Catalytic bioscavengers as countermeasures against organophosphate nerve agents

Catalytic bioscavengers as countermeasures against organophosphate nerve agents

Chemico-Biological Interactions 292 (2018) 50–64 Contents lists available at ScienceDirect Chemico-Biological Interactions journal homepage: www.els...

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Chemico-Biological Interactions 292 (2018) 50–64

Contents lists available at ScienceDirect

Chemico-Biological Interactions journal homepage: www.elsevier.com/locate/chembioint

Catalytic bioscavengers as countermeasures against organophosphate nerve agents

T

Moshe Goldsmith∗, Yacov Ashani Dept. of Biomolecular Sciences, Weizmann Institute of Science, Rehovot, Israel

A R T I C LE I N FO

A B S T R A C T

Keywords: Organophosphates Nerve agents Bioscavengers Intoxication Enzymes Medical countermeasures

Recent years have seen an increasing number of incidence, in which organophosphate nerve agents (OPNAs) have been used against civilians with devastating outcomes. Current medical countermeasures against OPNA intoxications are aimed at mitigating their symptoms, but are unable to effectively prevent them. In addition, they may fail to prevent the onset of a cholinergic crisis in the brain and its secondary toxic manifestations. The need for improved medical countermeasures has led to the development of bioscavengers; proteins and enzymes that may prevent intoxication by binding and inactivating OPNAs before they can reach their target organs. Noncatalytic bioscavengers such as butyrylcholinesterase, can rapidly bind OPNA molecules in a stoichiometric and irreversible manner, but require the administration of large protein doses to prevent intoxication. Thus, many efforts have been made to develop catalytic bioscavengers that could rapidly detoxify OPNAs without being inactivated in the process. Such enzymes may provide effective prophylactic protection and improve post-exposure treatments using much lower protein doses. Here we review attempts to develop catalytic bioscavengers using molecular biology, directed evolution and enzyme engineering techniques; and natural or computationally designed enzymes. These include both stoichiometric scavengers and enzymes that can hydrolyze OPNAs with low catalytic efficiencies. We discuss the catalytic parameters of evolved and engineered enzymes and the results of in-vivo protection and post-exposure experiments performed using OPNAs and bioscavengers. Finally, we briefly address some of the challenges that need to be met in order to transition these enzymes into clinically approved drugs.

1. Introduction Current medical countermeasures for treating organophosphate nerve agent (OPNA) intoxication include: atropine, oxime reactivators, and anticonvulsants. If applied in time, they can prevent lethality and mitigate the intoxication symptoms. However, they may fail to prevent a cholinergic crisis that would lead to loss of consciousness and permanent brain damage [1–3]. In addition, they are not suitable as preventive measures prior to intoxication, as these drugs produce severe side effects such as: CNS impairment, increased blood pressure and increased heart rate if administered prior to an OPNA intoxication [4]. In fact, apart from the stoichiometric bioscavenger human butyrylcholinesterase (HuBChE), purified from human blood, there is no available prophylactic treatment that can prevent OPNA intoxication and the onset of its symptoms. In recent years, catalytic bioscavengers have been proposed as the next generation of medical countermeasures that may allow efficient prophylactic protection from high doses of OPNAs using small doses of protein [5,6].



Catalytic bioscavengers are enzymes that can detoxify OPNAs by performing multiple cycles of OPNA binding and hydrolysis. They have a clear advantage over stoichiometric ones since unlike the latter, which inactivate OPNAs by binding to them irreversibly, their interaction with OPNAs results in reversible binding and rapid hydrolysis of the OPNAs. In principle, this should enable small amounts of catalytic scavenger to detoxify lethal doses of nerve agents in-vivo before the latter can inactivate acetylcholinesterase (AChE) at important physiological sites; and to afford protection from multiple OPNA exposures without being consumed. In contrast, the high molecular weight of stoichiometric scavengers such as butyrylcholinesterase (BChE) or AChE and the requirement for a one-to-one ratio of non-catalytic scavenger to OPNA molecule, imply that large protein doses (i.e. hundreds of mgs) are required to provide protection from OPNA intoxication using stoichiometric scavengers [7,8]. Use of high protein doses of BChE or AChE for protection can be costly [9], and may also increase the chances of adverse physiological reactions following their administration [10]. Thus, during the past decade an increasing number of

Corresponding author. Department of Bimolecular Sciences, Weizmann Institute of Science, Rehovot 7610001, Israel. E-mail address: [email protected] (M. Goldsmith).

https://doi.org/10.1016/j.cbi.2018.07.006 Received 20 May 2018; Received in revised form 4 July 2018; Accepted 6 July 2018 Available online 07 July 2018 0009-2797/ © 2018 Published by Elsevier B.V.

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esterase, endowed with a high turnover rate for OPNA binding and hydrolysis, are great. Such a bioscavenger would have all the advantages of a stoichiometric scavenger (e.g. high OPNA binding rates, long circulatory residence times, biocompatibility), without the loss of activity that follows its interaction with OPs. This has prompted many attempts to design or engineer B-esterase variants that could perform multiple turnovers with OPNAs, alone, or in combination with specific reactivators.

efforts have been devoted to the development of catalytic bioscavengers [5,6]. Pseudo-catalytic bioscavengers are enzymes or combinations of enzymes and chemical reactivators that can jointly perform multiple cycles of binding and hydrolysis of organophosphates. The term usually refers to stoichiometric scavengers from the family of B-esterases, such as AChE, BChE or carboxylesterase (CaE) [11], complexed with an oxime reactivator in their active site. Following the interaction of the enzyme with an OP inhibitor, it may undergo two consecutive reactions: First, covalent binding of the OP to its catalytic serine residue. Second, hydrolysis and release of the inactivating OP [12]. The rates of spontaneous detachment of the inhibitor from the catalytic serine depend on the type of enzyme and OP, but are usually very slow for Besterases interacting with OPNAs. Therefore, external nucleophiles (i.e. chemical reactivators) are required to turn B-esterases into pseudocatalytic OP hydrolyzing systems [13–15]. There are several advantages to pseudo-catalytic bioscavengers: First, their protein component is normally present in the circulation (e.g. BChE [16] in human serum and AChE [17] on red blood cells). Second, they rapidly sequester all types of OPNAs [18] as the latter are designed as cholinesterase inhibitors in the first place. Lastly, their reactivating oximes can reach inhibited AChE in peripheral nerve tissues and in some cases, in the central nervous system (CNS) [19–21]. However, there are several disadvantages to currently available reactivators: They exhibit narrow OPNA specificities and enable effective reactivation of only a number of OPNAs per reactivator [22,23]. They become ineffective following aging of the OP-cholinesterase bond, which is a rapid process in the case of OPNAs such as soman [24]. They have short circulatory residence times, and need replenishing in cases of continuing OPNA exposures [14]. Finally, the rate of OPNA hydrolysis by oxime-mediated reactivation, even by the most effective combinations of oxime, cholinesterase and OP (e.g. cyclosarin-inhibited HuAChE reactivated with HLö7, kr2 = 9.3 × 104 [M−1min−1] [25]), is ∼2–3 orders of magnitude slower than the hydrolytic rate required to efficiently prevent intoxication (see bellow). Therefore, the current status of pseudo-catalytic scavengers, suggests they are not efficient enough to detoxify OPNAs and prevent the inhibition of AChE at peripheral and CNS sites. A more effective way of preventing OPNA intoxication would be to employ a highly efficient, broad-spectrum catalytic bioscavenger, which could rapidly inactivate the toxic isomers of OPNAs in the circulation. In order to do so using a low enzyme treatment-dose (< 1 mg/ kg), theoretical models predict that catalytic bioscavengers must have a catalytic efficiency (kcat/KM) of ≥1 × 107 [M−1 min−1] with the toxic isomers of OPNAs [26,27]. Using enzymes with higher catalytic efficiencies (i.e. (kcat/KM) of ≥5 × 107 [M−1 min−1]) as prophylactics would detoxify 96% of the OP in the circulation in less than 10 s and provide sign-free protection from intoxication [5,28]. Unfortunately, OPNAs are xenobiotic compounds that serve as promiscuous substrates for natural OP hydrolyzing (OPH) enzymes and are hydrolyzed with low catalytic efficiencies [29,30]. In addition, most OPH enzymes isolated so far tend to bind and inactivate the less toxic isomers of OPNAs more efficiently than the toxic ones [31,32]. Therefore, in order to obtain catalytic bioscavengers that could be used as effective medical countermeasures for OPNA intoxication, there is a need to greatly enhance both the activity and selectivity of natural enzymes towards the toxic isomers of OPNAs. In recent years, directed evolution and protein engineering techniques have been used successfully to generate catalytic bioscavengers with such properties [33–38]. Here we aim to discuss proteins that have been suggested as candidates for catalytic bioscavenging, their activities and the efforts made to increase their OPNA hydrolyzing capabilities.

2.1. Carboxylesterases Carboxylesterases (CaEs) are a ubiquitous family of esterases (EC 3.1.1.1) that hydrolyze structurally diverse carboxylic esters to their corresponding alcohols and carboxylic acids [39,40]. Their activity, invivo, can either activate or inactivate compounds, but since their endogenous substrates are unknown, they are considered to play a protective physiological role by detoxifying xenobiotics [40]. Of the 5 human isoforms of CaE, only two have been extensively studied: Human carboxyl esterase 1 (HuCaE1) that is present mostly in human liver, macrophages, and lung epithelia and human intestinal carboxylesterase (HuiCaE) that is expressed especially in the small intestine, kidney, heart and skeletal muscle [41]. HuCaE1 is a 60 kDa protein that possess a much larger active-site pocket relative to HuAChE and can bind larger substrates [41]. It catalyzes ester hydrolysis using a catalytic triad (Ser221, Glu353, His 464) and was suggested to have a “side door” to facilitate the exit of its hydrolysis products [42]. CaEs lack a catalytic site tryptophan residue that is conserved in the cholinesterase family (e.g. Trp 86 in HuAChE), and so the enzyme reacts slowly with charged OPs and oxime reactivators. The abilities of CaEs to bind and detoxify OPNAs and pesticides were investigated primarily using rodent CaEs such as mouse or rat plasma carboxylesterase (CES1C) [43–45]. CES1C binds uncharged OPNAs such as sarin and soman, and pesticides such as paraoxon and DFP more rapidly than charged OPs such as VX. Both HuCaE and CES1C do not undergo aging regardless of the structure of the inactivating OP [44,46]. HuCaE1 is completely inhibited by soman and cyclosarin and undergoes very slow, spontaneous reactivation following inhibition by sarin with a half-time of 45 h [46]. Attempts to increase the reactivation rates of OPNA-inhibited CaEs using oxime nucleophiles, revealed that they react poorly with charged oxime compounds and much better with uncharged ones [46,47]. However, even in the presence of uncharged oximes, the half-life time for reactivation of HuCaE1 (t1/2) was too slow to promote rapid catalytic detoxification (e.g. the t1/2 of serine-inhibited HuCaE1 in the presence of 2,3-butanedione monoxime is 41 min [46]). Investigations of insect resistance to OP pesticides have revealed that while some insects overproduce esterases, such as CaE, to elicit resistance by sequestering the intoxicating OP [48], others have evolved mutants of CaE that are capable of hydrolyzing the pesticides [49,50]. In the case of the E3 CaE isozyme from L. cuprina, a single active site mutation, G137D, was sufficient to confer OP resistance by acting as a general base catalyst that can hydrolyze the OP-bound enzyme [50,51]. To enhance the OPNA hydrolase activities of HuCaE1, two active-site amino-acid substitutions were rationally designed based on the crystal structure of the enzyme. They were aimed to facilitate the reactivation of the OP-bound catalytic Ser 221 residue via the positioning of a water molecule in its vicinity [52]. Indeed, the V146H/ L363E HuCaE1 mutant replaced two hydrophobic residues on opposite sides of the active site pocket with charged residues that increased the reactivation rates of the enzyme without reducing its affinity to the OPNAs [52]. As a result, the reactivation rates of sarin, soman, and cyclosarin-inhibited HuCaE1 increased by 5-, 20-, and ∼33,000-fold and their half-life times decreased to 9.5 h, 11.5 h and 1.2 h respectively [52]. This CaE mutant was significantly improved in reactivation rates relative to the wt protein, and was able to hydrolyze the toxic isomer of a cyclosarin analog at a rate of 5.3 × 104 [M−1 min−1]. However, this

2. Turning stoichiometric bioscavengers into catalytic ones The antidotal and therapeutic potentials of a human B-family 51

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rate is still ∼3 orders of magnitude slower than the rate required from an effective catalytic bioscavenger. A different attempt to enhance the OP hydrolase activities of a CaE employed a structural homolog of HuCaE1 from Bacillus subtilis, p-nitrobenzyl esterase (pNBE), that was modified using rational design and directed evolution methods [53]. This homolog shares a common fold with AChE, BChE and HuCaE1, but different substrate specificities. In addition, pNBE does not contain some of the residues that promote aging of bound OPs in AChE (e.g. W86, F338 [24]) and BChE (e.g. W82, F329 [54]). Unlike human cholinesterases and carboxylesterase, pNBE is a bacterial enzyme that is easier to manipulate and express in bacterial cells. Thus, a library of pNBE mutants was created by targeting 5 residues at the vicinity (7 Å) of the active site Ser residue of pNBE for saturation mutagenesis, in combination with a A107H mutation designed to increase spontaneous reactivation [53]. Following screening with different substrates, a double pNBE mutant (A107H/A190C), which was improved 4000-fold over the wt. protein in its rate of reactivation following soman inhibition, was identified [53]. However, these mutations did not produce a similar improvement when transferred to HuCaE1 [53], and the reactivation rate of the pNBE mutant was still orders of magnitude slower than that of the BChE - G117H variant [55].

