Insights into the mechanism of paraoxonase-1: Comparing the

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Insights into the mechanism of paraoxonase-1: Comparing the reactivity of the six-bladed #-propeller hydrolases Timothy John Grunkemeyer, David Garcia Mata, Kiran Doddapaneni, Srividya Murali, and Thomas J. Magliery Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.8b01115 • Publication Date (Web): 14 Dec 2018 Downloaded from http://pubs.acs.org on December 16, 2018

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Biochemistry

Insights into the mechanism of paraoxonase-1: Comparing the reactivity of the six-bladed β-propeller hydrolases Timothy J. Grunkemeyer1,2, David G. Mata1, Kiran Doddapaneni1, Srividya Murali1, Thomas J. Magliery1,* 1

Department of Chemistry & Biochemistry, The Ohio State University, Columbus, Ohio, USA

2

Ohio State Biochemistry Program

*

To whom correspondence should be addressed: Department of Chemistry & Biochemistry, 100 W. 18th Ave., Columbus, OH 43210; email [email protected]; phone +1 (614) 247-8425; fax +1 (614) 292-1685

ACS Paragon Plus Environment

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Abstract The mammalian protein paraoxonase-1 (PON1) has been explored as a promising bioscavenger treatment for organophosphorus (OP) agent poisoning, but it is not active enough to protect against many agents. Engineering is limited because PON1’s catalytic mechanism is poorly understood; moreover, its native activity and substrate are unknown. PON1 is a calcium-bound six-bladed βpropeller hydrolase that shares high structural homology, a conserved metal-coordinating active site, and substrate specificity overlap with other members of a superfamily that includes squid diisopropylfluorophosphatase (DFPase), bacterial drug responsive protein 35 (Drp35), and mammalian senescence marker protein 30 (SMP30). We hypothesized that, by examining the reactivity of all four hydrolases using a common set of conservative mutations, we could gain further insight into the catalytic mechanism of PON1. We chose a set of mutations to examine conserved Asp and Glu residues in the hydrolase active sites, as well as the ligation sphere around the catalytic calcium and a His-His dyad seen in PON1. The wild-type (WT) and mutant hydrolases were assayed against a set of lactones, aryl esters, and OPs that PON1 is known to hydrolyze. Surprisingly, some mutations of Ca2+ coordinating residues, previously thought to be essential for turnover, resulted in significant activity toward all substrate classes examined. Additionally, merely maintaining WT-like charge in the active site of PON1 was insufficient for high activity. Finally, the H115-H134 dyad does not appear to be essential for catalysis against any substrate. Therefore, previously proposed mechanisms must be re-evaluated.


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Biochemistry

Introduction Three million people are exposed to high levels of neurotoxic organophosphorus (OP) compounds each year, approximately 10% of whom die from the exposure.1 The mechanism of toxicity for OPs is chemical modification of the active site serine of acetylcholinesterase (AChE). Over time, this modification becomes irreversible through a process called “aging,” preventing any future hydrolysis of acetylcholine, a critical signaling compound. This results in an overstimulation of the nicotinic acetylcholine receptors in the central nervous system, after which symptoms including muscle weakness, hypoxia, paralysis, and eventually coma and death occur.2 The standard of care for OP poisoning is a competitive antagonist (e.g., atropine) combined with an AChE re-activator (e.g., an oxime such as 2-PAM).3,4 Furthermore, the stoichiometric bioscavenger butyrylcholinesterase5 (BuChE) is in advanced development as well. However, the atropine/oxime combination is sedating and does not prevent seizures from reoccurring,6 and the BuChE treatment requires a large (stoichiometric) amount of enzyme to be administered.7 With the use of OP compounds as pesticides in developing countries1 as well as the recent re-emergence of OP chemical weapon use, there is a great need for improved interventions. A number of enzymes have been proposed as catalytic bioscavengers, to combine the effectiveness of BuChE with the smaller doses afforded by multiple turnover, including the bacterial protein organophosphatase hydrolase (OPH) and mammalian protein paraoxonase-1 (PON1, GenBank AAR95986). Because PON1 is a long-lived serum enzyme, it has many potential advantages in terms of safety and pharmacokinetics, but its inherent OP hydrolase activity is too low to be protective for most threat compounds. Consequently, we and others have used rational and random mutagenesis to improve the activity of PON1 as a potential therapeutic8– 15

, but this has been hampered by a poor understanding of PON1’s mechanism.


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PON1 is a calcium-bound mammalian enzyme with a six-bladed β-propeller fold that exhibits promiscuous hydrolytic ability against lactones, aryl esters, and OPs.16 While its physiological function is not fully understood, it has been shown to be secreted from the liver and is bound to HDL in mammalian serum.17,18 It is hypothesized that, while bound to HDL, PON1 plays a role in helping neutralize toxic components of plasma, specifically oxidized lipids.19 It also may mediate cholesterol efflux from macrophages, and along with its role in the hydrolysis of oxidized lipids, it may play a role in the prevention of atherosclerosis.20,21 It has been shown to hydrolyze lactones better than any other substrate, and therefore it is believed that its native activity is as a lactonase, although the native substrate is unclear.16 PON1 is also able to hydrolyze aryl esters and OPs.12,15,16,22,23 For example, PON1 exhibits some activity toward the pesticides paraoxon, chlorpyrifos oxon, diazinon oxon, certain G-agents, and to a much lesser extent Vagents.9,10,12,22–24 After extensive mutagenesis and screening campaigns, highly active pan-G agents variants were isolated, but only weak activity against V agents was elicited. Part of the problem is that the mechanism by which PON1 acts as a hydrolase remains elusive. It is even unclear whether PON1 hydrolyzes all substrates using one overarching mechanism. A great deal of work has targeted critical residues in the active site of PON1 from which three essential mechanistic hypotheses have emerged.19,21–26 These residues include a “histidine dyad,” H115 and H134, along the wall of the active site, as well as a group of residues in the base of the active site (E53, N168, N224, D269) that coordinate what is referred to as the “catalytic calcium,” which is believed to be critical for activity.24,27-27 It has been proposed that the histidine dyad mediates lactonase/esterase activity of PON1 by acting as a water-activating proton shuttle (Figure 1A).25,30 While N168 and N224 are believed to participate merely in calcium binding, it has also been suggested that E53 and D269 are critical for phosphotriesterase


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Biochemistry

activity.23,24,28,31 D269 has been proposed to either participate directly in the mechanism as a nucleophile, or indirectly as a general base, where it would activate water (Figure 1B,C).31 Mutagenic and simulation data support a key role for D269,23,24,26,31 but there is evidence for all three mechanisms.16,23,25–27,30–32 Although it fundamentally seems unlikely that two different hydrolytic mechanisms are at work in the same active site, mutations to H115 that ablate lactonase activity but enhance OPase activity raise questions.

