthesis and evaluation of N8-acetylspermidine analogues as inhibitors of bacterial acetylpolyamine amidohydrolase. Bioorg. Med. Chem. 21, 4530−4540. (42) Battye, T. G. G., Kontogiannis, L., Johnson, O., Powell, H. R., and Leslie, A. G. W. (2011) iMosflm: a new graphical interface for diffraction-image processing with Mosflm. Acta Crystallogr., Sect. D: Biol. Crystallogr. 67, 271−281. (43) Winn, M. D., Ballard, C. C., Cowtan, K. D., Dodson, E. J., Emsley, P., Evans, P. R., Keegan, R. M., Krissinel, E. B., Leslie, A. G. W., McCoy, A., McNicholas, S. J., Murshudov, G. N., Pannu, N. S., Potterton, E. A., Powell, H. R., Read, R. J., Vagin, A., and Wilson, K. S. (2011) Overview of the CCP4 suite and current developments. Acta Crystallogr., Sect. D: Biol. Crystallogr. 67, 235−242. (44) Terwilliger, T. C., Grosse-Kunstleve, R. W., Afonine, P. V., Moriarty, N. W., Zwart, P. H., Hung, L.-W., Read, R. J., and Adams, P. D. (2008) Iterative model building, structure refinement and density modification with the PHENIX AutoBuild wizard. Acta Crystallogr., Sect. D: Biol. Crystallogr. 64, 61−69. (45) Emsley, P., Lohkamp, B., Scott, W. G., and Cowtan, K. (2010) Features and development of Coot. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 486−501. (46) Adams, P. D., Afonine, P. V., Bunkóczi, G., Chen, V. B., Davis, I. W., Echols, N., Headd, J. J., Hung, L., Kapral, G. J., GrosseKunstleve, R. W., McCoy, A. J., Moriarty, N. W., Oeffner, R., Read, R. J., Richardson, D. C., Richardson, J. S., Terwilliger, T. C., and Zwart, P. H. (2010) PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr., Sect. D: Biol. Crystallogr. 66, 213−221. (47) Chen, V. B., Arendall, W. B., III, Headd, J. J., Keedy, D. A., Immormino, R. M., Kapral, G. J., Murray, L. W., Richardson, J. S., and Richardson, D. C. (2010) MolProbity: all-atom structure validation L


Structure and Function of the Acetylpolyamine Amidohydrolase from

Recommend Documents