purpose, researchers introduced a nucleophilic histidine residue into the oxyanion hole of AChE similar to the G117H mutation in HuBChE, which conferred OP hydrolase activity on BChE [81]. Since the G122HBfAChE mutant had completely lost its acetylcholine hydrolase activity, additional mutations were introduced into enzyme's active site based on a molecular modelling analysis [81]. This analysis suggested that the G122H mutation had distorted the active site by causing steric hindrance. Thus, two additional mutations, Y124Q and S125T, were introduced to increase the space in the active site and to mimic additional BChE residues at this region [81]. The resulting triple mutant H122H/ Y124Q/S125T-BfAChE displayed a 4 order of magnitude reduction in catalytic efficiency with actylthiocholine with respect to wt BfAChE, but was able to slowly hydrolyze the OP pesticides DFP and paraoxon in addition to OP Echothiophate [81]. Still, the efficiency of hydrolysis of these OPs was 1–2 orders of magnitude lower than that of G117HHuBChE and the mutant was suggested to resist OP inhibition mostly due to poor OP binding affinities. An attempt to graft the same set of mutations onto HuAChE did not generate an improved variant and was not further studied [81]. 2.3. Butyrylcholinesterase BChE (EC 3.1.1.8) is a 574 residue (85 kDa) serine hydrolase that is expressed primarily in human plasma, liver, skin and leg muscle and is estimated to be ∼10 fold more abundant than AChE in human adults (see recent review [16]). The physiological role of BChE is yet unknown, but it is generally accepted to have a role in detoxification of neurotoxins that can inactivate AChE and may act to protect it [82]. In addition, it was found to be able to substitute for AChE when the latter does not function [83] and to regulate the hormone ghrelin that regulates food intake [84]. BChE is assembled in-vivo into soluble, globular tetramers both in plasma and when bound to membranes via an interaction of a four-helix bundle tetramerization domain with a polyproline-rich peptide. The activity of BChE depends on the catalytic triad: Ser 198, Glu 325 and His 438 and is inhibited upon binding of Ser 198 to an OP [85]. An omega loop connecting residues Asp70 and Trp82 plays an important role in substrate binding in HuBChE. Phosphorylated HuBChE is reactivated less efficiently with oximes than phosphorylated HuAChE [82]. In an attempt to increase the spontaneous reactivation rates of OPNA-inhibited BChE, several activesite residues were mutated based on rational design and modelling of the enzyme onto the crystal structure of Torpedo californica AChE. One of the variants explored, BChE-G117H, resisted inhibition by sarin and VX 1000- and 7900- fold more than wt BChE and exhibited 100- and 2000-fold faster dephosphorylation rates following inhibition by sarin and VX respectively [86]. To further increase the reactivation rates of BChE, a residue that is important for aging, Glu197, was rationally mutated to abolish its charge [55]. Indeed, unlike the single G117H mutant, the double mutant BChE-G117H/E197Q was capable of spontaneously regenerating its catalytic activity following inhibition by soman. It also displayed improved reactivation rates with sarin and VX, but its overall rates of OPNA hydrolysis were still too slow to afford catalytic bioscavenging [55]. Attempts to find more improved HuBChE mutants by screening 62 rationally designed single, double or triple mutants of G117 failed to obtain better variants [87]. In conclusion, the esterases described above share a number of properties that make them ideal as catalytic bioscavengers: They are endogenous proteins with long circulatory residence times, high OPNA binding affinities, an ideal physiological location for serving as the first line of defense against OPNAs by detoxification (e.g. plasma BChE and Erythrocyte AChE) and they display very high catalytic efficiencies with their native substrates (e.g. AChE and BChE). However, attempts made so far using protein engineering and site-directed mutagenesis methods, have failed to generate variants of these proteins that could hydrolyze the toxic isomers of OPNAs at high catalytic efficiency rates (i.e. kcat/ KM ≥ 1 × 107 [M−1 min−1]) either by themselves or in combination

2.2. Acetylcholinesterase Acetylcholinesterase (EC 3.1.1.7) is an α/β fold, serine hydrolase that terminates the transmission of nerve impulses at cholinergic synapses by hydrolyzing the neurotransmitter acetylcholine (for recent reviews see Refs. [56–58]). It is a highly efficient enzyme that operates at a rate close to diffusion limit (e.g. kcat/KM = 9.6 × 1010 [M−1min−1] with acetylthiocholine [59]) and is expressed in various isoforms [60] and in different tissue (e.g. Ref. [17]). In vertebrates it functions mostly as a membrane-bound tetramer that is associates with either the collagenous protein ColQ or with a transmembrane proline-rich anchor (PRiMA) protein [61]. Both AChE and its paralog BChE rapidly bind OPNAs with bi-molecular rate constants (ki) of ∼107-1010 [M−1 min−1], display slow or no spontaneous reactivation kinetics and are permanently inactivated following aging of the enzyme-OPNA complex [13,62–64]. Since AChE is the physiological target of OPNAs, it has been a prime target for structural and functional investigations aiming, among other things, to identify amino-acid substitutions that could promote its reactivation following OP binding. A number of insect species have been shown to resist OP pesticide toxicity by evolving mutated forms of AChE [65–68]. However, these mutants confer resistance by reducing the binding of the OP to the mutant AChE active site and not by promoting the hydrolysis of the bound OP. Attempts to change the binding parameters of AChE with different substrates and OP inhibitors were made using site-directed mutagenesis and recombinant AChE from different sources (e.g. Refs. [69–73]). Unfortunately, these failed to identify mutations that would significantly increase the spontaneous reactivation rates of the OP-bound enzyme. In contrast, a number of site-specific mutations were found to significantly improve the efficiency of oxime-mediate reactivation of AChE following inhibition by OPNAs and to decrease the rate of aging [74–77]. Thus, by combining pretreatment with a PEGylated F338A-HuAChE mutant and post-exposure treatment with an HI-6 oxime, mice were shown to survive two sequential intoxications with high doses of soman (i.e. 5.3 LD50 and then 4 LD50 [78]). However, this kind of treatment still relies on the use of high bioscavenger doses and specific oxime-AChE mutant combinations for different OPNAs. Unlike HuAChE, AChE obtained from the venom of snakes such as Bungarus Fasciatus (BfAChE; Uniprot Q92035) [79] can be expressed as a monomeric protein, at high levels in mammalian cells [80]. This made it an attractive target for protein engineering for the purpose of improving its reactivation rates following OPNA inhibition. For this 52

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Table 1 Summary of in-vivo protection and post-exposure treatments of OPNA intoxications using catalytic bioscavengers as a stand-alone treatment. OPNA

Exposure rout

OPNA Dose

Bio-scavenger

Protein Dose (mg/Kg)

Timing (min)a

Animal

% survivalb

Intox. Symp.c

Experiment length [days]

Ref.

GB

Inhalation

HuPON1

4

−30

Guinea pigs

67%

I

1

[113]

GD

Inhalation

HuPON1

4

−30

Guinea pigs

75%

I

1

[113]

GB

Inhalation

RPON1

2

−30

Guinea pigs

73%

I

1

[113]

GD

Inhalation

RPON1

2

−30

Guinea pigs

71%

I

1

[113]

rePON1 (4E9)

2.2/1.1

−60

Mice

63%/45%

I/II

14

[26]

rePON1 (IIG1) rePON1(I-F11) rePTE (C23) rePTE (C23AL) DFPase DFPase-Mut Human liver prolidase Human liver prolidase Human liver prolidase

1/0.2 0.3–12g 5/2 1/2

−60 NDh +5/15 +5

Guinea pigs Mice Guinea pigs Guinea pigs

100% 90–100% 100% 100%

I/II I I/II - III III

0.125 4 0.125 0.125

[132] [143] [179] [180]

71 35.8 ND

−5 −5 ND

Rats Rats Mice

50% 100% 50%/0%j

III I ND

>1 >1 1

[208] [208] [230]

ND

ND

Mice

0%/0%

ND

1

[230]

ND

1

[230]

Sp-CMPCoumarind GF GA, GB, GD, GF VX VX

i.v.

1.2 × LC50 (846 mg/m3) 1.2 × LC50 (841 mg/m3) 1.2 × LC50 (846 mg/m3) 1.2 × LC50 (841 mg/m3) 2 × LD50

s.c. s.c. s.c. s.c.

2 5 2 2

GD GD GB

s.c. s.c. s.c.

3 × LD50 3 × LD50 1/2i x LD50

GD

s.c.

1/2i x LD50

GF

s.c.

× × × ×

i

LD50 LD50f LD50 LD50

1/2 x LD50

e

j

ND

ND

Mice

33%/0% j

a

Negative values = time (min) before intoxication. Positive values = time (min) post intoxication. Survival is reported at the end of the experimental observation time. c Intoxication symptoms ranked as: I – light, II – moderate, III – sever. d The toxic isomer of a GF surrogate. e The 24 h survival rate was 75%. f The same mice were sequentially intoxicated with 5 × LD50 doses of GD, GF, GB and GA at 24 h intervals. g The protein was expressed endogenously following infection with and adenovirus expression vector. Protein concentration range was estimated in this table using an average blood volume of 1.65 ml per mouse and the reported range of 0.2–7.6 mg/ml protein concentration obtained in the blood. h ND – not determined. i Two cumulative 1 × LD50 doses were given within a 1 h interval. j Survival is reported after the first and second 1 × LD50 intoxication dose. b

observations and in-vivo experiments: Initially, the susceptibility of animals to OP-pesticide intoxications was found to correlate with their plasma PON1 activities (e.g. low levels in birds, high in rabbits [99,100]). Then, injections (i.v.) of purified rabbit PON1 were shown to protect mice and rats from intoxication by lethal doses of chlorpyrifos (100,150 mg/kg) or chlorpyrifos-oxon (14 mg/kg) [101,102]. Finally, PON1-knockout mice were found to be markedly more susceptible to OP-toxicity [103], but were able to recover their resistance to pesticide metabolites such as diazoxon and chlorpyrifos-oxon following injections of exogenous PON1 [104]. Paradoxically, the catalytic activity of PON1 did not provide in-vivo protection from paraoxon in mice due to its low catalytic efficiency of paraoxon hydrolysis relative to other pesticide metabolites [104]. Attempts to correlate between the in-vivo OP-hydrolyzing activity of HuPON1 and the susceptibility of humans to OP toxicity are complicated by genetic and environmental factors. First, differences in PON1 genetic polymorphs and their expression levels between individuals (e.g. L/M55, Q/R 192, -108T/C allele) result in differences of up to 13fold in serum protein levels and up to 40-fold in plasma PON1 activity [105]. Then, there are substantial inter-individual differences in the capacity to metabolize thion (P=S)-containing, OP pesticides to their active oxon (P=O) forms (e.g. parathion to paraoxon, chlorpyriphos to chlorpyrifphos-oxon) that are the toxic metabolites of these pesticides. These are due to differences in the activities of metabolic enzymes (e.g. P450 cytochromes [106]) and to factors such as: age, sex, health and lifestyle [107]. Finally, individual occupational, secondary or accidental OP exposure levels are difficult to assess. Yet, there are sufficient indications that low level PON1 activity is a risk factor for humans exposed to OPs [105]. The fact that PON1 is a naturally abundant protein in the circulation

with chemical reactivators. Whether achievement of this goal is feasible or not, due to inherent limitations imposed by their active site architecture, is not clear. 3. Enhancing the efficiency of OPNA hydrolysis by natural OP hydrolases 3.1. Serum paraoxonase 1 Serum paraoxonase 1 (PON1; EC 3.1.8.1) is a 355 amino-acid (40 kDa), calcium-dependent hydrolase, structured as a six-bladed βpropeller [88]. It is the most characterized member of the mammalian PON gene family that includes also PON2 and PON3 [89]. Human PON1 (HuPON1) is synthesized in the liver and is secreted as a glycoprotein into the blood where it is associated with cholesterol-carrying HDL particles [90]. Its concentration in human blood can vary significantly between individuals due to genetic polymorphism and physiological state [90,91]. It was first discovered as an OP-hydrolyzing enzyme in animal tissue and was named after its capacity to hydrolyze paraoxon [92,93]. However, it can also hydrolyze a broad range of esters, phospholipids and organophosphates and is most efficient at hydrolyzing lactones [94,95]. The primary physiological role of HuPON1 is considered to be the prevention of low and high-density lipoprotein (LDL, HDL) oxidation by hydrolysis of pro-inflammatory, oxidized phospholipids and hydroxides of cholesteryl linoleate [96,97]. In doing so, PON1 plays an important role in the prevention of atherosclerosis and cardiovascular diseases [98]. The OP hydrolyzing activity of PON1 is a promiscuous activity of the enzyme. Yet, it provides protection in-vivo from the toxicity of OP pesticides and insecticides. This was established following a line of 53

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promoted the survival of ∼50% of the intoxicated animals, with minimal symptoms [26]. A similar administration of wt rePON1 provided no survival and a prophylactic treatment using the standard chemical antidotes, Atropine plus 2-PAM 5 min prior to intoxication, provided only transient survival [26]. A larger dose of 4E9 (2.2 mg/kg) increased animal survival to 75% initially and 63% after 14 days (Table 1). This supported the model that correlated between the catalytic efficiency of the bioscavenger, its concentration in-vivo and its ability to promote survival following intoxication with a specific OPNA dose [26,131]. In a follow-up work that also used directed evolution, the catalytic efficiency of rePON1 was improved by 5- to 3400-fold with the toxic isomers of all G-type nerve agents (i.e. sarin, soman, cyclosarin and tabun) [33]. The most efficient, evolved rePON1 variant (IIG1) contained 8 mutations relative to wt rePON1 and was best at hydrolyzing the toxic isomers of soman and cyclosarin (Table 2) [33]. This variant was injected i. v. as a purified protein at a dose of 1 mg/kg, 1 h prior to the intoxication of guinea pigs by a 2 × LD50 dose of cyclosarin (s.c.); providing complete survival with minimal intoxication symptoms and significant retention of brain ChE activities (Table 1) [132]. Complete survival and retention of brain ChE activities were also obtained when the prophylactic dose of IIG1 was reduced to 0.2 mg/kg, although with more sever intoxication symptoms [132]. The use of a catalytic bioscavenger for post-exposure treatment was tested using a rePON1 variant that was evolved to efficiently hydrolyze the pesticide metabolite of parathion, paraoxon (kcat/KM = 2.2 × 107 M−1min−1). In this work, the evolved variant was shown to promote full survival from a 1.4 × LD50 i. m. dose of paraoxon in rats, when administered together with the enzyme glutamate-oxaloacetate transaminase (GOT), 1 min postintoxication at a 2 mg/kg dose [133]. GOT1 converts glutamate (Glu) and oxaloacetate (OxAc) to 2-ketoglutarate and aspartate. It can be used as a catalytic Glu bioscavenger to rapidly decrease blood and brain Glu concentrations; improving neuronal survival and reducing brain damage following head injuries, ischemic strokes and gliomas [134–136]. The co-administration of purified human GOT1 with OxAc has also been shown to provide significant neuronal protection from OP intoxications in animal models [137,138]. In this work [133], it was administered together with an evolved rePON1 variant and OxAc to improve survival and prevent long-term brain damage following an acute paraoxon intoxication in rats. However, it was concluded that more studies were required to establish its efficacy. G-type nerve agents (i.e. sarin, soman, cyclosarin) have a small and highly reactive fluoride leaving group (pKa = 3.1) that makes them easier to hydrolyze. In contrast, V-type nerve agents, i.e. VX, Russian VX (RVX) and Chinese VX (CVX), have bulky, positively charged and less labile N,N-dialkylaminoethylthiolate leaving groups (pKa = 7.9), which make them harder to inactivate by enzymatic hydrolysis. Indeed, while the catalytic efficiencies of hydrolysis of the toxic isomers of Gtype agents by rePON1 (Table 2) were low, with V-agents they were bellow detection limit (i.e. < 2 M−1 min−1; Goldsmith et al. unpublished data). However, an examination of evolved variants from the previously described directed evolution works [26,33] identified several variants improved at hydrolyzing Sp-VX (kcat/ KM ∼ 102 M−1min−1) [33]. These variants were further evolved to hydrolyze Sp-VX, using directed evolution and computational design methods (Goldsmith et al. unpublished data). However, since the highest catalytic efficiency obtained by an evolved variant (kcat/KM ∼105 M−1min−1) was two orders of magnitude lower than required from an efficient catalytic bioscavenger, these attempts were not pursued further. Another manner of introducing a bioscavenger into animals is by infection with an adenovirus vector that can induce the expression of the protein in-vivo [139]. The adenovirus mediated expression of HuPON1 in mice was found to prevent intoxication by chlorpyrifos oxon [140] and diazoxon [141], and to maintain whole-brain AChE