Figure 1. Proposed mechanisms for the six-bladed β-propeller hydrolases. (A) PON1 proton shuttling mechanism implicating the H115-H134 dyad in abstraction of a proton from water leading to attack on the bound lactone.21 (B) PON1 water activation mechanism proposed involving D269. D269 abstracts a proton from a nearby water, leading to attack of the substrate by the generated OH-.28 (C) PON1 mechanism implicating D269 in direct attack of substrate. A newly generated OH- is free to attack the acyl-enzyme intermediate which has been formed, to regenerate the enzyme and release the product.28 (D) DFPase D229 direct attack mechanism proposed to follow the same reaction pathway as the PON1 D269 direct attack mechanism shown in (C).40 (E) The mechanism for DHC hydrolysis by Drp35 implicates two Asp residues, D236 (homologous to D269 and D229 in PON1 and DFPase) and D138 (homologous to N168 and N120 in PON1 and DFPase). D138 initiates the reaction by abstracting a proton from a nearby water, leading to attack on the carboxyl center by the generated OH-.31

We decided to approach this problem from a different perspective, by comparing the effects of mutations in PON1 to similar mutations in three other six-bladed β-propeller hydrolases:


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diisopropylfluorophosphatase (DFPase, UniProtKB Q7SIG4), drug responsive protein-35 (Drp35, UniProtKB Q7A338), and senescence marker protein 30 (SMP30, UniProtKB Q15493). Despite minimal sequence homology and identity, all four enzymes share a highly conserved fold (Figure 2A) as well as a nearly identical active site.33–36 They all bind calcium in the base of the putative active site using four conserved amino acids (with one exception: Drp35 has an Asp in the position of Asn168 in PON1) (Figure 2B,C).34,37,38 Despite originating from bacteria, squid and humans (Staphylococcus aureus, Luligo vulgaris, and Homo sapiens for Drp35, DFPase and PON1/SMP30, respectively), they all have been shown to hydrolyze some esters, OPs, or both. This has been attributed to the similarities in their binding pockets. But the differences in their binding pockets have also been shown to dictate their substrate specificities, accounting for the differences in activity profiles between the enzymes.39 DFPase and SMP30 exhibit phosphotriesterase activity, while Drp35 and SMP30 exhibit lactonase activity.26,34,38–44 Finally, a variety of mechanisms have been proposed for PON1, DFPase, and Drp35 (no mechanisms have been proposed for SMP30 to our knowledge) as shown in Figure 1B,C,D,E,25,31,34,43 but in principle all of these proposed mechanisms could operate in each enzyme. It has been suggested that PON1 and DFPase share the same mechanism,26 and Drp35 has been proposed to be the “bacterial counterpart of eukaryotic PON1.”34,44 We therefore hypothesized that the same mechanism may be at work in all four enzymes.


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Biochemistry

Figure 2. Alignment of proteins and active sites. Global backbone (A) and active site (B,C) alignment of DFPase (PDB ID: 3O4P, blue), Drp35 (PDB ID: 2DG1, green), G3C9 PON1 (PDB ID: 1V04, red), and SMP30 (PDB ID: 3G4E, purple). Residues of interest are shown in sticks and Ca2+ are shown in spheres. Conserved residues are labeled in black, non-conserved residues are labeled in the respective protein’s colors. Rendered with PyMOL.

To test this, residues corresponding to E53, D269, and H115 in PON1 were identified in DFPase, Drp35, and SMP30 via an active site alignment model from which conservative mutations were made. The WT and mutant enzymes were screened against a panel of substrates including an aryl ester, lactones, and OPs, all of which PON1 is known to hydrolyze. Furthermore, it has been shown that charge balance of the calcium binding residues in DFPase is critical for its phosphotriesterase activity.37,45 We also created a similar set of mutations in PON1 to see if a similar effect on activity is observed. Together, these experiments identified common critical residues in and around the hydrolase active sites and also rule out some commonly-accepted beliefs about the PON1 mechanism.

MATERIALS AND METHODS Materials SMP30 cDNA was purchased from Origene (Rockville, MD). The genes for DFPase and Drp35 were synthesized by Genewiz (South Plainfield, NJ). The PON1 template DNA (G3C9) for


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rational variant clonings was obtained from the Tawfik group at the Weizmann Institute of Science (Rehovot, Israel). Oligonucleotide primers were designed and ordered from Sigma Aldrich (St. Louis, MO). Herculase II Fusion DNA Polymerase (600675) was purchased from Agilent Technologies (Santa Clara, CA). T4 DNA Ligase (M0202T), dNTPs, and all restriction enzymes were purchased from New England Biolabs (Ipswich, MA). Ampicillin (A-301-10), IPTG (I2481C50), and phenylmethylsulfonyl fluoride (PMSF) (P-470-10) were purchased from Gold Biotechnology Inc. (St. Louis, MO). HisPur Ni-NTA resin (88222) was purchased from Thermo Scientific. Protein molecular weight marker came from USB and Bradford reagent was purchased from Bio-Rad Laboratories (Hercules, CA). Paraoxon-Ethyl (D9286), Tergitol NP-10 Nonionic, D-(+)-gluconic acid δ-lactone (G4750), diisopropylfluorophosphate (D0879), dihydrocoumarin (D104809), phenyl acetate (108723) were obtained from Sigma Aldrich (St. Louis, MO). Chlorpyrifos oxon (MET-674B) and the diazinon-O-analogue (MET-11621A) were obtained from Chem Service Inc. (West Chester, PA). O-cyclohexylmethylphosphonyl coumarin and thiobutyl butyrolactone were kindly supplied by Prof. Christopher Hadad (Department of Chemistry & Biochemistry, The Ohio State University). Standard 96-well microplates and UV transparent microplates were purchased from USA Scientific (Ocala, FL). All other reagents used were of analytical grade purchased from Fisher Scientific. Buffers used were prepared in double distilled water and filtered using various filtration units.

Molecular Cloning and Mutagenesis DFPase and Drp35 genes were cloned into the pHLIC expression vector using the NcoI and BamHI restriction sites,46 directly downstream of a Tobacco Etch Virus (TEV)-protease cleavable Nterminal 6×His tag. Constructs were sequence-confirmed at Genewiz. SMP30 was cloned into the


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Biochemistry

pHMT vector (a gift of Prof. Mark Foster, Department of Chemistry & Biochemistry, The Ohio State University) using the EcoRI and XhoI restriction sites. This added a TEV-protease cleavable N-terminal 6×His-MBP fusion tag to SMP30. This scheme was used for all subsequent SMP30 mutants. The PON1 G3C9 variant is cloned into pET32b and has a C-terminal 6×His tag. All single mutants were cloned with the above noted WT constructs as templates by overlap PCR using complementary mutagenic primers (the SMP30 E18Q mutation was close enough to the C-terminus of the protein that the mutation was simply added into the forward amplification primer). All multi-mutants were made sequentially by adding single mutations to previously cloned single, double, or triple mutants. For instance, E53Q was cloned first after which the E53Q/D269N mutant was made by adding the D269N mutation to the E53Q template. Briefly, 5’ and 3’ amplicons were made by PCR from the 5’ cloning site to the site of mutation, and from the site of mutation to the 3’ cloning site. These were digested with DpnI, mixed, and amplified with the primers at the cloning sites after only a few cycles of primerless amplification to generate the full-length mutated fragment.