that can hydrolyze a broad range of OP pesticides [108] and nerve agents such as sarin, soman, tabun and VX [109–112], made it an ideal candidate for development as a catalytic bioscavenger. Indeed, protection experiments using purified human, rabbit and recombinant PON1 (∼2–4 mg/kg, i. v.), introduced as a stand-alone treatment, 30 min prior to intoxication, were able to significantly improve survival rates and recovery following intoxication by 1.2 × LCt50 of sarin or soman in guinea pigs (Table 1) [113,114]. In these experiments, the prophylactic treatment with purified PON1 not only increased survival rates from ∼20% to ∼75%, but also improved blood O2 saturation levels, pulse and respiratory rates and protected ChE activity both in the circulation and in the brain, following intoxication [113,114]. However, attempts to protect guinea pigs from higher doses (i.e. 2 × LD50) of sarin, soman, cyclosarin or tabun proved unsuccessful even when the dose of recombinant HuPON1 was increased to 10 mg/kg and the animals were intoxicated only 5 min post-injection of PON1 [115]. In addition, mice with 10- to 30-fold greater PON1 concentrations in their plasma, following an infection with an adenovirus-PON1 expression vector, were also not protected from 2 × LD50 doses of soman or VX [115]. The fact that other animals in these experiments, were able to survive a 4 × LD50 dose of chlorpyrifos oxon or a 2 × LD50 dose of diazoxon, highlighted the fact that high bioscavenger blood concentration is not sufficient to provide effective prophylactic protection, unless its catalytic efficiency is sufficiently high with the intoxicating OP. The low catalytic efficiencies of HuPON1 with OPNAs prevented it from providing effective protection from lethal nerve-agent doses. Site directed mutagenesis of HuPON1 has been employed to design mutants with enhanced OP hydrolyzing capabilities [116,117]. However, the resulting single and double-mutant variants displayed improvements of only 1.5- to 8.9-fold in hydrolytic efficiency on paraoxon or on racemic mixtures of sarin, soman, tabun and VX [116,117]. The expression of HuPON1 in human 293T embryonic kidney cells precluded high-throughput mutagenesis and screening efforts, and attempts to express it in E. coli cells produced an unstable protein at very low yields [118,119]. In addition, the initial absence of a crystalstructure of PON1 delayed rational design of improved variants. These problems were circumvented following the application of directed evolution methods to PON1. Directed evolution and computational design methods have been successfully used in the past two decades to redesign the properties of natural proteins and enzymes for different purposes (reviewed in Refs. [120–129]). They rely on experimental systems that facilitate genetic manipulations (e.g. gene mutagenesis), expression of protein variants and examination of their catalytic efficiencies and other desired properties. In most cases, unicellular organisms such as bacteria or yeast, or artificial cell-like compartments such as water-in-oil emulsions [130] are best suited for these purposes. In the case of HuPON1, bacterial expression was enabled by selection of a recombinant PON1 variant following gene-shuffling of 4 mammalian PON1 genes from: Human, mouse, rabbit and rat. The resulting protein (i.e. rePON1) expressed as a soluble protein at high yields in E. coli and exhibited similar catalytic properties to wt HuPON1 [118]. Initially, wild type-like rePON1 was used as a template for directed evolution aimed at increasing its catalytic efficiency with OPs using a fluorogenic analog of paraoxon (DEPCyC) [118]. The examination of several evolved variants using: cyclosarin, soman, a sarin analog, DFP and chlorpyrifos-oxon, found variants improved ∼10-fold with cyclosarin and soman and identified the key residues involved [31]. In a later work, rePON1 variants were screened for their ability to hydrolyze fluorogenic analogs of sarin and cyclosarin as well as the in-situ generated nerve agents. Following several rounds of mutagenesis and selection, a rePON1 variant (i.e. 4E9), which was improved by 135-fold at cyclosarin hydrolysis relative to rePON1, was isolated (Table 2) [26]. When a low dose (1.1 mg/kg) of purified 4E9 was administered to mice as a prophylactic treatment, 1 h before intoxication with a 2 × LD50 dose of the toxic isomer of a cyclosarin analog (Sp-CMP-coumarin), it 54

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Table 2 Catalytic efficiencies of natural and most improveda OPNA hydrolases. Enzyme

Variant nameb e

GA

PON1 PON1

wt 4E9

0.043 ND

PON1

IIG1

0.23 (5×)

PTE PTE

wtg YT

69 ND

PTE

C23

PTE

d1-IVA1

PTE

GB

GD-fc

GD-sc

GF

VX

RVX −7

ND ND

f

CVX-fd

CVX-sd

Ref.

ND ND

ND ND

[33] [26]

0.008 0.03 (4×) 0.32 (40×)

0.0055 0.74 (135×) 5.1 (927×)

0.0015 0.058 (39×) 5.1 (3400×)

0.013 1.75 (135×) 3.4 (262×)

< 2 × 10 8 × 10−6 (> 40×) ND

ND

ND

ND

[33]

ND ND

[38] [36]

0.5 (500) 0.35 (350×) 5.1 (5100×)

0.07 (1000×) 1.2 (17,143×) 0.27 (3857×)

0.17 (850×) 0.75 (3750×) 0.34 (1700×)

ND

[37]

0.19

[38]

5.5 (6.7×)

0.005 4.8 (960×) 0.46 (92×) 0.15 (30×) 0.32 (64×)

2 × 10−4 ND

12.6 (0.2×)

0.01 3 (300×) 0.83 (83×) 0.11 (11×) 0.07 (7×)

7 × 10−5 ND

10-1-D11

0.1 3 (30×) 0.83 (8.3×) 0.38 (3.8×) 0.07 (0.7×)

0.001 ND

15.8 (0.2×) ND

0.82 12 (15×) 14.8 (18×) ND

0.11

[38]

DFPase DFPase

wt Mut1

ND ND

0.25 1.4 (6×)

ND ND

ND ND

0.1 2.9 (29×)

ND ND

ND ND

ND ND

ND ND

[203] [203]

OPAA OPAA

wt FLYD

ND ND

ND ND

ND ND

ND ND

ND ND

ND ND

5.5 × 10−5 8.9 × 10−4 (16×)

ND ND

ND ND

[222] [222]

Prolidaseh

wt

ND

8 × 10−3

2 × 10−3

3 × 10−3

3 × 10−3

ND

ND

ND

ND

[230]

Hydrolysis of the toxic (Sp) isomers of nerve agents kcat/KM (x107 M−1 min−1); fold improvements relative to the wt enzyme in parentheses. a The most improved and kinetically defined variants, with at least one of the nerve agents listed, per enzyme. b wt – wild type enzyme. c Attributed to the two toxic isomers of GD: SpRc and SpSc; f – fast, s – slow. d CVX was assayed as a racemic mixture. Isomers were identified by rate of hydrolysis; f – fast, s – slow. e Measured using wt-like rePON1 variant G3C9 (see Aharoni, A. et al., PNAS, 2004, 101 (2), 482–487.). f ND – not determined. g Measured using wt –like rePTE variant S5 (see Roodveldt, C. et al., PNAS 2005, 18 (1), 51–58.). h Values reported for recombinant human liver prolidase.

encoding PTE (opd) was found on a plasmid, meant that it could be transferred to other bacterial strains. Indeed, PTE was also found independently in unrelated soil bacteria, isolated from different parts of the world (e.g. Sphingobium fuliginis (former Flavobacterium sp. ATCC 27551) identified in the Philippines [150–152] and in Brevundimonas diminuta and Pseudomonas putida identified in the USA [145,153]). Plasmid pCMS1 from B. diminuta was found to be self-transmissible and responsible for the horizontal transfer of its opd gene between soil bacteria [154]. OP pesticides can become a valuable source of phosphorous for soil bacteria. However, they need to be imported into the cell and degraded to simpler molecules such as phosphoric acid, in order to be utilized for growth and energy. Accordingly, the expression of PTE from B. diminuta was found to be targeted to the inner membrane of the bacteria, where it becomes membrane-bound and associated with phosphatases and with a phosphate transporter [155]. Since the native activity of PTE resulted in rapid hydrolysis of OP pesticides, it was soon recognized as a potential enzyme for OPNA detoxification. The purified enzyme hydrolyzed racemic soman and cyclosarin with moderate efficiencies (i.e. kcat/KM = 4.8 × 106 and 5.8 × 105 M−1 min−1, respectively) [156], but was less efficient with VX (kcat/KM = 4.5 × 104 M−1 min−1) [157] or RVX (kcat/ KM = 6.3 × 104 M−1 min−1) [158]. When used in animal protection experiments, PTE was found to effectively protect mice from multiple lethal doses of paraoxon (i.e. up to 7.3 × LD50, i. v.) and O,O-diethyl fluorophosphates (DEFP) (i.e. up to 2.9 × LD50, i. v.) using low enzyme doses (i.e. 0.2–0.6 mg/kg) administered (i.v.) only 10 min prior to intoxication and without any additional treatment [159]. PTE was shown to protect mice from intoxication by tabun in a similar manner; i.e. from 4 to 5.7 × LD50 doses of tabun, administered (i.v.) 5 min after the prophylactic administration of PTE (0.36 and 0.53 mg/kg PTE, i. v.) without additional treatment [160]. Prophylactic protection, using low

activity. When directly evolved rePON1 variants (i.e. VII-D11, I-F11), from the work previously described [33], were cloned into an adenovirus vector and used to infect mice, they provided complete and symptomless protection from a cumulative 6.3 × LD50 dose of paraoxon (VII-D11) [142] or from 4 repeating doses of 5 × LD50's of either sarin, soman, cyclosarin or tabun (I-F11; Table 1) [143]. The expressed rePON1 variants were found to associate with HDL in the circulation of the infected animals, as does wt PON1, and were expressed for 6–7 days at high concentrations (i.e. up to 4.1 mg/ml) in plasma [142,143]. Thus, while HuPON1 itself is insufficiently active to provide effective prophylactic protection from lethal doses of OPNAs, directly evolved variants of rePON1 have been shown in animal models to do so. Although limited to G-type nerve agents, these variants (e.g. IIG1, IID11, I-F11) may serve as drug candidates for further development as medical countermeasures. 3.2. Phosphotriesterase Phosphotriesterase (PTE), also termed organophosphate hydrolase (OPH), (EC 3.1.8.1) is a 336 amino-acid (36 kDa), zinc-dependent hydrolase, structured as an (αβ)8 TIM-barrel [144]. It is a bacterial enzyme that belongs to the amidohydrolase superfamily and was first identified in soil bacteria that hydrolyzed the pesticide parathion [145]. Since man-made organophosphate pesticides were introduced into the environment only in the 1950's, and since its catalytic efficiency of hydrolysis approaches diffusion limit (e.g. kcat/KM ∼109 M−1 min−1 with paraoxon), it was suggested that PTE is the product of recent and rapid natural evolution of a lactonase [146,147]. PTE is encoded on a natural plasmid (pCMS1) in a bacterial strain that was originally classified as Pseudomonas diminuta strain MG [145,148] and later reclassified as Brevundimonas diminuta [149]. The fact that the gene 55

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also be used for efficacious post-exposure treatments, when OPNA concentration buildup in the circulation is delayed by slow absorption processes (e.g. skin absorption of V-agents or prolonged exposure by inhalation to low OPNA concentrations). In conclusion, although PTE is the most efficient, natural OP hydrolase found to date, its hydrolytic activity with the toxic isomers of OPNAs had to be increased for it to serve as an effective catalytic bioscavenger. Following directed evolution and computational design efforts, more efficient PTE variants had been evolved, and were successfully used to promote survival and recovery even under conditions in which intoxication has preceded their administration (i.e. as a postexposure treatment). The broad-spectrum activity of some evolved PTE variants (e.g. 10-1-D11, Table 2) suggests that effective prophylactic protection from all currently known G- and V-type nerve agents, may be achieved using a combination of 2–3 PTE variants (e.g. 10-1D11 + d1IVA1 +YT). The exploration of PTE mutants with improved catalytic efficiencies has revealed that different mutational trajectories could be used to obtain the same goal in PTE (discussed in Ref. [38]). Thus, PTE may also serve as a good starting point for the development of active variants against other, less known OPNAs (e.g. GP [181]) or ones that may become a threat in the future.

doses of purified PTE was effective in those two experiments since the catalytic efficiencies of the enzyme with the OPs used (i.e. paraoxon, DEFP and tabun) are high at physiological temperature (i.e. ≥ 5 × 107 M−1 min−1, 38 °C) and the combination of circulating enzyme levels and high catalytic efficiency resulted in rapid detoxification [28]. However, in-vivo protection of mice from soman required much higher PTE doses (i.e. 100 mg/kg, i. v.) [161], probably due to the insufficient catalytic efficiency of PTE with soman [156]. Investigations of the substrate specificity of PTE revealed that despite its high catalytic efficiency with a diverse group of OPs [162–165], OPNAs comprise a more challenging group of substrates for PTE [166]. This is due to the fact that nerve agents, which are all methylphosphonates (except tabun), are different from pesticides that are phosphotriesters. In addition, they have a chiral phosphorous atom with an Sp isomer that is more toxic than the Rp isomer. Finally, the detoxification rates of SpOPNAs using PTE, apart from GA and GB, are several orders of magnitude lower than with paraoxon (Table 2). Thus, as with previously described enzymes, in order to obtain an efficient, broad-spectrum catalytic bioscavenger based on PTE, directed evolution and computational design methods had to be applied to improve its catalytic efficiencies with OPNAs. The task of evolving PTE variants with improved OPNA hydrolyzing capabilities was undertaken by a number of different labs (e.g. Refs. [36,37,167–171]). Since the original strains of soil bacteria, from which PTE was isolated are difficult to grow and to genetically manipulate in lab, the opd gene expressing PTE was cloned into expression plasmids and transformed into E. coli cells. In some cases, directed evolution was used to improve the expression levels of PTE in E. coli, as an initial step [172,173]. In all cases, the original 29 amino acid signal peptide sequence of opd was removed during cloning. Different approaches have been applied to create libraries of PTE mutants. For example: random mutagenesis [174], DNA shuffling [171], site-specific mutagenesis [37,169], rationally designed mutations [32,168,170], site saturation mutagenesis [36,167], computational design [37,38] and different combinations of these methods. In most cases, library design relied on the crystal structures of PTE (e.g. Ref. [175]) and on an analysis of its catalytic machinery (e.g. Ref. [176]). The resulting variant libraries were screened for improved OP hydrolase activities using either: pesticides, chromogenic analogs of OPNAs or the real nerve agents. Activities were detected in different manners: release of chromogenic compounds following the hydrolysis of OPNA analogs [32,171,174,177], detection of released fluoride ions by a fluoridespecific electrode [168], detection of released thiols using Ellman's reagent for V-agent hydrolysis [35,169,170], or AChE inhibition [37,38,178]. While these efforts yielded PTE variants with different degrees of catalytic improvement, some resulted in variants that can hydrolyze the toxic isomers of G- and V-type nerve agents with catalytic efficiencies that are sufficient for effective and sign-free prophylactic protection (Table 2) [36–38,178]. The utility of two evolved PTE variants (i.e. C23 and C23AL) as a post-exposure treatment following VX intoxication, was examined in guinea pigs (Table 1) [179,180]. In these experiments, a 2 × LD50 s c. dose of VX administered to anesthetized animals (medetomidine, midazolam and fentanyl) was followed by an i. v. or i. o. injection of purified PTE variants 5 or 15 min following intoxication. The animals were monitored for 3 h for clinical signs, mortality, brain and blood ChE activity, and blood VX concentrations. The two variants were used separately and differed in catalytic efficiency of VX hydrolysis (C23- kcat/ KM = 5 × 106 M−1min−1; C23AL-kcat/KM = 1.2 × 107 M−1min−1) and in molecular weight (C23 = 79.2 kDa, C23AL = 36.9 kDa). The protein doses used for post-exposure treatment were 2 and 5 mg/kg and no additional treatment was administered post-intoxication. In both experiments, post-exposure treatment with a catalytic bioscavenger was able to prevent mortality and systemic toxicity by VX, and to partially preserve brain AChE activity while rapidly decreasing the concentration of VX in the blood [179,180]. Thus, catalytic bioscavengers may

3.3. PTE homologs A closely related homolog of PTE named OpdA (accession number EU002557), which has a 90% amino-acid sequence identity to PTE, was isolated from a soil bacterial strain called Agrobacterium radiobacter P230 in Australia [182]. They mainly differ in the C-terminal part of the protein, where OpdA is longer than PTE by 20 amino-acids. Unlike PTE, OpdA can hydrolyze the pesticides fenthion and phosmet and displays higher turnover rates with OPs that have shorter side chains [182]. Similar to PTE the OpdA was found to hydrolyze the toxic isomers of soman and cyclosarin with moderate efficiencies (i.e. kcat/KM = 3.5 and 2.3 × 104 M−1 min−1, respectively), but was less efficient with the toxic isomers of VX (kcat/KM = 2.7 × 102 M−1 min−1), RVX (kcat/ KM = 2 × 102 M−1 min−1) or CVX (kcat/KM = 2.3 × 102 M−1 min−1) [183]. The ability of OpdA to increase survival and recovery from lethal doses of pesticides (i.e. a 3 × LD50 dose of dichlorovos, ethyl parathion [184] or methyl parathion [185]) was examined in a rat animal model. OpdA hydrolyzes these pesticides with high catalytic efficiencies (i.e. ≥ 5 × 107 M−1min−1) [184]. A single dose of purified OpdA (0.15 mg/kg) given i. v. immediately after dichlorovos poisoning (3 × LD50 dose; p. o.), resulted in 100% survival at 24 h following intoxication [184]. In the case of an ethyl parathion poisoning (i.e. the commercial parathion), a single OpdA dose (0.15 mg/kg) administered i. v. immediately or 10 min after intoxication (p.o.) did not affect survival. However, repeated dosing 45 and 90 min post intoxication resulted in 62.5% survival at 24 h [184]. In the case of methyl parathion, a similar single dose of OpdA (0.15 mg/kg) given i. v. 10 min post intoxication increased the 4 h survival rates to 100% but not at 24 h [185]. These results highlighted the ability of OpdA to act as effective post-intoxication treatment, providing there is a timely correlation between its pharmacokinetic properties (i.e. rate of clearance) and the buildup of toxic OP concentrations in-vivo. A similar experiment in nonhuman primates resulted in complete animal survival at 4 h, the duration of the experiment, using as low as 1.2 mg/kg dose of OpdA injected i. v. immediately after a 75 mg/kg (p.o.) intoxication with dichlorvos [186]. A number of homologs of PTE termed, PTE-like lactonases (PLLs) [147], have been isolated from extremophile bacteria such as: Sulfolobus solfataricus (SsoPox) [187], Sulfolobus acidocaldarius (SacPox) [188], Sulfolobus islandicus (SisLac) [189], Vulcanisaeta moutnovskia (VmutPLL) [190], Deinococcus radiodurans (Dr0930) [191] and Geobacillus stearothermophilus (GsP) [192]. Since all of them exhibit different degrees of OP hydrolysis (e.g. Refs. [189,193–195]), and since they all 56

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catalytic efficiency of the described DFPase mutant with the toxic isomers of sarin and cyclosarin (kcat/KM > 1 × 107 M−1min−1) qualify it as a potential candidate for prophylactic protection from intoxication by these nerve agents. At the same time, this work also highlights the challenges associated with attempts to improve the pharmacokinetic and immunogenic properties of non-human therapeutic protein-drugs.

exhibit enhanced stabilities relative to proteins from mesophilic bacteria (e.g. PTE, PON1), they were proposed to serve as candidates for the development of catalytic scavengers for bioremediation and surface decontamination purposes [196]. However, as with catalytic bioscavengers developed for medical countermeasures, the catalytic efficiency of these enzymes with OPNAs needs to be improved in order to make their usage applicable and their production economically feasible. Using directed evolution and selection for paraoxon hydrolysis, the catalytic efficiencies of SsoPox were improved by 2210-, 163-, 58- and 16- fold with methyl-parathion, malathion, ethyl-paraoxon and methylparaoxon respectively, in a single variant [197]. The activity of this variant with OPNAs is expected to be improved as previous work has shown that the turnover rate of Ssopox was improved 4-fold with the toxic isomer of cyclosarin following selection for improved paraoxon hydrolysis [198]. Thus, while the catalytic efficiency goal for effective surface decontamination and bioremediation using a catalytic bioscavenger has not yet been determined, it seems there are a number of promising candidates for the development of such bioscavengers.

3.5. SMP30 Human senescence marker protein-30 (HuSMP-30; EC 3.1.1.17) is a 299 amino-acid (33 kDa) protein, structured as a 6-bladed β-propeller. It has a single metal-ion binding site that can bind different di-valent ions (e.g. Ca2+, Zn2+, Mg2+, Mn2+) [209]. It is primarily expressed in liver and kidney tissues and was suggested to have antiapoptotic and antioxidant roles, as well as roles in Ca2+ regulation, and in the occurrence and development of cataracts [210,211]. It is also highly conserved (70–90% identity) in vertebrates, suggesting it is important physiologically [210]. SMP30 was first characterized in rat liver extract where it was shown to hydrolyze sarin > soman > tabun > DFP and was reported to hydrolyze all 4 isomers of soman at the same rate (∼2 × 104 M−1min−1) [212]. The DFPase activity of SMP30 is dependent on Mg2+, Mn2+, or Co2+, but it was reported to be unable to hydrolyze DFP with Ca2+ [213]. The efficiency of OPNA hydrolysis was examined using mouse SMP30 (MoSMP30), which has an amino acid sequence that is identical to HuSMP30 in 89% [214]. At G-agent concentrations of 10 μM, equivalent to ∼3 × LD50 in humans, MoSMP30 was unable to hydrolyze any of the nerve agents, probably due to high KM values; only at a much higher concentration (1 mM) MoSMP30 hydrolyzed them with the following preference: sarin > cyclosarin > soman and was unable to hydrolyze tabun [214]. Finally, a recombinant human SMP30(HuSMP30) was expressed using chaperones as a soluble protein in E. coli [215]. This may enable future directed evolution and computational engineering efforts to develop an efficient catalytic bioscavenger using SMP30.

3.4. DFPase The enzyme DFPase (EC 3.1.8.2) is a 314 amino acid (35 kDa), calcium-dependent hydrolase, structured as a six-bladed β-propeller [199]. It was initially isolated from the giant axon of the North Atlantic squid Loligo pealei [200,201], due to its ability to hydrolyze the sarin analog diisopropylfluorophophate (DFP). The enzyme, later isolated from the axon of the European squid Loligo vulgaris, was found to be able to hydrolyze also tabun, soman, sarin and cyclosarin [201,202]. In spite of its preference for the less toxic Rp isomers of nerve agents, the catalytic efficiencies of wt DFPase with the toxic isomers of sarin, and cyclosarin are quite high: kcat/KM = 2.5 and 1 × 106 M−1min−1, respectively (Table 2) [203]. In addition, DFPase can be expressed in large amounts as a soluble and active protein in E. coli [204]. Using site-directed mutagenesis, a mutation (S271A) was discovered to increase the hydrolysis of DFP by 30% relative to wt DFPase [199]. Using x-ray and neutron scattering, crystal structures of DFPase [205,206] combined with rational design, a quadruple mutant of DFPase (Mut1; Table 2) was made with increased hydrolytic rates of: DFP (2.5-fold), with the toxic isomer of sarin (5.5-fold) and with that of cyclosarin (29-fold) [203]. The mutant also exhibited reversed stereospecificity relative to wt DFPase, preferring the Sp isomers over the Rp isomers of sarin and cyclosarin by 4 and 5-fold, respectively [203]. Since DFPase is a non-mammalian protein, its clearance rate from the circulation of mammals is expected to be high and to restrict its ability to provide prophylactic protection from OPNAs. One way to increase the half-life time of proteins in the circulation is to modify them with inert polymers such as polyethylene glycol (PEG), which can reduce fast renal clearance by increasing the molecular weight of the protein, reduce immunogenicity, protect from proteolysis and lower cytotoxicity [207]. Thus, wild-type and the rationally designed DFPase mutant were conjugated to PEG, injected (i.v.) 5 min prior to intoxication and tested for their ability to protect rats from a 3 × LD50 (s.c.) dose of soman [208]. While untreated control animals died within 15 min of intoxication, rats injected with 71 mg/kg PEGylated wt DFPase and rats injected with 35.8 mg/kg PEGylated mutant DFPase all survived > 24 h post intoxication (Table 1) [208]. Rats pretreated with the PEGylated mutant DFPase showed less intoxication symptoms and less body weight loss compared to ones pretreated with the PEGylated wt DFPase. PEGylation enabled a second dosing of the protein although it was unable to inhibit the formation of anti-DFPase antibodies [208]. Thus, this study addressed not only the ability of an improved DFPase variant to protect animals from toxic doses of OPNAs, but also examined its immunogenic properties by binding it to a non-immunogenic polymer (i.e. PEG). It showed that DFPase could be conjugated to several PEG chains without causing a significant loss of catalytic efficiency, however this did not prevent the formation of anti-DFPase antibodies. The high

3.6. Organophosphate acid anhydrolase (OPAA) Organophosphate acid anhydrolase (OPAA; EC 3.1.8.2) is a 517 amino-acid (59 kDa), bimetalohydrolase, containing a small N-terminal domain and a large C-terminal domain, structured as a “pita bread” fold [216]. It also belongs to the family of prolidases that cleave imidodipeptides and imidotripeptides with a C-terminal proline or hydroxyproline (EC 3.4.13.9) [217]. It was initially isolated from a moderately-halophilic bacterium (Alteromonas Sp. strain JD6.5) found near Salt Lake City, Utah, USA, and exhibited high levels of enzymatic activity against several OPs [218]. OPAA binds two Mn2+ ions in its Cterminal domain and can catalyse the hydrolysis of DFP (kcat = 1.1 × 105 min−1), soman (kcat/KM = 1.6 × 107 M−1 min−1), sarin (kcat/KM = 1.3 × 106 M−1 min−1), tabun (kcat = 5.1 × 103 min−1), cyclosarin (kcat = 9.9 × 104 min−1), GP (2,2-dimethylcyclopentyl methylphosphonofluoridate; kcat/KM = 1.3 × 107 M−1 min−1) and paraoxon (kcat = 7.4 × 103 min−1), favoring P-F bond hydrolysis over P-CN, P-O or P-S bonds [181,216,217,219]. It was initially reported to favor the more toxic Sp isomers of OPs [220], however was recently shown it might prefer the less toxic Rp isomers [181]. OPAA was encapsulated in sterically stabilized liposomes (SL) and used to protect mice from DFP intoxication [221]. Free and SL encapsulated OPAA were administered, together with a combination of atropine and 2-PAM to mice, 1 h prior to intoxication with DFP (s.c.). The LD50 doses of DFP in mice were determined for atropine, 2-PAM, atropine plus 2-PAM, and their combination with free or liposome-encapsulated OPAA. While the free enzyme provided only a slight increase in LD50 value (33.2 vs. 29.3 mg/kg DFP), with the encapsulated OPAA the LD50 value increased 3.3-fold (i.e. 98.6 vs. 29.3 mg/kg DFP). Untreated control animals displayed an LD50 of 4.2 mg/kg DFP. In addition, the encapsulated enzyme was stable for > 2 days in the 57

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[229,230]. When recombinant human liver prolidase was purified from adenovirus infected mice livers, it was shown to hydrolyze the toxic isomers of sarin, soman, and cyclosarin with moderate catalytic efficiencies (kcat/KM) of ∼104 M−1 min−1, but was unable to hydrolyze tabun or VX (Table 2) [230]. Thus, conflicting reports remain about the ability of Prolidase from human liver to hydrolyze tabun and VX: While some have observed hydrolysis of tabun [29,226,227] others have not [230], and while some have observed hydrolysis of VX [227], others have not [226,230]. The ability of recombinant HuProlidase to protect mice from intoxication by DFP [229] or by OPNAs [230] was tested using the adenovirus expression vector system in mice. Although the gene was produced in high levels in the circulation of mice and increased the levels of prolidase by 100- to 150-fold, it delayed the death of mice from two, cumulative 1 × LD50 doses of DFP by only 4–8 h [229]. Similarly, the protein was unable to protect mice from two, cumulative 1 × LD50 doses of sarin, soman or cyclosarin (Table 1) [230]. Mice that expressed mouse BChE following an adenovirus-vector infection, were able to survive for 24 h past similar challenges with sarin, soman and cyclosarin. Since the expressed levels of BChE on the 5th day were 500–1500-fold higher than control animals and those of HuProlidase were only 30-fold higher then controls in that experiment, it was concluded that the blood concentration of reHuProlidase was not high enough to provide sufficient protection from sarin, soman or cyclosarin [230]. It is also likely that the low catalytic efficiency of prolidase compared with the high biomolecular rate of BChE binding to OPs, resulted in the observed differences in survival. In summary, wt HuProlidase is insufficiently active to provide protection from intoxication by OPNAs. It could serve, however, as a starting point for computational design and directed evolution efforts that would change and increase its specificity towards the toxic isomers of OPNAs.