Protein Expression The expressions of both DFPase and Drp35 (and their mutants) were carried out in the same manner as previously described from BL21(DE3) E. coli.34,47 PON1 and SMP3048 were expressed in the same manner as DFPase and Drp35, with two differences. The cultures were supplemented with 1 mM CaCl2, and the SMP30 expression media was also supplemented with 1% glycerol. At induction, the growth temperature for PON1 was decreased to 30 °C rather than 16 °C, after which the culture was only grown for four hours instead of overnight. The cells were centrifuged and the pellet was harvested and stored at -80 °C the same day. For SMP30, after the OD600 had reached


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0.7, the culture was cold shocked at 4 °C for 4 h. The remaining expression of SMP30 was done in the same manner as DFPase and Drp35.

Purification DFPase and Drp35 were purified and TEV cleaved using the same protocols.34,47 All steps were carried out on ice unless otherwise noted. The pellets were re-suspended in the appropriate lysis buffers (10 mM Tris pH 7.4, 500 mM NaCl, 2 mM CaCl2, 2 mM β-mercaptoethanol (BME) and 50 mM Tris pH 7.4, 300 mM NaCl, 1 mM CaCl2, 2 mM BME for DFPase and Drp35, respectively) at a ratio of 20 mL/L culture. Once re-suspended, the culture was supplemented with 1 mM PMSF and lysed using an Avestin Emulsiflex-C3 homogenizer. The lysate was centrifuged for one hour at 15,000 RPM. One mL of Ni-NTA resin/L culture was added to the supernatant and incubated for 2 h at 4 °C. This suspension was added to a fritted column, and the flow through was collected and set aside. The column was washed with 50 column volumes (CV) of lysis buffer supplemented with 10 mM imidazole. The protein was then eluted off the column by washing the resin twice with 10 CV of lysis buffer supplemented with 200 mM imidazole. The flow through, wash, and elutions were analyzed by reducing SDS-PAGE gel to ensure the presence and purity of protein. The fractions containing protein were dialyzed against 4 L of TEV cleavage buffer (50 mM Tris pH 8.0, 50 mM NaCl, 1 mM CaCl2) for 4 h at 4 °C. The TEV cleavage reaction contained 10 mg of protein, 5 mM DTT, and 1 mg of TEV protease. This reaction was carried out overnight at 4 °C. The reaction was then dialyzed against 4 L of the desired assay buffer (10 mM Tris pH 7.0, 100 mM NaCl, 2 mM CaCl2 and 50 mM Tris pH 7.0, 300 mM NaCl, 1 mM CaCl2 for DFPase and Drp35, respectively) for 4 h at 4 °C in order to remove the DTT. The liberated His tag was rebound to 1 mL of Ni-NTA resin by nutating the resin with the reaction for 2 hours at 4 °C. The


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Biochemistry

flow through from this column contained the protein of interest and was subsequently dialyzed into the appropriate assay buffer. PON1 and SMP3048 were purified in a similar manner with minor differences. The purification buffers used for PON1 and SMP30 were 50 mM Tris pH 8.0, 50 mM NaCl, 1 mM CaCl2, 2 mM BME and 50 mM Tris pH 7.4, 200 mM NaCl, 1 mM CaCl2, 2 mM BME, respectively. SMP30 required the addition of a cOmplete protease inhibitor tablet (Roche) to the lysis buffer before pellet re-suspension. Additionally, since PON1 and SMP30 are hypothesized to be membrane-bound, they required detergent extraction. 0.1% Tergitol was added to the lysate after cell lysis, with stirring/nutating for 2.5 h for PON1 and 4 h for SMP30 at 4 °C. Additionally, since the 6x-His tag on PON1 is not TEV cleavable and since the SMP30-MBP construct was observed to have the same activity as SMP30 by itself, TEV cleavages were not carried out for either protein or their mutants. SMP30 required additional purification steps to achieve homogeneity. After elution from the Ni-NTA column, the solution was added to 2 mL of Amylose resin/L of culture and nutated for 2 h at 4 °C. The flow through was collected, and the resin was washed with 50 CV of lysis buffer, and eluted 3 times with 5 CV of 10 mM maltose. After all purification steps, both enzymes were dialyzed into their respective assay buffers (50 mM Tris pH 7.0, 50 mM NaCl, 1 mM CaCl2 and 25 mM Tris pH 7.0, 100 mM NaCl, 1 mM CaCl2 for PON1 and SMP30, respectively). Protein concentrations were determined by Bradford assay and confirmed by SDSPAGE with Coomassie Brilliant Blue staining and comparison to BSA.

Enzyme Kinetics All kinetic assays were carried out at room temperature (25 ºC). To screen each enzyme, a panel of aryl esters, lactones, and organophosphates with chromogenic leaving groups was assembled


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(Figure 3). The concentrations of substrates were confirmed by following their hydrolysis using NaOH or the enzyme 3B3 for CMP, which hydrolyzes only one enantiomer.12 Each enzyme was then tested independently against each substrate. The highest concentrations of substrates allowed by solubility limits were used to achieve maximum signal intensity and avoid possible detection limitations. All substrates were dissolved in methanol except TBBL (acetonitrile) and DFP (2propanol). The alcohol concentrations in the assays were kept constant (4 μL or 2% of the total reaction volume) by making a substrate dilution series in methanol (or acetonitrile and 2-propanol for TBBL and DFP respectively). Enzymes were concentrated to 1 mg/mL per protein, then diluted 1:10, 1:100, and 1:1000. The concentrated protein and dilutions were then screened against the substrates at the appropriate wavelengths (Figure 3) in a Molecular Devices M5 Spectramax plate reader. To do this, 10 μL of enzyme was added to 186 μL buffer and 4 μL of substrate and product formation was followed for 10 minutes at the desired wavelength. Separate reactions with enzyme and buffer only, and substrate and buffer only, were run as noted above for enzyme-only and substrate-only negative controls. Active enzymes exhibited a slope greater than the sum of the slopes of the two blanks.


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Biochemistry

Figure 3. Substrates used for activity assays. The method of observation of turnover is noted (wavelength refers to where we are observing the formation of the hydrolysis product). Abbreviations and method of reaction tracking are noted.