circulation. Thus, OPAA was shown to have considerable antidotal activity against DFP intoxication when encapsulated into sterically stabilized liposomes in contrast to the free enzyme under the same conditions [221]. Wild type OPAA has very low hydrolytic activity with RVX (kcat/ KM = 5.5 × 102 M−1 min−1) [222]. In an attempt to increase its efficiency of RVX hydrolysis, the crystal structure of OPAA from (Alteromonas Sp. strain JD6.5) [216] was used for rational design of site specific mutations. Designed mutants were cloned, expressed and tested for RVX hydrolysis [222]. Following several rounds of mutagenesis and screening, including the crystallization and structural elucidation of an improved variant, a triple-mutant of OPAA that was improved ∼36fold in catalytic efficiency with the less-toxic Rp isomer of RVX (kcat/ KM = 2 × 104 M−1 min−1), was identified (FLYD; Table 2). In order to increase its activity with the more toxic, Sp isomer, of RVX, an additional mutation was rationally designed into the active site. The catalytic activity of the resulting quadruple-mutant towards the Sp isomer of RVX (kcat/KM = 8.9 × 103 M−1 min−1) was only 16-fold improved relative to wt OPAA, but the variant was now able to hydrolyze both isomers of RVX [222]. When the double mutants, created in that study, were examined for their ability to hydrolyze G-type nerve agents, it was found that the Y212F/V342L (FL) mutant was improved 6-fold over wt OPAA at hydrolyzing sarin (kcat/KM = 8.5 × 106 M−1 min−1), 5-fold relative to wt OPAA with soman (kcat/KM = 7.5 × 107 M−1 min−1) and 10-fold with GP (kcat/KM = 1.3 × 108 M−1 min−1) [181]. However, it was also found that in the case of sarin, this improvement was due to an increase with the less toxic Rp isomer. The activity of OPAA-FL with the more toxic isomers of soman and GP was not defined [181]. In conclusion, OPAA can exhibit very high catalytic efficiencies when hydrolyzing OPNAs (e.g. soman, GP), however, it may also have a strong preference for their less toxic Rp isomers. Variants evolved so far have mostly been able to increase the activity with these isomers. Thus, future attempts to improve the catalytic efficiency of OPAA will require screening or selection methods aiming to identify variants with preference for binding and hydrolyzing the Sp isomers of nerve agents.

3.8. Methylparathion hydrolase Methylparathion hydrolase (MPH; EC 3.1.8.1) is a 331 amino-acid (35 kDa), zinc-dependent aryldialkylphosphatase that is structured as a metallo-β-lactamase. It was initially identified in a rod-shaped gramnegative bacterium called Plesiomonas sp. Strain M6, and its gene was named mpd (methyl parathion degrading) [231]. An almost identical gene (99.5% DNA sequence identity) was identified on a plasmid pZWL0 from Pseudomonase sp. Strain WBC-3 and shown to hydrolyze the OPs: methyl parathion, parathion and fenitrothion [232]. The gene, named mph, enables this bacteria to utilize methyl parathion as its sole source of carbon, nitrogen and energy and was isolated from 7 additional bacterial strains [233]. The protein can be expressed and purified from E. coli [232], and its structure was found to contain a binuclear center of zinc and cadmium ions in a monomer described as an αβ/βα sandwich [234] although the wild type protein is likely to contain only zinc ions. The catalytic efficiency of MPH with methyl parathion is high (kcat/KM = 6 × 107 M−1 min−1) [234] however, its activity with other OPs such as ethyl paraoxon was found to be 1 to 3-orders of magnitude lower [235]. Using directed evolution, the activity of an MPH variant was increased by 4-fold [236] with chlorpyrifos and by 100-fold with ethyl paraoxon (kcat/KM = 3.5 × 107 M−1 min−1) [235]. MPH could be targeted for surface display, as shown with Pseudomonas putida strain JS444, to facilitate decontamination [237] and stabilized using rational design [238]. Recently, the catalytic mechanism of MPH has been investigated using a computational approach [239]. In order to assess the potential of MPH as a catalytic scavenger, its activity with OPNAs in general, and with their Sp isomers, in particular, needs to be examined.

3.7. Prolidase Human prolidase (HuProlidase; EC 3.4.13.9) is a 493 amino acid (54 kDa), bi-metallic, Mn2+ dependent dipeptidase that catalyzes the hydrolysis of specific C-terminal dipeptides [223]. It is a homodimer with two domains: N-terminal domain (184 residues) and a C-terminal domain (309 residues) that is structured as a “pita bread” fold. It plays an important role at the final stage of protein catabolism and is responsible for the hydrolysis of dipeptides containing a proline or a hydroxyproline residue at their C-terminals and especially the Gly-Pro bond [224]. Although it shares only 30% amino-acid sequence identity with OPAA from Alteromonas Sp. strain JD6.5, HuProlidase is related to it, and to other bacterial OPAAs (e.g. OPAA from Alteromonas macleodii [225]), in function and structure. Human prolidase is ubiquitously expressed and is mainly implicated in the degradation of dietary and endogenous proteins such as collagen. It also plays a role in the regulation of peptidic hormones. Its deficiency results in skin ulcerations, recurrent infections of the respiratory tract and mental retardation of different degrees [224]. Initially, human liver prolidase purified from human liver or as a recombinant protein from E. coli cells, was found to hydrolyze soman, sarin, cyclosarin, tabun and DFP but not VX or paraoxon [29,226]. In a later work, it was shown to hydrolyze DFP and the toxic isomers of all G-type agents and VX [227]. In contrast, recombinant human skin and kidney prolidases, purified from Trichoplusia ni larva, were found to hydrolyze DFP and sarin but were inefficient with other OPNAs (e.g. soman, tabun, VX); presumably due to sequence differences between them and liver prolidase [227]. Human liver prolidase could be expressed and purified from COS-7 or yeast cells [228] and from mice liver, following its infection with an adenovirus expression vector

4. De-novo designed catalytic bioscavengers Advancements in the computational design of proteins and specifically, enzymes (reviewed in Refs. [125,127,240,241]), have enabled 58

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In spite of the complicating factors described above, the past 40 years has seen an increasing application of protein drugs such as: antibodies, fusion proteins and enzymes in the treatment of a wide range of human diseases. For example, between 2011 and 2016, sixty-two new therapeutic proteins had been approved for use by the U.S. Food and Drug Administration (FDA) [250]. The facilitated application of protein drugs has been enabled by the development of novel strategies for the reduction of immunogenicity, increasing stability, improving delivery and for the assessment of product safety. These include: chemical modifications of therapeutic proteins by polymers such as PEG to mask immunogenic epitopes and reduce clearance rates [251,252], delivery of protein drugs inside vesicles [253], improvement of cellular penetration by conjugation to cell-penetrating peptides [254], protein engineering [255] and the addition of excipients to protect protein structure during production and storage [256]. Thus, the challenges of developing catalytic bioscavengers as medical countermeasures may be successfully met using the strategies that have enabled the clinical application of other enzymes.

the creation of man-made enzymes with desired properties. Such enzymes have been shown to catalyse chemical reactions, for which there is no known natural enzyme catalyst, or that utilize unnatural substrates. Since OPNAs constitute xenobiotic compounds, not known to have a natural hydrolase that has evolved to detoxify them, they are prime targets for such design efforts. In an attempt to create an efficient OP hydrolyzing enzyme, the active site of a zinc-containing, mouse adenosine deaminase was computationally redesigned to enable OP hydrolysis by stabilizing the modeled transition state of an organophosphate [242]. The resulting protein, PT3, was able to hydrolyze a model OP compound, diethyl 7-hydroxycoumarinyl phosphate (DECP), with catalytic efficiency (kcat/KM) of 2.4 × 102 [M−1 min−1], while the w. t scaffold protein was 3 orders of magnitude less active [242]. Following only two rounds of directed evolution, a variant with ∼2500fold higher catalytic activity was obtained (kcat/KM = 5.9 × 105 [M−1 min−1]). Thus, the combination of computational design and directed evolution yielded an improvement of more than 6 orders of magnitude in catalytic efficiency with DECP. De-novo computational design methods can provide many potential benefits for creating catalytic bioscavengers. They may enable enzyme engineers to graft the active site architecture of an efficient OPNA hydrolyase into a new scaffold with desired properties such as: reduced immunogenicity, improved stability, smaller size, and longer circulatory life-time. In this manner, computational design may also help circumvent some of the potential problems that may arise during attempts to employ catalytic bioscavengers, evolved from non-human proteins, as prophylactic drugs.

6. Summary The importance of research and development of catalytic bioscavengers as medical countermeasures stems from the following: First, OPNA intoxications are difficult to treat, and protein bioscavengers are currently the only molecules shown to provide effective prophylactic protection from lethal OPNA intoxications in-vivo, as a stand-alone treatment. Second, despite almost 70 years of research, attempts to design improved drugs for treatment of OP intoxications have yielded only modest advances. Most countries still maintain atropine, 2-PAM and valium as their standard therapeutic regimen for such conditions and the need for a broad-spectrum, highly efficacious and fully bioavailable ChE reactivator, has not been met yet. Stoichiometric bioscavengers may enable both effective prophylaxis and post-intoxication treatment. However, the need to repetitively introduce large doses of protein drugs in order to counteract an unknown amount of OP, will continue to burden the application of stoichiometric bioscavengers even after problems of production and cost are overcome. In contrast, catalytic bioscavengers may provide effective prophylactic protection and improve post-exposure treatment of OPNA intoxications using low protein doses. Unlike current treatments, they may promote survival while preventing the onset of intoxication symptoms and their manifestations. In the past decade, considerable progress has been made in the research of catalytic bioscavengers; transitioning from attempts to identify new enzymes with nerve agent hydrolyzing capabilities to improving previously identified enzymes and testing them in-vivo. On one hand, there has been a growing realization that natural enzymes are not likely to exhibit high catalytic efficiencies with OPNAs, which are manmade, xenobiotic compounds; on the other hand, progress in directed evolution and computational design methods have facilitated the ability to redesign natural enzymes and to obtain specific and highly active OPNA-hydrolyzing variants. As a result, a number of enzymes, PON1, PTE and DFPase, have been evolved to hydrolyze the toxic isomers of Gand V-type nerve agents with high catalytic efficiencies (kcat/ KM) > 1 × 107 M−1 min−1. This has also enabled to test the validity of a theoretical model [26] that predicted the dose of a bioscavenger required to prevent intoxication from a specific lethal dose of OPNA, by knowing its catalytic efficiency for the toxic isomers of the intoxicating agent. The accumulated data from animal protection experiments that employed different enzymes, OPs and animals [28], has demonstrated the generality of the catalytic bioscavenger model. It suggests that the protective dose required to provide prophylactic protection from lethal doses of OPNAs, as a stand-alone medical countermeasure in humans, could be reliably estimated. In addition, animal experiments indicate that catalytic bioscavengers may also be used effectively for post-exposure treatments, in cases where the increase in circulatory OPNA

5. Catalytic bioscavengers – from bench to bedside The applicability of any newly developed drug depends primarily on the ability to demonstrate its safety and efficacy in humans. Protein based drugs require special attention due to their high molecular weight, susceptibility to proteolytic breakdown, rapid plasma clearance, immunogenicity and tendency for denaturation and aggregation [243,244]. These properties may complicate drug production, formulation, storage, delivery and efficacy in addition to increasing manufacturing costs. For example, the oral delivery of proteins is complicated by the need to protect them from proteolytic enzymes and low pH in the gastro-intestinal tract while transdermal delivery is restricted by their low skin penetration properties [245]. The tendency of proteins to denature and aggregate may require storage of therapeutic proteins at low temperatures (e.g. 4 Deg C) right up to use and may significantly limit their shelf-life relative to those of small-molecule drugs [246]. Immunogenicity is a particular concern when dealing with protein drugs, especially in cases of repeated administrations. Many factors influence the immunogenicity of a therapeutic protein, primarily, its similarity to endogenous proteins. If it is completely “foreign” (e.g. a bacterial, plant or yeast protein inside a human body), it is likely to elicit an adaptive immune response upon its introduction. If the protein is highly identical or compatible with an endogenous protein, than it will induce an antibody response only if it can break B-cell tolerance [247]. This may occur by: protein aggregation, differences in glycosylation patterns and the presence of impurities and contaminants from the manufacturing process. In addition, other factors such as the route of administration (e.g. intravenous or intramuscular), dose levels, the duration of treatment and the genetic background of the patient can influence its immunogenicity [244,248]. Enzymes from non-human sources are likely to be recognized by the immune system as foreign antigens and elicit an immune response that may lead to their inactivation by neutralizing antibodies. This problem is likely to increase with repeated dosing and can render the drug ineffective and perhaps harmful. On the other hand, the administration of large doses of human enzymes may elicit an autoimmune response that may lead to the inactivation of both the exogenously introduced and endogenously generated enzyme [249] [10]. 59

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concentration is slow due to absorption processes (e.g. VX skin penetration or prolonged exposure to low-concentrations of toxic vapors). However, there are many issues relating to the safety, efficacy, production and medical use of catalytic bioscavengers that are yet unresolved. Research focusing on: immunogenicity, pharmacokinetic and pharmacodynamics properties (i.e. absorption, distribution, metabolism and excretion), cross-reactivity, stability, production, storage and delivery is required in order to transition catalytic bioscavengers from promising proto-types to bona-fide drug candidates.