The substrates diisopropylfluorophosphate (DFP) and d-gluconic acid δ-lactone (GL) required use of a pH STAT titrator to measure kinetics due to their lack of a chromogenic leaving group, and protocols were designed as previously described.49 The same screening protocol was used as described above. Maximum substrate concentrations were tested against concentrated, 1:10, 1:100, and 1:1000 dilutions of enzyme. For all reactions, a reaction volume of 5 mL (250 μL of enzyme was mixed with 4,650 μL assay buffer—10 mM Tris pH 7.0, 50 mM NaCl, 1 mM CaCl2—with 100 μL substrate) was used and the reactions were carried out at room temperature under nitrogen atmosphere for 5 min. The instrument was programmed to perform a “pre-STAT” measurement where the assay buffer and substrate mixture were adjusted to pH 7.000 with NaOH. After enzyme addition, the titrant (2 mM NaOH) was added into the reaction at 50 μL/min in order to keep the pH constant at 7.000 for 5 min. From this, volume of titrant was plotted against time and used to calculate product formation based on the ratio of hydrogen ions produced per substrate


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molecule hydrolyzed (2:1 in the case of DFP and GL). To calculate corrected kinetic parameters, non-enzymatic hydrolysis rates were taken for both DFP and GL as described above, but without the addition of enzyme. This rate was subtracted from the enzymatic reactions to obtain corrected enzymatic rates. Once active variants had been identified, they were subjected to a full battery of kinetics.50 The substrate concentration ranges were used in activity assays for dihydrocoumarin (DHC), TBBL,

GL,

phenyl

acetate

(PA),

O-cyclohexyl-O-(3-cyano-4-methyl-7-coumarinyl)-

methylphosphonate (CMP), paraoxon (PX), chlorpyrifos oxon (CPO), diazinon oxon (DZO), and DFP were 0.016-1, 0.05-0.5, 0.063-1, 0.031-2, 0.007-0.052, 0.016-1, 0.016-0.5, 0.016-1, and 0.063-1 mM, respectively. Enzyme concentration against each substrate was optimized to ensure that the assumptions of the Michaelis-Menten kinetic model were satisfied (enzyme concentrations used are noted in Table S6). The assays to determine kcat/Km for the hydrolysis of PA, DHC, TBBL, CMP, PX, CPO, and DZO were performed using an M5 Spectramax plate reader. A minimum of four substrate concentrations per substrate was required for accurate fitting to the Michaelis-Menten kinetic model and subsequent calculations. Reactions were recorded in triplicate. Each reaction contained 186 μL assay buffer, 4 μL substrate, and 10 μL enzyme, added in that order. Product formation was then followed for 5 min at the desired wavelength. To measure TBBL kinetics, DTNB was mixed with the assay buffer in a 1:20 volumetric ratio and the assay was conducted while detecting product formation at the wavelength of formation of DTN- (405 nm). Kinetics for DFP and GL were carried out in duplicate using the pH STAT titrator as described above. Values were replicated with at least one independent enzyme preparation, and replicate values were within 2-fold. The greatest source of replicate error is likely estimation of enzyme concentration.


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Microsoft Excel and Kaleidagraph were used to calculate kinetic parameters. The slopes (absorbance vs. time) of each trial and blank within a substrate concentration were calculated and averaged. The background signal given from the enzyme and substrate blanks were subtracted from this value. This value was then converted to the rate of product formation using a path length of 1.76 cm and the appropriate leaving group extinction coefficients. Since most of the substrates assayed here are sparingly soluble in water, they had to be in very low concentrations to avoid precipitation. Therefore, for most variants, only kcat/Km could be obtained from the initial linear portion of the concentration versus rate plot. Non-chromogenic kinetic measurements taken via pH STAT titration were compiled into Excel from which kinetic parameters were calculated. Rate of base (2 mM NaOH) titration was calculated for both enzymatic and non-enzymatic hydrolysis by taking the slope of time vs. volume of base added. The molar concentrations of substrate hydrolyzed were then extrapolated for both types of reaction (taking into account that for every 2 mol base added, 1 mol of substrate was hydrolyzed). The non-enzymatic rate of product formation was subtracted from that of the enzymatic rate to obtain corrected rates.

RESULTS Cloning, Expression, and Purification of Hydrolases We expressed DFPase and Drp35 from our T7 pHLIC expression vector, removing the N-terminal 6×His tag with TEV protease.46 SMP30 was cloned into pHMT, creating a MBP fusion that greatly enhanced its solubility and allowed for successful purification. The mammalian hybrid PON1 (G3C9) was expressed from pET32b with a C-terminal 6×His tag that was not removed. DFPase, Drp35, and PON1 were expressed in BL21(DE3) E. coli cells, while SMP30 was expressed in C43(DE3) Walker series E. coli cells.51 All variants were purified by NiNTA chromatography,


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with an additional amylose affinity step for the MBP-SMP30 fusion. This yielded samples with >95% purity as opposed to the ~70% purity off the Ni-NTA column. All proteins were ultimately purified to >95% purity.

Kinetic Screening and Characterization All WT enzymes were screened against the set of lactones, aryl esters, and organophosphates in Figure 3. It has been previously shown that PON1 will readily hydrolyze all of them with reasonable activity.9,10,12,15,16,22,23,50 Active enzyme-substrate pairs were identified, and were further characterized with a series of substrate concentrations to determine kinetic constants. PON1 showed high catalytic efficiency against the lactones DHC (4.7 × 106 M-1 s-1) and TBBL (1.0 × 105 M-1 s-1), and the aryl ester PA (1.8 × 105 M-1 s-1). More modest activity was observed against the OP pesticide oxons PX (5.6 × 103 M-1 s-1), chlorpyrifos oxon (CPO) (7.0 × 105 M-1 s-1), and diazinon oxon (DZO) (2.1 × 104 M-1 s-1). Additionally, PON1 was shown to hydrolyze the OP nerve agent simulants CMP (4.4 × 104 M-1 s-1) and DFP (1.3 × 103 M-1 s-1) (Figure 4, Table S1). kcat and Km could only be determined independently for WT PON1 (G3C9) against DHC, CMP, PX, and CPO (Table S4); the enzyme could not be saturated by TBBL, PA, DZO or DFP at accessible concentrations. The Km values for the saturating substrates were overall fairly high and in a fairly narrow range from 6 μM to 210 μM, while the kcat values varied over a 16-fold larger range from 0.3 s-1 to 172 s-1. In general, it is difficult to saturate PON1 with its poorly water soluble substrates, and differences in kcat probably matter more in determining catalytic efficiency.


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Biochemistry

Figure 4. Six-bladed β-propeller hydrolase wild type, active site Asp, and Glu variant activities. A. Lactonase and aryl esterase catalytic efficiencies. B. Phosphotriesterase catalytic efficiencies. All values presented in M-1 s-1 were obtained using a linear fit to the rate versus initial substrate data. Error bars indicate standard error to a linear fit (except variants listed in Table S4, for which kinetic parameters and errors were obtained by fitting data to the Michaelis Menten equation using the graphing software Kaleidagraph). *Indicates weak activity was observed, but a value was not calculated as it was near the detection limit of the experiment.

DFPase exhibited strong activity against DFP (6.8 × 105 M-1 s-1), agreeing well with previous values shown in literature. It also exhibited moderate catalytic efficiency against DHC (1.0 × 103 M-1 s-1) and low activity against TBBL (10 M-1 s-1). We report for the first time its ability


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to hydrolyze CPO (2.9 × 102 M-1 s-1) and GL (3.3 × 103 M-1 s-1). Drp35 also showed moderate to low activity against the lactones DHC (2.7 × 104 M-1 s-1) and TBBL (20 M-1 s-1), and a minimal ability to hydrolyze DFP. (‘Minimal’ refers to activity that can be seen above background, but it is too low to be accurately quantified.) SMP30 was only able to hydrolyze DHC (3.0 × 102 M-1 s1

) and GL (2.4 × 104 M-1 s-1) as expected, indicating that it is only a lactonase (Figure 4, Table S1).