organophosphorus compounds, Pharmacol. Ther. 139 (2013) 249–259. [23] M. Katalinic, N. Macek Hrvat, K. Baumann, S. Morasi Pipercic, S. Makaric, S. Tomic, O. Jovic, T. Hrenar, A. Milicevic, D. Jelic, S. Zunec, I. Primozic, Z. Kovarik, A comprehensive evaluation of novel oximes in creation of butyrylcholinesterase-based nerve agent bioscavengers, Toxicol. Appl. Pharmacol. 310 (2016) 195–204. [24] A. Shafferman, A. Ordentlich, D. Barak, D. Stein, N. Ariel, B. Velan, Aging of phosphylated human acetylcholinesterase: catalytic processes mediated by aromatic and polar residues of the active centre, Biochem. J. 318 (Pt 3) (1996) 833–840. [25] F. Worek, G. Reiter, P. Eyer, L. Szinicz, Reactivation kinetics of acetylcholinesterase from different species inhibited by highly toxic organophosphates, Arch. Toxicol. 76 (2002) 523–529. [26] R.D. Gupta, M. Goldsmith, Y. Ashani, Y. Simo, G. Mullokandov, H. Bar, M. BenDavid, H. Leader, R. Margalit, I. Silman, J.L. Sussman, D.S. Tawfik, Directed evolution of hydrolases for prevention of G-type nerve agent intoxication, Nat. Chem. Biol. 7 (2011) 120–125. [27] D. Josse, O. Lockridge, W. Xie, C.F. Bartels, L.M. Schopfer, P. Masson, The active site of human paraoxonase (PON1), J. Appl. Toxicol. 21 (Suppl 1) (2001) S7–S11. [28] Y. Ashani, H. Leader, N. Aggarwal, I. Silman, F. Worek, J.L. Sussman, M. Goldsmith, In vitro evaluation of the catalytic activity of paraoxonases and phosphotriesterases predicts the enzyme circulatory levels required for in vivo protection against organophosphate intoxications, Chem. Biol. Interact. 259 (2016) 252–256. [29] R.C. diTargiani, L. Chandrasekaran, T. Belinskaya, A. Saxena, In search of a catalytic bioscavenger for the prophylaxis of nerve agent toxicity, Chem. Biol. Interact. 187 (2010) 349–354. [30] R. Iyer, B. Iken, A. Damania, A comparison of organophosphate degradation genes and bioremediation applications, Environ Microbiol Rep 5 (2013) 787–798. [31] G. Amitai, L. Gaidukov, R. Adani, S. Yishay, G. Yacov, M. Kushnir, S. Teitlboim, M. Lindenbaum, P. Bel, O. Khersonsky, D.S. Tawfik, H. Meshulam, Enhanced stereoselective hydrolysis of toxic organophosphates by directly evolved variants of mammalian serum paraoxonase, FEBS J. 273 (2006) 1906–1919. [32] P.C. Tsai, A. Bigley, Y. Li, E. Ghanem, C.L. Cadieux, S.A. Kasten, T.E. Reeves, D.M. Cerasoli, F.M. Raushel, Stereoselective hydrolysis of organophosphate nerve agents by the bacterial phosphotriesterase, Biochemistry 49 (2010) 7978–7987. [33] M. Goldsmith, Y. Ashani, Y. Simo, M. Ben-David, H. Leader, I. Silman, J.L. Sussman, D.S. Tawfik, Evolved stereoselective hydrolases for broad-spectrum g-type nerve agent detoxification, Chem. Biol. 19 (2012) 456–466. [34] A.N. Bigley, M.F. Mabanglo, S.P. Harvey, F.M. Raushel, Variants of phosphotriesterase for the enhanced detoxification of the chemical warfare agent VR, Biochemistry 54 (2015) 5502–5512. [35] A.N. Bigley, C. Xu, T.J. Henderson, S.P. Harvey, F.M. Raushel, Enzymatic neutralization of the chemical warfare agent VX: evolution of phosphotriesterase for phosphorothiolate hydrolysis, J. Am. Chem. Soc. 135 (2013) 10426–10432. [36] P.C. Tsai, N. Fox, A.N. Bigley, S.P. Harvey, D.P. Barondeau, F.M. Raushel, Enzymes for the homeland defense: optimizing phosphotriesterase for the hydrolysis of organophosphate nerve agents, Biochemistry 51 (2012) 6463–6475. [37] I. Cherny, P. Greisen Jr., Y. Ashani, S.D. Khare, G. Oberdorfer, H. Leader, D. Baker, D.S. Tawfik, Engineering V-type nerve agents detoxifying enzymes using computationally focused libraries, ACS Chemical Biology 8 (2013) 2394–2403. [38] M. Goldsmith, N. Aggarwal, Y. Ashani, H. Jubran, P.J. Greisen, S. Ovchinnikov, H. Leader, D. Baker, J.L. Sussman, A. Goldenzweig, S.J. Fleishman, D.S. Tawfik, Overcoming an optimization plateau in the directed evolution of highly efficient nerve agent bioscavengers, Protein Eng. Des. Sel. 30 (2017) 333–345. [39] T. Satoh, M. Hosokawa, Structure, function and regulation of carboxylesterases, Chem. Biol. Interact. 162 (2006) 195–211. [40] M.J. Hatfield, R.A. Umans, J.L. Hyatt, C.C. Edwards, M. Wierdl, L. Tsurkan, M.R. Taylor, P.M. Potter, Carboxylesterases: general detoxifying enzymes, Chem. Biol. Interact. 259 (2016) 327–331. [41] M.R. Redinbo, P.M. Potter, Mammalian carboxylesterases: from drug targets to protein therapeutics, Drug Discov. Today 10 (2005) 313–325. [42] S. Bencharit, C.L. Morton, E.L. Howard-Williams, M.K. Danks, P.M. Potter, M.R. Redinbo, Structural insights into CPT-11 activation by mammalian carboxylesterases, Nat. Struct. Biol. 9 (2002) 337–342. [43] D.M. Maxwell, The specificity of carboxylesterase protection against the toxicity of organophosphorus compounds, Toxicol. Appl. Pharmacol. 114 (1992) 306–312. [44] D.M. Maxwell, K.M. Brecht, Carboxylesterase: specificity and spontaneous reactivation of an endogenous scavenger for organophosphorus compounds, J. Appl. Toxicol. 21 (Suppl 1) (2001) S103–S107. [45] E.G. Duysen, F. Koentgen, G.R. Williams, C.M. Timperley, L.M. Schopfer, D.M. Cerasoli, O. Lockridge, Production of ES1 plasma carboxylesterase knockout mice for toxicity studies, Chem. Res. Toxicol. 24 (2011) 1891–1898. [46] A.C. Hemmert, T.C. Otto, M. Wierdl, C.C. Edwards, C.D. Fleming, M. MacDonald, J.R. Cashman, P.M. Potter, D.M. Cerasoli, M.R. Redinbo, Human carboxylesterase 1 stereoselectively binds the nerve agent cyclosarin and spontaneously hydrolyzes the nerve agent sarin, Mol. Pharmacol. 77 (2010) 508–516. [47] D.M. Maxwell, C.N. Lieske, K.M. Brecht, Oxime-induced reactivation of carboxylesterase inhibited by organophosphorus compounds, Chem. Res. Toxicol. 7 (1994) 428–433. [48] A.L. Devonshire, L.M. Field, Gene amplification and insecticide resistance, Annu. Rev. Entomol. 36 (1991) 1–23. [49] C. Claudianos, R.J. Russell, J.G. Oakeshott, The same amino acid substitution in orthologous esterases confers organophosphate resistance on the house fly and a blowfly, Insect Biochem. Mol. Biol. 29 (1999) 675–686. [50] R.D. Newcomb, P.M. Campbell, D.L. Ollis, E. Cheah, R.J. Russell, J.G. Oakeshott, A

Appendix A. Supplementary data Supplementary data related to this article can be found at https:// doi.org/10.1016/j.cbi.2018.07.006. References [1] T.M. Shih, D.E. Lenz, D.M. Maxwell, Effects of repeated injection of sublethal doses of soman on behavior and on brain acetylcholine and choline concentrations in the rat, Psychopharmacology (Berl) 101 (1990) 489–496. [2] V. Aroniadou-Anderjaska, T.H. Figueiredo, J.P. Apland, E.M. Prager, V.I. Pidoplichko, S.L. Miller, M.F. Braga, Long-term neuropathological and behavioral impairments after exposure to nerve agents, Ann. N. Y. Acad. Sci. 1374 (2016) 17–28. [3] M.A. Dunn, F.R. Sidell, Progress in medical defense against nerve agents, Jama 262 (1989) 649–652. [4] P. Taylor, Anticholinesterase agents, in: L.L. Brunton, Bruce A. Chabner, B.C. Knollmann (Eds.), Goodman & Gilman's the Pharmacological Basis of Therapeutics, McGraw-Hill, New York, 2011, p. 2084. [5] F. Worek, H. Thiermann, T. Wille, Catalytic bioscavengers in nerve agent poisoning: a promising approach? Toxicol. Lett. 244 (2016) 143–148. [6] P. Masson, S.V. Lushchekina, Emergence of catalytic bioscavengers against organophosphorus agents, Chem. Biol. Interact. 259 (2016) 319–326. [7] Y. Ashani, S. Pistinner, Estimation of the upper limit of human butyrylcholinesterase dose required for protection against organophosphates toxicity: a mathematically based toxicokinetic model, Toxicol. Sci. 77 (2004) 358–367. [8] O. Cohen, C. Kronman, L. Raveh, O. Mazor, A. Ordentlich, A. Shafferman, Comparison of polyethylene glycol-conjugated recombinant human acetylcholinesterase and serum human butyrylcholinesterase as bioscavengers of organophosphate compounds, Mol. Pharmacol. 70 (2006) 1121–1131. [9] J.M. Corbin, B.I. Hashimoto, K. Karuppanan, Z.R. Kyser, L. Wu, B.A. Roberts, A.R. Noe, R.L. Rodriguez, K.A. McDonald, S. Nandi, Semicontinuous bioreactor production of recombinant butyrylcholinesterase in transgenic rice cell suspension cultures, Front. Plant Sci. 7 (2016) 412. [10] J. Descotes, A. Gouraud, Clinical immunotoxicity of therapeutic proteins, Expet Opin. Drug Metabol. Toxicol. 4 (2008) 1537–1549. [11] W.N. Aldridge, Serum esterases. I. Two types of esterase (A and B) hydrolysing pnitrophenyl acetate, propionate and butyrate, and a method for their determination, Biochem. J. 53 (1953) 110–117. [12] J. Estevez, E. Vilanova, Model equations for the kinetics of covalent irreversible enzyme inhibition and spontaneous reactivation: esterases and organophosphorus compounds, Crit. Rev. Toxicol. 39 (2009) 427–448. [13] F. Worek, H. Thiermann, L. Szinicz, P. Eyer, Kinetic analysis of interactions between human acetylcholinesterase, structurally different organophosphorus compounds and oximes, Biochem. Pharmacol. 68 (2004) 2237–2248. [14] F. Worek, N. Aurbek, T. Wille, P. Eyer, H. Thiermann, Kinetic prerequisites of oximes as effective reactivators of organophosphate-inhibited acetylcholinesterase: a theoretical approach, J. Enzym. Inhib. Med. Chem. 26 (2011) 303–308. [15] R. Sharma, B. Gupta, N. Singh, J.R. Acharya, K. Musilek, K. Kuca, K.K. Ghosh, Development and structural modifications of cholinesterase reactivators against chemical warfare agents in last decade: a review, Mini Rev. Med. Chem. 15 (2015) 58–72. [16] O. Lockridge, Review of human butyrylcholinesterase structure, function, genetic variants, history of use in the clinic, and potential therapeutic uses, Pharmacol. Ther. 148 (2015) 34–46. [17] C. Saldanha, Human Erythrocyte acetylcholinesterase in health and disease, Molecules 22 (2017). [18] H.P. Benschop, L.P.A. Dejong, Nerve agent stereoisomers - analysis, isolation, and toxicology, Accounts Chem. Res. 21 (1988) 368–374. [19] D.E. Lorke, H. Kalasz, G.A. Petroianu, K. Tekes, Entry of oximes into the brain: a review, Curr. Med. Chem. 15 (2008) 743–753. [20] V.A. Voicu, J. Bajgar, A. Medvedovici, F.S. Radulescu, D.S. Miron, Pharmacokinetics and pharmacodynamics of some oximes and associated therapeutic consequences: a critical review, J. Appl. Toxicol. 30 (2010) 719–729. [21] Z. Radic, R.K. Sit, E. Garcia, L. Zhang, S. Berend, Z. Kovarik, G. Amitai, V.V. Fokin, K. Barry Sharpless, P. Taylor, Mechanism of interaction of novel uncharged, centrally active reactivators with OP-hAChE conjugates, Chem. Biol. Interact. 203 (2013) 67–71. [22] F. Worek, H. Thiermann, The value of novel oximes for treatment of poisoning by

60

Chemico-Biological Interactions 292 (2018) 50–64

M. Goldsmith, Y. Ashani

[51]

[52]

[53]

[54]

[55]

[56] [57] [58] [59] [60] [61] [62]

[63]

[64] [65]

[66]

[67]

[68]

[69]

[70]

[71]

[72]

[73]

[74]

[75]

[76]

[77]

[78]

755–763. [79] V. Kumar, W.B. Elliott, The acetylcholinesterase of Bungarus fasciatus venom, Eur. J. Biochem. 34 (1973) 586–592. [80] X. Cousin, S. Bon, N. Duval, J. Massoulie, C. Bon, Cloning and expression of acetylcholinesterase from Bungarus fasciatus venom. A new type of cooh-terminal domain; involvement of a positively charged residue in the peripheral site, J. Biol. Chem. 271 (1996) 15099–15108. [81] T. Poyot, F. Nachon, M.T. Froment, M. Loiodice, S. Wieseler, L.M. Schopfer, O. Lockridge, P. Masson, Mutant of Bungarus fasciatus acetylcholinesterase with low affinity and low hydrolase activity toward organophosphorus esters, Biochim. Biophys. Acta 1764 (2006) 1470–1478. [82] P. Masson, O. Lockridge, Butyrylcholinesterase for protection from organophosphorus poisons: catalytic complexities and hysteretic behavior, Arch. Biochem. Biophys. 494 (2010) 107–120. [83] E. Boudinot, L. Taysse, S. Daulon, A. Chatonnet, J. Champagnat, A.S. Foutz, Effects of acetylcholinesterase and butyrylcholinesterase inhibition on breathing in mice adapted or not to reduced acetylcholinesterase, Pharmacol. Biochem. Behav. 80 (2005) 53–61. [84] V.P. Chen, Y. Gao, L. Geng, S. Brimijoin, Butyrylcholinesterase regulates central ghrelin signaling and has an impact on food intake and glucose homeostasis, Int. J. Obes. 41 (2017) 1413–1419. [85] Y. Nicolet, O. Lockridge, P. Masson, J.C. Fontecilla-Camps, F. Nachon, Crystal structure of human butyrylcholinesterase and of its complexes with substrate and products, J. Biol. Chem. 278 (2003) 41141–41147. [86] C.B. Millard, O. Lockridge, C.A. Broomfield, Design and expression of organophosphorus acid anhydride hydrolase activity in human butyrylcholinesterase, Biochemistry 34 (1995) 15925–15933. [87] L.M. Schopfer, A.T. Boeck, C.A. Broomfield, O. Lockridge, Mutants of human butyrylcholinesterase with organophosphate hydrolase activity; Evidence that His117 is a general base catalyst for hydrolysis of echothiophate, J Med Chem Def 2 (2004) 1–21. [88] M. Harel, A. Aharoni, L. Gaidukov, B. Brumshtein, O. Khersonsky, R. Meged, H. Dvir, R.B. Ravelli, A. McCarthy, L. Toker, I. Silman, J.L. Sussman, D.S. Tawfik, Structure and evolution of the serum paraoxonase family of detoxifying and antiatherosclerotic enzymes, Nat. Struct. Mol. Biol. 11 (2004) 412–419. [89] S.L. Primo-Parmo, R.C. Sorenson, J. Teiber, B.N. La Du, The human serum paraoxonase/arylesterase gene (PON1) is one member of a multigene family, Genomics 33 (1996) 498–507. [90] P.N. Durrington, B. Mackness, M.I. Mackness, Paraoxonase and atherosclerosis, Arterioscler. Thromb. Vasc. Biol. 21 (2001) 473–480. [91] M.C. Blatter Garin, X. Moren, R.W. James, Paraoxonase-1 and serum concentrations of HDL-cholesterol and apoA-I, J. Lipid Res. 47 (2006) 515–520. [92] A. Mazur, An enzyme in animal tissues capable of hydrolysing the phosphorusfluorine bond of alkyl fluorophosphates, J. Biol. Chem. 164 (1946) 271–289. [93] W.N. Aldridge, Serum esterases. II. An enzyme hydrolysing diethyl p-nitrophenyl phosphate (E600) and its identity with the A-esterase of mammalian sera, Biochem. J. 53 (1953) 117–124. [94] D.I. Draganov, J.F. Teiber, A. Speelman, Y. Osawa, R. Sunahara, B.N. La Du, Human paraoxonases (PON1, PON2, and PON3) are lactonases with overlapping and distinct substrate specificities, J. Lipid Res. 46 (2005) 1239–1247. [95] O. Khersonsky, D.S. Tawfik, Structure-reactivity studies of serum paraoxonase PON1 suggest that its native activity is lactonase, Biochemistry 44 (2005) 6371–6382. [96] M.I. Mackness, P.N. Durrington, B. Mackness, How high-density lipoprotein protects against the effects of lipid peroxidation, Curr. Opin. Lipidol. 11 (2000) 383–388. [97] K. Kowalska, E. Socha, H. Milnerowicz, Review: the role of paraoxonase in cardiovascular diseases, Ann. Clin. Lab. Sci. 45 (2015) 226–233. [98] D.A. Chistiakov, A.A. Melnichenko, A.N. Orekhov, Y.V. Bobryshev, Paraoxonase and atherosclerosis-related cardiovascular diseases, Biochimie 132 (2017) 19–27. [99] C.J. Brealey, C.H. Walker, B.C. Baldwin, A-esterase activities in relation to the differential toxicity of pirimiphos-methyl to birds and mammals, Pestic. Sci. 11 (1980) 546–554. [100] L.G. Costa, C.L. Galli, S.D. Murphy, North Atlantic Treaty Organization. Scientific Affairs Division., Toxicology of Pesticides: Experimental, Clinical, and Regulatory Perspectives, Springer-Verlag, Berlin; New York, 1987. [101] W.F. Li, L.G. Costa, C.E. Furlong, Serum paraoxonase status: a major factor in determining resistance to organophosphates, J. Toxicol. Environ. Health 40 (1993) 337–346. [102] W.F. Li, C.E. Furlong, L.G. Costa, Paraoxonase protects against chlorpyrifos toxicity in mice, Toxicol. Lett. 76 (1995) 219–226. [103] D.M. Shih, L. Gu, Y.R. Xia, M. Navab, W.F. Li, S. Hama, L.W. Castellani, C.E. Furlong, L.G. Costa, A.M. Fogelman, A.J. Lusis, Mice lacking serum paraoxonase are susceptible to organophosphate toxicity and atherosclerosis, Nature 394 (1998) 284–287. [104] W.F. Li, L.G. Costa, R.J. Richter, T. Hagen, D.M. Shih, A. Tward, A.J. Lusis, C.E. Furlong, Catalytic efficiency determines the in-vivo efficacy of PON1 for detoxifying organophosphorus compounds, Pharmacogenetics 10 (2000) 767–779. [105] L.G. Costa, G. Giordano, T.B. Cole, J. Marsillach, C.E. Furlong, Paraoxonase 1 (PON1) as a genetic determinant of susceptibility to organophosphate toxicity, Toxicology 307 (2013) 115–122. [106] F.M. Buratti, M.T. Volpe, A. Meneguz, L. Vittozzi, E. Testai, CYP-specific bioactivation of four organophosphorothioate pesticides by human liver microsomes, Toxicol. Appl. Pharmacol. 186 (2003) 143–154. [107] G. Kaur, A.K. Jain, S. Singh, CYP/PON genetic variations as determinant of

single amino acid substitution converts a carboxylesterase to an organophosphorus hydrolase and confers insecticide resistance on a blowfly, Proc. Natl. Acad. Sci. U. S. A. 94 (1997) 7464–7468. P.D. Mabbitt, G.J. Correy, T. Meirelles, N.J. Fraser, M.L. Coote, C.J. Jackson, Conformational disorganization within the active site of a recently evolved organophosphate hydrolase limits its catalytic efficiency, Biochemistry 55 (2016) 1408–1417. A.C. Hemmert, T.C. Otto, R.A. Chica, M. Wierdl, J.S. Edwards, S.M. Lewis, C.C. Edwards, L. Tsurkan, C.L. Cadieux, S.A. Kasten, J.R. Cashman, S.L. Mayo, P.M. Potter, D.M. Cerasoli, M.R. Redinbo, Nerve agent hydrolysis activity designed into a human drug metabolism enzyme, PLoS One 6 (2011) e17441. P.M. Legler, S.M. Boisvert, J.R. Compton, C.B. Millard, Development of organophosphate hydrolase activity in a bacterial homolog of human cholinesterase, Front Chem 2 (2014) 46. P. Masson, P.L. Fortier, C. Albaret, M.T. Froment, C.F. Bartels, O. Lockridge, Aging of di-isopropyl-phosphorylated human butyrylcholinesterase, Biochem. J. 327 (Pt 2) (1997) 601–607. C.B. Millard, O. Lockridge, C.A. Broomfield, Organophosphorus acid anhydride hydrolase activity in human butyrylcholinesterase: synergy results in a somanase, Biochemistry 37 (1998) 237–247. I. Silman, J.L. Sussman, Acetylcholinesterase: how is structure related to function? Chem. Biol. Interact. 175 (2008) 3–10. I. Silman, J.L. Sussman, Recent developments in structural studies on acetylcholinesterase, J. Neurochem. 142 (Suppl 2) (2017) 19–25. R.L. Rotundo, Biogenesis, assembly and trafficking of acetylcholinesterase, J. Neurochem. 142 (Suppl 2) (2017) 52–58. M. Bazelyansky, E. Robey, J.F. Kirsch, Fractional diffusion-limited component of reactions catalyzed by acetylcholinesterase, Biochemistry 25 (1986) 125–130. C. Legay, Why so many forms of acetylcholinesterase? Microsc. Res. Tech. 49 (2000) 56–72. H. Dvir, I. Silman, M. Harel, T.L. Rosenberry, J.L. Sussman, Acetylcholinesterase: from 3D structure to function, Chem. Biol. Interact. 187 (2010) 10–22. N. Aurbek, H. Thiermann, L. Szinicz, P. Eyer, F. Worek, Analysis of inhibition, reactivation and aging kinetics of highly toxic organophosphorus compounds with human and pig acetylcholinesterase, Toxicology 224 (2006) 91–99. A. Bartling, F. Worek, L. Szinicz, H. Thiermann, Enzyme-kinetic investigation of different sarin analogues reacting with human acetylcholinesterase and butyrylcholinesterase, Toxicology 233 (2007) 166–172. D.M. Quinn, J. Topczewski, N. Yasapala, A. Lodge, Why is aged acetylcholinesterase so difficult to reactivate? Molecules 22 (2017). A. Mutero, M. Pralavorio, J.M. Bride, D. Fournier, Resistance-associated point mutations in insecticide-insensitive acetylcholinesterase, Proc. Natl. Acad. Sci. U. S. A. 91 (1994) 5922–5926. K.Y. Zhu, S.H. Lee, J.M. Clark, A point mutation of acetylcholinesterase associated with azinphosmethyl resistance and reduced fitness in Colorado potato beetle, Pestic. Biochem. Physiol. 55 (1996) 100–108. M.S. Williamson, G.D. Moores, S. Walsh, T. Dolden, A. Mullaley, R. Taylor, A.L. Devonshire, Mutations in the housefly acetylcholinesterase gene that confer resistance to insecticides, Struct. Func. Cholinesterases Related Proteins (1998) 548. Z. Chen, R. Newcomb, E. Forbes, J. McKenzie, P. Batterham, The acetylcholinesterase gene and organophosphorus resistance in the Australian sheep blowfly, Lucilia cuprina, Insect Biochem. Mol. Biol. 31 (2001) 805–816. M. Harel, J.L. Sussman, E. Krejci, S. Bon, P. Chanal, J. Massoulie, I. Silman, Conversion of acetylcholinesterase to butyrylcholinesterase: modeling and mutagenesis, Proc. Natl. Acad. Sci. U. S. A. 89 (1992) 10827–10831. A. Shafferman, A. Ordentlich, D. Barak, C. Kronman, H. Grosfeld, D. Stein, N. Ariel, Y. Segall, B. Velan, Enzyme engineering towards novel OP-hydrolases based on the human acetylcholinesterase template, Phosphorus Sulfur 109 (1996) 393–396. A. Vaughan, T. Rocheleau, R. ffrench-Constant, Site-directed mutagenesis of an acetylcholinesterase gene from the yellow fever mosquito Aedes aegypti confers insecticide insensitivity, Exp. Parasitol. 87 (1997) 237–244. J. Pleiss, N. Mionetto, R.D. Schmid, Probing the acyl binding site of acetylcholinesterase by protein engineering1Dedicated to Professor Hideaki Yamada in honor of his 70th birthday.1, J. Mol. Catal. B Enzym. 6 (1999) 287–296. D. Barak, D. Kaplan, A. Ordentlich, N. Ariel, B. Velan, A. Shafferman, The aromatic "trapping" of the catalytic histidine is essential for efficient catalysis in acetylcholinesterase, Biochemistry 41 (2002) 8245–8252. Y. Ashani, Z. Radic, I. Tsigelny, D.C. Vellom, N.A. Pickering, D.M. Quinn, B.P. Doctor, P. Taylor, Amino-acid-residues controlling reactivation of organophosphonyl conjugates of acetylcholinesterase by monoquaternary and bisquaternary oximes, J. Biol. Chem. 270 (1995) 6370–6380. C. Chambers, C. Luo, M. Tong, Y. Yang, A. Saxena, Probing the role of amino acids in oxime-mediated reactivation of nerve agent-inhibited human acetylcholinesterase, Toxicol. Vitro 29 (2015) 408–414. Z. Kovarik, N. Macek Hrvat, M. Katalinic, R.K. Sit, A. Paradyse, S. Zunec, K. Musilek, V.V. Fokin, P. Taylor, Z. Radic, Catalytic soman scavenging by the Y337A/F338A acetylcholinesterase mutant assisted with novel site-directed aldoximes, Chem. Res. Toxicol. 28 (2015) 1036–1044. N. Macek Hrvat, S. Zunec, P. Taylor, Z. Radic, Z. Kovarik, HI-6 assisted catalytic scavenging of VX by acetylcholinesterase choline binding site mutants, Chem. Biol. Interact. 259 (2016) 148–153. O. Mazor, O. Cohen, C. Kronman, L. Raveh, D. Stein, A. Ordentlich, A. Shafferman, Aging-resistant organophosphate bioscavenger based on polyethylene glycolconjugated F338A human acetylcholinesterase, Mol. Pharmacol. 74 (2008)