For these enzymes and substrates, only kcat/Km values were obtained as saturation was never achieved. Interestingly, none of the enzymes except PON1 could hydrolyze the aryl ester PA. Once the WT enzymes had been characterized, mutations were made to what have been hypothesized to be critical residues in the active site. Specifically, the Asp and Glu in the base of the active site were targeted, as they have been hypothesized to participate either directly or indirectly in catalysis and Ca2+ binding (Figure 2B,C)23,25. In PON1, D269 was mutated to both Asn and Glu. The D269N mutation resulted in a three order of magnitude activity decrease against DHC, four order loss against TBBL, and near complete loss in observable activity for PA, CMP, PX, CPO, and DFP. The D269E mutation exhibited a more moderate effect on activity. Only one order of magnitude of activity was lost against DHC, along with two orders against TBBL and CMP. Yet, nearly all activity was still lost against PA and PX (CPO, DZO, DFP, GL were not tested). E53Q was found to have a large effect on activity as well, decreasing DHC hydrolysis by three orders of magnitude and TBBL and PA activity by two orders of magnitude. Little to no phosphotriesterase activity was observed, as the only activity maintained was against CPO (7.8 × 102 M-1 s-1), a three order of magnitude decrease. The corresponding mutations in DFPase (D229N and E21Q), Drp35 (D236N and E48Q), and SMP30 (D204N and E18Q) were also made. Nearly all activity was lost for all six of these for all substrates tested (Figure 4, Table S1). But it is worth noting that the WT activities were also lower for all of these substrates, and two-to-four order of


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Biochemistry

magnitude losses as seen with PON1 would be beyond the detection threshold. Mutation of these two key residues results in a profound loss in activity for all substrates, but the residual activities for the best substrates are larger than expected for mutation of a residue that directly participates in catalysis. We next turned our attention to the importance of charge balance in the active site of PON1. In 2010, Blum and coworkers characterized the importance of charge balance in DFPase.37 The mutations altered the number of charged and neutral residues and moved the positions of charged residues to explore changes in net charge and also the pattern of charge around the catalytic cation. The present experiment was designed in the same manner, creating charge balance alterations among the five residues closest to Ca2+ in the base of the active site: E53, N168, D183, N224, and D269 (Figure S3). The canonical total charge of these five residues and Ca2+ is taken to be -1. These variants were tested against DHC, TBBL, PA, CMP, and PX (Figure 5, Table S2). The E53Q/D269N (+1 charge, removed 2 negative charges) mutation, which neutralizes both residues hypothesized to participate mechanistically, resulted in a two order of magnitude loss of activity against all five substrates in question—large, but not as large as the single mutants, above. The E53Q/D269E (0 charge, removed 1 negative charge) mutation exhibited nearly the exact same effect on the enzyme. When a single negative charge was moved from E53 to N168 by making the E53Q/N168D (-1 charge, WT-like) mutation, it exhibited a nearly identical effect to the first two mutations tested, except for an additional ten-fold loss of activity against TBBL (Figure 5). These first three mutations all retained the ability to be saturated by DHC, TBBL, and CMP (except E53Q/N168D against TBBL). These mutations all have roughly the same effect on kcat and Km for each substrate (Table S4).


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Figure 5. G3C9 PON1 wild type and charge balance variant activities. See notes in Fig. 4.

Additionally, a negative charge was moved from D269 to N168 with the N168D/D269N mutation, from D269 to N224 with the N224D/D269N mutation, and from D183 to N168 with the N168D/D183N mutation. These all maintain the overall -1 charge of G3C9, while moving the charge around the active site. All of these mutations have a very significant effect on catalysis of DHC and PX, reducing them to minimal or undetectable levels. A significant effect was also observed on TBBL, with only N168D/D269N hydrolyzing substrate in a detectable range (80 ± 5 M-1 s-1). A similar effect was observed on the hydrolysis of PA and CMP, with only N224D/D269N hydrolyzing them in the detectable range, 1.0 × 102 M-1 s-1 and 2.0 × 102 M-1 s-1, respectively (Table S4). The N168D/D183N variant is the only PON1 variant tested in which a complete loss in activity was observed (Figure 5). An additional negative charge was introduced into the active site in order to determine if loss of the negative charge at D269 could be compensated for by additional negative ligating residues. This led to the creation of N168D/N224D/D269N, giving the active site an overall charge of -2. But this variant was nearly inactive, with minimal or undetectable activity for all substrates


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Biochemistry

except TBBL, which decreased three orders of magnitude. The last mutant made in this set was a mutant that completely reversed the charges in the core four residues, moving negative charges from E53 and D269 to N168 and N224 respectively. The resulting E53Q/N168D/N224D/D269N mutation had a nearly identical profile to that of the triple mutant, maintaining only activity against TBBL three orders of magnitude below WT (Figure 5). Net charge alone is clearly not sufficient to maintain high activity, and variants with similar net charges vary considerably. There is a multitude of possible effects on Ca2+ binding, the electronic structure around the metal, and how the ligating residues interact with near neighbors. But our results make it unlikely that E53 and D269 are uniquely mechanistically important, or on the other hand that Ca2+ binding alone is sufficient for high level activity. The effect of a higher calcium concentration was also examined with several charge balance

variants

(D269N,

D269E,

E53Q/N168D,

N168D/D269N,

N224D/D269N,

N168D/N224D/D269N, and E53Q/N168D/N224D/D269N). The enzymes were dialyzed into assay buffer that contained 10 mM CaCl2 (instead of 1 mM used in all other assays) and activity assays were carried out using this buffer. The resulting catalytic efficiencies were nearly all within an order of magnitude of the 1 mM CaCl2 assays (data not shown). The only significant difference observed was for the E53Q/N168D variant, which exhibited slightly decreased activity against PA (~20x) but a slight increase in activity against CMP (~14x). The His-dyad (H115/H134) on the side of the active site has also been proposed to participate directly in hydrolysis, particularly for lactones and esters.25 The ablation of lactonase and esterase activity in H115 mutants has led to the suggestion that two mechanisms might be at work in the PON1 active site. To investigate this, sterically conservative His to Phe mutations were made in PON1. Surprisingly, the H115F variant had a fairly modest effect on catalysis, with


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minimal effects on CMP and CPO hydrolysis, a single order of magnitude decrease in DHC, PX, and a three order of magnitude decrease in TBBL and PA hydrolysis. This mutation improved PON1’s ability to hydrolyze DFP, increasing its activity by a single order of magnitude (Figure 6). Individual kinetic parameters (kcat and Km) were able to be obtained for H115F against DHC, TBBL, CMP, PX, and CPO. The Km was adversely affected against DHC, CMP, and PX, increasing approximately three-fold. But the Km decreased against CPO and TBBL, about two-fold and to determinable levels against CPO and TBBL (Table S4). The H134F mutation had a slightly greater effect, decreasing CMP, PX, and CPO hydrolysis two orders of magnitude, and decreasing TBBL hydrolysis four orders of magnitude, while having the same effect as the H115F mutation on DHC, PA, and DZO hydrolysis (Figure 6). This mutation increased the Km enough to prohibit the calculation of individual kinetic parameters except in the case of PX, for which Km was increased by about 1.5-fold (Table S4).


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Biochemistry

Figure 6. Six-bladed β-propeller hydrolase wild type and “His-dyad-like” variant activities. A. Lactonase and aryl esterase catalytic efficiencies. B. Phosphotriesterase catalytic efficiencies. See notes in Fig. 4.

The H115F/H134F double mutant, on the other hand, had a surprisingly small effect on catalysis, with activity closer to the H115F variant. The CPO and DZO activities were only minimally affected, DHC activity was decreased only a single order of magnitude, while CMP activity was decreased two orders of magnitude, and TBBL and PA activity were decreased three orders or magnitude each. PX activity increased by an order of magnitude to 1.3 × 104 M-1 s-1 and


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DFP hydrolysis increased more than either single mutant to 5.2 × 104 M-1 s-1 (Figure 6). Additionally, individual kinetic parameters were able to be determined for DHC, PX, and CPO for the double mutant, showing lower Km values against PX and CPO (three-fold) but a higher Km for DHC (three-fold). The kcat was significantly decreased for all three substrates (Table S4). Finally, the double mutant (H115F/H134F) was combined with D269N from PON1 to make the triple mutant H115F/H134F/D269N in order to remove all hypothesized catalytic residues. Surprisingly, this variant showed a two order of magnitude decrease in DHC and PA activity, a three order of magnitude decrease in TBBL activity, and a near complete loss of phosphotriesterase activity (Figure 6). For DHC, there was a large decrease in kcat (~200-fold) but only a small increase in Km (Table S4). DFPase, Drp35 and SMP30 do not have a histidine in sequence or three-dimensional space analogous to PON1 H115. We wondered if mutations analogous to those produced at the His residues of PON1, or whether inserting His residues in a PON1 H115-like position, would have an effect on activity (Figure 2B). In DFPase, H287 is on a different side of the active site, but in a similar position above the bound cation to that of H115 in PON1. It has previously been suggested that H287 may participate in catalysis, perhaps in the same manner as PON1 H115F.25,42,52 As a result, H287F was made and yielded only a small effect on catalysis, decreasing DHC, TBBL, and CPO activity a single order of magnitude while losing only five-fold activity against DFP. Interestingly, most of the GL activity was lost for this variant relative to WT (Figure 6). This set of values indicates that H287 may not participate in catalysis, but may play a role similar to H115 in PON1. There are alanines present in DFPase and Drp35 that have a backbone position very similar to that of PON1 H115. Additionally, in SMP30 the side chain of R101 protrudes into the active site where we would expect to see the side chain of PON1 H115 (Figure 2B,C). The resulting


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Biochemistry

A74H, A90H and R101H mutations were made in DFPase, Drp35, and SMP30, respectively. DFPase A74H was the only active variant; while this mutation did not rescue any activity, it did maintain WT-like activity against DHC. Minimal activity was observed for DFPase A74H and Drp35 A90H against CPO, but the activity levels were too low to determine a value. The R101H variant of SMP30 also maintained minimal GL activity (Figure 6). These data together with the PON1 His-dyad mutagenesis data indicate that this group of residues likely does not participate directly in catalysis (especially considering DFPase and Drp35 lack the residues to do so), but may play an important role in substrate binding or orientation.

DISCUSSION Despite very little sequence homology, the six-bladed β-propeller hydrolases share a highly conserved active site and hydrolyze a wide range of substrates. Previous work has suggested that their remarkable ability to tolerate different classes of substrate stems from their adaptable active site tunnels.24,26,30,34,39,53 However, it is still unclear if these four hydrolases share a common mechanism, and, if they do, what that mechanism is. Here, we conducted a comprehensive interrogation of the six-bladed β-propeller hydrolase active sites, where critical variants were tested against a panel of esters, lactones, and organophosphates to uncover trends that could lead to a stronger mechanistic hypothesis. We first tested the wild-type hydrolases (PON1, DFPase, Drp35, SMP30) against the ester, lactone and OP substrates shown in Figure 3. PON1 was also the only enzyme capable of simple ester hydrolysis, and is capable of hydrolyzing all three substrate classes. DFPase was the only other enzyme capable of significant OP hydrolysis. Previously, SMP30 has been observed to hydrolyze DFP, as well,54 but we were unable to detect that. All four of these enzymes have been


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suggested as a possible lead for an OP poisoning therapeutic, but the broader OP activity of PON1suggests it is the best choice.39,47,53,55 In agreement with literature, SMP30 and Drp35 exhibited GL and DHC hydrolysis, respectively,34,54,56 and we show for the first time that SMP30 is also capable of DHC hydrolysis. In addition to DHC, Drp35 is weakly capable of TBBL hydrolysis. The only substrate class that all four enzymes hydrolyze is the lactones. The OPase activity of these enzymes may arise from an overlap in the transition states leading to lactone and OP hydrolysis that is not necessarily present in ester hydrolysis. It has been hypothesized these proteins may have originated from a common ancestor and were transferred to other organisms through horizontal gene transfer (from mammals).57–61 Differences in substrate specificity may have come from thousands of years of evolution resulting in specialization for certain functions.39,52,57–61 It is difficult to know how much substrate binding versus mechanism leads to the extant specificities, but we believe that the common lactonase activity suggests the ancestral, if not contemporary, function of these four enzymes. An examination of the DFPase and PON1 binding pockets in complex with inhibitors sheds some light on their promiscuous activities.53 The binding pocket for DFPase is fairly static22 and highly complementary for DFP or a similar substrate in Loligo vulgaris (Figure S1). We would expect that a planar substrate such as DHC would be able to enter this active site pocket, but it may not make sufficient contacts for tight binding or proper positioning. TBBL is a small, nearly planar lactone, but the flexible thio-alkyl “tail” might introduce steric clashes. We have also shown that DFPase also hydrolyzes the OP CPO, but not DZO. On the periphery of the binding pocket and in the base of the active site there are cavities that might accommodate the three groups, the specifics of binding must be remarkably highly tuned. Studies have suggested that DFPase has a relatively rigid active site and entrance tunnel, but that the opposite is true of PON1’s active


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Biochemistry

site.26,37,61 There are also fewer residues surrounding the active site tunnel (Figure S2), possibly leading to lower substrate selectivity. This may account for PON1’s ability to hydrolyze OP substrates with larger leaving groups, mostly in contrast to the other three enzymes here. Interrogation of D269 (PON1) and corresponding residues in DFPase, Drp35, and SMP30 demonstrates its importance to the enzymes’ catalytic ability. Upon a conservative mutation to asparagine, PON1 loses activity against all substrates tested except for the lactones DHC and TBBL (where activity decreases four orders of magnitude). The AspGlu mutation also resulted in a significant loss in activity. The D269E mutant only retained activity against DHC, TBBL, and CMP, with most residual activity against lactones. The AspAsn mutation is not tolerated in any of the other hydrolases, corresponding to a complete loss in activity against all substrates. Of course, the other hydrolases showed poorer hydrolytic ability overall, so it is not surprising that less residual activity can be detected. Additionally, this aspartic acid has been shown in all four cases to be responsible in part for calcium binding.33,35,38,47 On the other hand, when mutating the neighboring residue E53 (PON1) and its counterparts in the other hydrolases, we see a less detrimental effect. While we still observe a complete loss in activity in DFPase, Drp35, and SMP30 (likely for the same reason noted above), PON1 retains significant activity against DHC, TBBL, PA, and CPO. This aligns well with previously proposed water-activation mechanisms, where it has been hypothesized that E53 may act to stabilize a transition state. In this mechanism we wouldn’t expect a conservative mutation such as GluGln to have as severe effect on enzymatic activity, if this is indeed this residue’s role. Strikingly, when these two mutants are combined in PON1 (E53Q/D269N) we do not observe an additive effect (Figure 5, Table S2). While both residues are important for enzymatic activity, we have shown that an acid-free active site mutant (E53Q/D269N) retains activity against


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all three substrate classes (all five substrates). Additionally, this acid-free variant generally exhibits the same or improved activity when compared to the E53Q single mutant (i.e., E53Q/D269) as well as the E53Q/D269E variant. This is contrary to previously proposed hypotheses that state the addition of potential general bases should improve activity of PON1.25 The activity of the Gln/Asn variant argues strongly against a highly specific role for these residues as the active-site nucleophile or direct water activation, and it also suggests that the Ca2+ ion is probably retained even with all carboxamide ligands. We also set out to test the importance of charge balance among the four coordinating residues around the calcium of PON1 in a manner similar to a previous study of DFPase.37,43 We define the canonical (summed) overall charge of the residues in and around the PON1 active site to be -1 (Figure S3). Notably, the N168D/D183N variant (-1 overall charge, WT-like) results in a complete loss in activity while E53Q/D269N (+1 overall charge) retains significant activity toward all substrates examined, indicating a net charge of -1 alone is not sufficient for high activity. On the other hand, mutants with changes to the overall charge can still retain significant activity, as observed in the E53Q/D269N (+1 overall charge) and E53Q/D269E (neutral overall charge) variants. This indicates while the overall charge of the active site may not be critical, alterations of the active site may instead result in altered binding (as previously shown25) or chemical environment for the calcium. It could also result in altered positioning of residues critical for hydrolysis around the calcium. Comparing these results to previous charge balance work on the DFPase active site,37,43 we see a similar conclusion. When a neutral overall active site charge is assumed (by only taking into account catalytic calcium coordinating residues), both addition and removal of charges around the metal significantly perturb (if not eliminate) the activity of DFPase. Furthermore, variants which retained the overall net neutral charge in the active site (as in the WT


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Biochemistry

DFPase) still lost all activity while retaining the ability to bind the catalytic calcium. Additionally, when using increased calcium concentrations in activity assays, no significant changes were observed in the catalytic efficiencies of the variants tested, implying that 1 mM Ca2+ was saturating. Our results combined with those of Blum et al. show that overall charge is not the sole determining factor for hydrolase activity. Rather, an ideal steric, geometric, and electronic environment needs to be maintained for productive substrate binding and subsequent turnover. Finally, we turned our attention to the role of the His dyad that has been suggested at least in the esterase/lactonase mechanism of PON1. We began by mutating the histidines to phenylalanine, sterically similar but not nucleophilic or hydrogen bonding. We observed a surprisingly small effect on catalysis across all substrates tested. The H115F mutation was far less detrimental than H134F, overall. This contrasts previous work where the same positions were mutated to glutamine. Tawfik et al. found that the H115Q mutant exhibited much stronger effects on OPase, esterase, and lactonase activities than H134Q, with the exception of DHC which exhibited the same trend seen here.25 With the H115F/H134F double mutant, we again observed surprisingly high activity against all substrates, and even increased activity toward DHC, DFP, PX, and CPO, compared to the single mutants. This aligns with the previous observation that the H115Q/H134Q double mutant retained significant levels OPase, esterase, and lactonase activity.25 Together, these results show that the His-dyad in PON1 is not necessary for catalysis, but may interact with the bound substrate and help position it properly for attack. Finally, we combined the H115F/H134F mutant with D269N to determine if the enzyme could function without any of the suggested activating residues. This triple mutant still turns over both lactones and aryl esters, but loses activity against OP’s (Figure 6). Therefore we conclude that H115 and D269 can be ruled


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out as direct nucleophiles or water activating residues. The data also strongly suggest a steric role for H115 and H134, participating in substrate binding and orientation. We identified residues aligning with H115 or H134 in PON1 in the other hydrolases (Figure 6). DFPase, Drp35, and SMP30 appear to lack a residue directly analogous to H115. We instead explored the role of residues in DFPase, Drp35, and SMP30 that had similar backbone and side chain positions to that of H115 in PON1. A74, A90, and R101 (respectively) were identified as potential candidates for a mutation to histidine, to see if we could rescue any PON1-like activity in these enzymes. The only variant to retain any activity was DFPase A74H against DHC. There are clearly many reasons why this simple idea might not work, but it rules out the recovery of activity by simply adding a nucleophilic, H-bonding residue in a similar position. DFPase H287F was made to see if it had any similar effect to H115F in PON1. When tested against the panel of substrates, this variant retained activity against DHC, CPO, and DFP. This effect is similar to that of what we see in PON1 H115F. Examination of the binding pocket of DFP (Figure S1) shows that this residue has the potential to come into direct contact with the substrate as it enters the active site, just as H115 in PON1 does. We can conclude that these residues probably play crucial roles in binding and orienting the substrate, thus having an effect on turnover, but likely do not participate directly in catalysis.

CONCLUSION Across the six-bladed β-propeller hydrolases studied here, none of the previously proposed activating residues appear to be singularly critical for hydrolysis. Key aspartic acid, glutamic acid, and histidines (when present) have significant effects on activity in the hydrolases, but our data appear to rule them out as direct participants as previously envisioned. The mild role of the H287F


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Biochemistry

mutant in DFPase, the lack of an H115 analog in other enzymes, and the high residual activity of H115, H134, and D269 single, double and triple mutants, and D269/E53 double mutants all support this conclusion. Mutations around the catalytic calcium in all hydrolases also have large effects on activity, arguing that the electronic and geometric environment of the metal is very important. Taken together, these data suggest that electrophilic activation of the substrate by the catalytic calcium is extremely important, It would also be consistent with the data for residues in the immediate vicinity to help perturb the pKa and activate bound or nearby water molecules.62–64 However, no specific residue has been identified as uniquely responsible for this. One mechanism that is consistent with the available data is that a metal-bound water with depressed pKa (by calcium and/or surrounding residues), when deprotonated, may either directly attack the substrate, or may deprotonate another nearby water which in turn acts as the nucleophile (Figure 7). There is evidence for such mechanisms in other metallohydrolases.62–65 There is considerable work required to affirmatively demonstrate elements of this possible mechanism, such as metal ion replacement, 18O-water hydrolysis studies, and structural elucidation of some of the key mutants presented here.

Figure 7. Proposed Ca2+-mediated water activation mechanisms. Lewis acid activation of the carbonyl (or, in OPs, phosphoryl) group and metal ion mediated activation of water are most


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consistent with the mutational data presented here. Ca2+ may activate a water directly (blue arrows), or it may activate a nearby water that is not directly bound to the ion (red arrows).

Supporting Information Sequences and accession identification numbers for each WT enzyme characterized (Figure S0), DFPase binding pocket (Figure S1), PON1 binding pocket (Figure S2), PON1 (G3C9) active site charge balance map (Figure S3), Active site alignment and individual active site arrangements of the six-bladed β-propeller hydrolases (Figure S4), PON1, DFPase, Drp35, and SMP30 wild-type, Glu, and Asp variant activities (Table S1), PON1 charge balance variant activities (Table S2), PON1, DFPase, Drp35, and SMP30 “His-dyad-like” variant activities (Table S3), Kinetic parameters of PON1 variants which achieved substrate saturation (Table S4), Primers used for cloning of characterized variants (Table S5), Enzyme Concentrations Used in Activity Assays (Table S6)

Accession Codes G3C9 Paraoxonase-1: AAR95986 (GenBank), 1V04 and 3SRG (PDB) Diisopropylfluorophosphatase: Q7SIG4 (UniProtKB), 3O4P (PDB) Drug responsive protein 35: Q7A338 (UniProtKB), 2DG1 (PDB) Senescence marker protein 30: Q15493, 3G4E (PDB)

Author Information Corresponding Author E-mail: [email protected]


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phone +1 (614) 247-8425 Author Contributions The manuscript was generated through contributions of all authors. T.J.G., K.D., D.G.M. and T.J.M. designed the research. T.J.G., D.G.M, and S.M. conducted the research. T.J.G. and T.J.M. analyzed the experimental data. T.J.G. and T.J.M. wrote the manuscript, and all authors edited and approve the manuscript. Funding NIH U54 NS058183; NIH R21 NS084899 Notes The authors declare no financial conflicts of interest.

Acknowledgements Funding for this work was generously provided by NIH grants U54 NS058183 to T.J.M. through a subcontract to OSU from USAMRAA, and to R21 NS084899 to TJM. We gratefully acknowledge the Christopher M. Hadad group at The Ohio State University and the Dan S. Tawfik group at The Weizmann Institute of Science for contributions of reagents to this work. We also appreciate many productive discussions with Dr. David P. Bowles and Dr. Nicholas E. Long.

Abbreviations


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OP(s): organophosphorus compound(s); PON1: paraoxonase-1; DFPase: diisopropylfluorophosphatase; Drp35: drug responsive protein 35; SMP30: senescence marker protein 30; AChE: acetylcholinesterase; BChE: butyrylcholinesterase; HDL: high-density lipoprotein; DHC: dihydrocoumarin; TBBL; 5-thiobutyl butyrolactone; PA: phenyl acetate; CMP: O-cyclohexyl-O-(3-cyano-4-methyl-7-coumarinyl)-methylphosphonate; CPO: chlorpyrifos oxon; DZO: diazinon oxon; PX: paraoxon; DFP: diisopropylfluorophosphate; GL: d-gluconic acid δ-lactone;

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For Table of Contents Use Only

Insights into the mechanism of paraoxonase-1: Comparing the reactivity of the six-bladed βpropeller hydrolases Timothy J. Grunkemeyer1,2, David G. Mata1, Kiran Doddapaneni1, Srividya Murali1, Thomas J. Magliery1,*


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Proposed mechanisms for the six-bladed β-propeller hydrolases. (A) PON1 proton shuttling mechanism implicating the H115-H134 dyad in abstraction of a proton from water leading to attack on the bound lactone.21 (B) PON1 water activation mechanism proposed involving D269. D269 abstracts a proton from a nearby water, leading to attack of the substrate by the generated OH-.28 (C) PON1 mechanism implicating D269 in direct attack of substrate. A newly generated OH- is free to attack the acyl-enzyme intermediate which has been formed, to regenerate the enzyme and release the product.28 (D) DFPase D229 direct attack mechanism proposed to follow the same reaction pathway as the PON1 D269 direct attack mechanism shown in (C).40 (E) The mechanism for DHC hydrolysis by Drp35 implicates two Asp residues, D236 (homologous to D269 and D229 in PON1 and DFPase) and D138 (homologous to N168 and N120 in PON1 and DFPase). D138 initiates the reaction by abstracting a proton from a nearby water, leading to attack on the carboxyl center by the generated OH-.31 161x62mm (600 x 600 DPI)


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Alignment of proteins and active sites. Global backbone (A) and active site (B,C) alignment of DFPase (PDB ID: 3O4P, blue), Drp35 (PDB ID: 2DG1, green), G3C9 PON1 (PDB ID: 1V04, red), and SMP30 (PDB ID: 3G4E, purple). Residues of interest are shown in sticks and Ca2+ are shown in spheres. Conserved residues are labeled in black, non-conserved residues are labeled in the respective protein’s colors. Rendered with PyMOL. 177x59mm (300 x 300 DPI)


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Substrates used for activity assays. The method of observation of turnover is noted (wavelength refers to where we are observing the formation of the hydrolysis product). Abbreviations and method of reaction tracking are noted. 83x80mm (600 x 600 DPI)


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Six-bladed β-propeller hydrolase wild type, active site Asp, and Glu variant activities. A. Lactonase and aryl esterase catalytic efficiencies. B. Phosphotriesterase catalytic efficiencies. All values presented in M-1 s-1 were obtained using a linear fit to the rate versus initial substrate data. Error bars indicate standard error to a linear fit (except variants listed in Table S4, for which kinetic parameters and errors were obtained by fitting data to the Michaelis Menten equation using the graphing software Kaleidagraph). *Indicates weak activity was observed, but a value was not calculated as it was near the detection limit of the experiment. 170x150mm (300 x 300 DPI)


Biochemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

G3C9 PON1 wild type and charge balance variant activities. See notes in Fig. 4. 170x72mm (300 x 300 DPI)


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Six-bladed β-propeller hydrolase wild type and “His-dyad-like” variant activities. A. Lactonase and aryl esterase catalytic efficiencies. B. Phosphotriesterase catalytic efficiencies. See notes in Fig. 4. 170x150mm (300 x 300 DPI)


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Proposed Ca2+-mediated water activation mechanisms. Lewis acid activation of the carbonyl (or, in OPs, phosphoryl) group and metal ion mediated activation of water are most consistent with the mutational data presented here. Ca2+ may activate a water directly (blue arrows), or it may activate a nearby water that is not directly bound to the ion (red arrows). 170x55mm (300 x 300 DPI)


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Insights into the mechanism of paraoxonase-1: Comparing the

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