61

Chemico-Biological Interactions 292 (2018) 50–64

M. Goldsmith, Y. Ashani

organophosphate pesticides toxicity, J. Genet. 96 (2017) 187–201. [108] R.J. Richter, G.P. Jarvik, C.E. Furlong, Paraoxonase 1 (PON1) status and substrate hydrolysis, Toxicol. Appl. Pharmacol. 235 (2009) 1–9. [109] H.G. Davies, R.J. Richter, M. Keifer, C.A. Broomfield, J. Sowalla, C.E. Furlong, The effect of the human serum paraoxonase polymorphism is reversed with diazoxon, soman and sarin, Nat. Genet. 14 (1996) 334–336. [110] D.T. Yeung, J.R. Smith, R.E. Sweeney, D.E. Lenz, D.M. Cerasoli, Direct detection of stereospecific soman hydrolysis by wild-type human serum paraoxonase, FEBS J. 274 (2007) 1183–1191. [111] M. Valiyaveettil, Y. Alamneh, L. Biggemann, I. Soojhawon, B.P. Doctor, M.P. Nambiar, Efficient hydrolysis of the chemical warfare nerve agent tabun by recombinant and purified human and rabbit serum paraoxonase 1, Biochem. Biophys. Res. Commun. 403 (2010) 97–102. [112] M.W. Peterson, S.Z. Fairchild, T.C. Otto, M. Mohtashemi, D.M. Cerasoli, W.E. Chang, VX hydrolysis by human serum paraoxonase 1: a comparison of experimental and computational results, PLoS One 6 (2011) e20335. [113] M. Valiyaveettil, Y. Alamneh, P. Rezk, L. Biggemann, M.W. Perkins, A.M. Sciuto, B.P. Doctor, M.P. Nambiar, Protective efficacy of catalytic bioscavenger, paraoxonase 1 against sarin and soman exposure in Guinea pigs, Biochem. Pharmacol. 81 (2011) 800–809. [114] M. Valiyaveettil, Y. Alamneh, P. Rezk, M.W. Perkins, A.M. Sciuto, B.P. Doctor, M.P. Nambiar, Recombinant paraoxonase 1 protects against sarin and soman toxicity following microinstillation inhalation exposure in Guinea pigs, Toxicol. Lett. 202 (2011) 203–208. [115] S.M. Hodgins, S.A. Kasten, J. Harrison, T.C. Otto, Z.P. Oliver, P. Rezk, T.E. Reeves, N. Chilukuri, D.M. Cerasoli, Assessing protection against OP pesticides and nerve agents provided by wild-type HuPON1 purified from Trichoplusia ni larvae or induced via adenoviral infection, Chem. Biol. Interact. 203 (2013) 177–180. [116] S.D. Kirby, J.R. Norris, J. Richard Smith, B.J. Bahnson, D.M. Cerasoli, Human paraoxonase double mutants hydrolyze V and G class organophosphorus nerve agents, Chem. Biol. Interact. 203 (2013) 181–185. [117] D.T. Yeung, D. Josse, J.D. Nicholson, A. Khanal, C.W. McAndrew, B.J. Bahnson, D.E. Lenz, D.M. Cerasoli, Structure/function analyses of human serum paraoxonase (HuPON1) mutants designed from a DFPase-like homology model, Biochim. Biophys. Acta 1702 (2004) 67–77. [118] A. Aharoni, L. Gaidukov, S. Yagur, L. Toker, I. Silman, D.S. Tawfik, Directed evolution of mammalian paraoxonases PON1 and PON3 for bacterial expression and catalytic specialization, Proc. Natl. Acad. Sci. U. S. A. 101 (2004) 482–487. [119] M. Trovaslet-Leroy, L. Musilova, F. Renault, X. Brazzolotto, J. Misik, L. Novotny, M.T. Froment, E. Gillon, M. Loiodice, L. Verdier, P. Masson, D. Rochu, D. Jun, F. Nachon, Organophosphate hydrolases as catalytic bioscavengers of organophosphorus nerve agents, Toxicol. Lett. 206 (2011) 14–23. [120] F.H. Arnold, Directed evolution: bringing new chemistry to life, Angew Chem. Int. Ed. Engl. (2017). [121] F. Cheng, L. Zhu, U. Schwaneberg, Directed evolution 2.0: improving and deciphering enzyme properties, Chem. Commun. (J. Chem. Soc. Sect. D) 51 (2015) 9760–9772. [122] M. Goldsmith, D.S. Tawfik, Directed enzyme evolution: beyond the low-hanging fruit, Curr. Opin. Struct. Biol. 22 (2012) 406–412. [123] J.L. Porter, R.A. Rusli, D.L. Ollis, Directed evolution of enzymes for industrial biocatalysis, Chembiochem 17 (2016) 197–203. [124] V. Sachsenhauser, J.C. Bardwell, Directed evolution to improve protein folding in vivo, Curr. Opin. Struct. Biol. 48 (2017) 117–123. [125] H. Kries, R. Blomberg, D. Hilvert, De novo enzymes by computational design, Curr. Opin. Chem. Biol. 17 (2013) 221–228. [126] G. Schreiber, S.J. Fleishman, Computational design of protein-protein interactions, Curr. Opin. Struct. Biol. 23 (2013) 903–910. [127] M.D. Smith, A. Zanghellini, D. Grabs-Rothlisberger, Computational design of novel enzymes without cofactors, Meth. Mol. Biol. 1216 (2014) 197–210. [128] H.J. Wijma, D.B. Janssen, Computational design gains momentum in enzyme catalysis engineering, FEBS J. 280 (2013) 2948–2960. [129] W. Yang, L. Lai, Computational design of ligand-binding proteins, Curr. Opin. Struct. Biol. 45 (2017) 67–73. [130] K. Bernath, S. Magdassi, D.S. Tawfik, Directed evolution of protein inhibitors of DNA-nucleases by in vitro compartmentalization (IVC) and nano-droplet delivery, J. Mol. Biol. 345 (2005) 1015–1026. [131] Y. Ashani, M. Goldsmith, H. Leader, I. Silman, J.L. Sussman, D.S. Tawfik, In vitro detoxification of cyclosarin in human blood pre-incubated ex vivo with recombinant serum paraoxonases, Toxicol. Lett. 206 (2011) 24–28. [132] F. Worek, T. Seeger, M. Goldsmith, Y. Ashani, H. Leader, J.S. Sussman, D. Tawfik, H. Thiermann, T. Wille, Efficacy of the rePON1 mutant IIG1 to prevent cyclosarin toxicity in vivo and to detoxify structurally different nerve agents in vitro, Arch. Toxicol. 88 (2014) 1257–1266. [133] M. Goldsmith, Y. Ashani, R. Margalit, A. Nyska, D. Mirelman, D.S. Tawfik, A new post-intoxication treatment of paraoxon and parathion poisonings using an evolved PON1 variant and recombinant GOT1, Chem. Biol. Interact. 259 (2016) 242–251. [134] F. Campos, T. Sobrino, P. Ramos-Cabrer, B. Argibay, J. Agulla, M. Perez-Mato, R. Rodriguez-Gonzalez, D. Brea, J. Castillo, Neuroprotection by glutamate oxaloacetate transaminase in ischemic stroke: an experimental study, J. Cerebr. Blood Flow Metabol.: Official Journal of the International Society of Cerebral Blood Flow and Metabolism 31 (2011) 1378–1386. [135] A. Ruban, T. Berkutzki, I. Cooper, B. Mohar, V.I. Teichberg, Blood glutamate scavengers prolong the survival of rats and mice with brain-implanted gliomas, Invest. N. Drugs 30 (2012) 2226–2235. [136] A. Zlotnik, S.E. Gruenbaum, A.A. Artru, I. Rozet, M. Dubilet, S. Tkachov,

[137]

[138]

[139]

[140]

[141]

[142]

[143]

[144]

[145]

[146]

[147]

[148]

[149]

[150] [151]

[152]

[153]

[154]

[155]

[156]

[157]

[158]

[159]

[160]

[161] [162]

62

E. Brotfain, Y. Klin, Y. Shapira, V.I. Teichberg, The neuroprotective effects of oxaloacetate in closed head injury in rats is mediated by its blood glutamate scavenging activity: evidence from the use of maleate, J. Neurosurg. Anesthesiol. 21 (2009) 235–241. A. Ruban, I.E. Biton, A. Markovich, D. Mirelman, MRS of brain metabolite levels demonstrates the ability of scavenging of excess brain glutamate to protect against nerve agent induced seizures, Int. J. Mol. Sci. 16 (2015) 3226–3236. A. Ruban, B. Mohar, G. Jona, V.I. Teichberg, Blood glutamate scavenging as a novel neuroprotective treatment for paraoxon intoxication, J. Cerebr. Blood Flow Metabol.: Off. J. Int. Soc. Cerebral Blood Flow Metab. 34 (2014) 221–227. L.D. Stratford-Perricaudet, P. Briand, M. Perricaudet, Feasibility of adenovirusmediated gene transfer in vivo, Bone Marrow Transplant. 9 (Suppl 1) (1992) 151–152. J. Cowan, C.M. Sinton, A.W. Varley, F.H. Wians, R.W. Haley, R.S. Munford, Gene therapy to prevent organophosphate intoxication, Toxicol. Appl. Pharmacol. 173 (2001) 1–6. E.G. Duysen, K. Parikh, V. Aleti, V. Manne, O. Lockridge, N. Chilukuri, Adenovirus-mediated human paraoxonase1 gene transfer to provide protection against the toxicity of the organophosphorus pesticide toxicant diazoxon, Gene Ther. 18 (2011) 250–257. D.G. Mata, P.E. Rezk, P. Sabnekar, D.M. Cerasoli, N. Chilukuri, Investigation of evolved paraoxonase-1 variants for prevention of organophosphorous pesticide compound intoxication, J. Pharmacol. Exp. Therapeut. 349 (2014) 549–558. D.G. Mata, P. Sabnekar, C.A. Watson, P.E. Rezk, N. Chilukuri, Assessing the stoichiometric efficacy of mammalian expressed paraoxonase-1 variant I-F11 to afford protection against G-type nerve agents, Chem. Biol. Interact. 259 (2016) 233–241. M.M. Benning, J.M. Kuo, F.M. Raushel, H.M. Holden, Three-dimensional structure of phosphotriesterase: an enzyme capable of detoxifying organophosphate nerve agents, Biochemistry 33 (1994) 15001–15007. C.M. Serdar, D.T. Gibson, D.M. Munnecke, J.H. Lancaster, Plasmid involvement in parathion hydrolysis by Pseudomonas diminuta, Appl. Environ. Microbiol. 44 (1982) 246–249. L. Afriat-Jurnou, C.J. Jackson, D.S. Tawfik, Reconstructing a missing link in the evolution of a recently diverged phosphotriesterase by active-site loop remodeling, Biochemistry 51 (2012) 6047–6055. L. Afriat, C. Roodveldt, G. Manco, D.S. Tawfik, The latent promiscuity of newly identified microbial lactonases is linked to a recently diverged phosphotriesterase, Biochemistry 45 (2006) 13677–13686. C.M. Serdar, D.T. Gibson, Enzymatic hydrolysis of organophosphates: cloning and expression of a parathion hydrolase gene from Pseudomonas diminuta, Bio Technol. 3 (1985) 567. P. Segers, M. Vancanneyt, B. Pot, U. Torck, B. Hoste, D. Dewettinck, E. Falsen, K. Kersters, P. De Vos, Classification of Pseudomonas diminuta Leifson and Hugh 1954 and Pseudomonas vesicularis Busing, Doll, and Freytag 1953 in Brevundimonas gen. nov. as Brevundimonas diminuta comb. nov. and Brevundimonas vesicularis comb. nov., respectively, Int. J. Syst. Bacteriol. 44 (1994) 499–510. N. Sethunathan, T. Yoshida, A Flavobacterium sp. that degrades diazinon and parathion, Can. J. Microbiol. 19 (1973) 873–875. W.W. Mulbry, J.S. Karns, Parathion hydrolase specified by the Flavobacterium opd gene: relationship between the gene and protein, J. Bacteriol. 171 (1989) 6740–6746. K. Kawahara, A. Tanaka, J. Yoon, A. Yokota, Reclassification of a parathione-degrading Flavobacterium sp. ATCC 27551 as Sphingobium fuliginis, J. Gen. Appl. Microbiol. 56 (2010) 249–255. R. Iyer, V.G. Stepanov, B. Iken, Isolation and molecular characterization of a novel Pseudomonas putida strain capable of degrading organophosphate and aromatic compounds, Adv. Biol. Chem. 3 (2013) 564–578. E.V. Pandeeti, D. Chakka, J.P. Pandey, D. Siddavattam, Indigenous organophosphate-degrading (opd) plasmid pCMS1 of Brevundimonasdiminuta is self-transmissible and plays a key role in horizontal mobility of the opd gene, Plasmid 65 (2011) 226–231. S. Parthasarathy, H. Parapatla, A. Nandavaram, T. Palmer, D. Siddavattam, Organophosphate hydrolase is a lipoprotein and interacts with pi-specific transport system to facilitate growth of Brevundimonas diminuta using OP insecticide as source of phosphate, J. Biol. Chem. 291 (2016) 7774–7785. D.P. Dumas, H.D. Durst, W.G. Landis, F.M. Raushel, J.R. Wild, Inactivation of organophosphorus nerve agents by the phosphotriesterase from Pseudomonas diminuta, Arch. Biochem. Biophys. 277 (1990) 155–159. J.E. Kolakowski, J.J. DeFrank, S.P. Harvey, L.L. Szafraniec, W.T. Beaudry, K.H. Lai, J.R. Wild, Enzymatic hydrolysis of the chemical warfare agent VX and its neurotoxic analogues by organophosphorus hydrolase, Biocatal. Biotransform. 15 (1997) 297–312. V.K. Rastogi, J.J. DeFrank, T.C. Cheng, J.R. Wild, Enzymatic hydrolysis of RussianVX by organophosphorus hydrolase, Biochem. Biophys. Res. Commun. 241 (1997) 294–296. Y. Ashani, N. Rothschild, Y. Segall, D. Levanon, L. Raveh, Prophylaxis against organophosphate poisoning by an enzyme hydrolysing organophosphorus compounds in mice, Life Sci. 49 (1991) 367–374. L. Raveh, Y. Segall, H. Leader, N. Rothschild, D. Levanon, Y. Henis, Y. Ashani, Protection against tabun toxicity in mice by prophylaxis with an enzyme hydrolyzing organophosphate esters, Biochem. Pharmacol. 44 (1992) 397–400. C.A. Broomfield, A purified recombinant organophosphorus acid anhydrase protects mice against soman, Pharmacol. Toxicol. 70 (1992) 65–66. D.P. Dumas, S.R. Caldwell, J.R. Wild, F.M. Raushel, Purification and properties of the phosphotriesterase from Pseudomonas diminuta, J. Biol. Chem. 264 (1989)

Chemico-Biological Interactions 292 (2018) 50–64

M. Goldsmith, Y. Ashani

e47028. [190] J. Hiblot, G. Gotthard, C. Champion, E. Chabriere, M. Elias, Crystallization and preliminary X-ray diffraction analysis of the lactonase VmoLac from Vulcanisaeta moutnovskia, Acta Crystallogr Sect F Struct Biol Cryst Commun 69 (2013) 1235–1238. [191] D.F. Xiang, P. Kolb, A.A. Fedorov, M.M. Meier, L.V. Fedorov, T.T. Nguyen, R. Sterner, S.C. Almo, B.K. Shoichet, F.M. Raushel, Functional annotation and three-dimensional structure of Dr0930 from Deinococcus radiodurans, a close relative of phosphotriesterase in the amidohydrolase superfamily, Biochemistry 48 (2009) 2237–2247. [192] R. Hawwa, J. Aikens, R.J. Turner, B.D. Santarsiero, A.D. Mesecar, Structural basis for thermostability revealed through the identification and characterization of a highly thermostable phosphotriesterase-like lactonase from Geobacillus stearothermophilus, Arch. Biochem. Biophys. 488 (2009) 109–120. [193] V. Kallnik, A. Bunescu, C. Sayer, C. Brasen, R. Wohlgemuth, J. Littlechild, B. Siebers, Characterization of a phosphotriesterase-like lactonase from the hyperthermoacidophilic crenarchaeon Vulcanisaeta moutnovskia, J. Biotechnol. 190 (2014) 11–17. [194] M. Elias, J. Dupuy, L. Merone, L. Mandrich, E. Porzio, S. Moniot, D. Rochu, C. Lecomte, M. Rossi, P. Masson, G. Manco, E. Chabriere, Structural basis for natural lactonase and promiscuous phosphotriesterase activities, J. Mol. Biol. 379 (2008) 1017–1028. [195] J. Hiblot, G. Gotthard, E. Chabriere, M. Elias, Characterisation of the organophosphate hydrolase catalytic activity of SsoPox, Sci. Rep. 2 (2012) 779. [196] P. Jacquet, D. Daude, J. Bzdrenga, P. Masson, M. Elias, E. Chabriere, Current and emerging strategies for organophosphate decontamination: special focus on hyperstable enzymes, Environ. Sci. Pollut. Res. Int. 23 (2016) 8200–8218. [197] P. Jacquet, J. Hiblot, D. Daude, C. Bergonzi, G. Gotthard, N. Armstrong, E. Chabriere, M. Elias, Rational engineering of a native hyperthermostable lactonase into a broad spectrum phosphotriesterase, Sci. Rep. 7 (2017) 16745. [198] L. Merone, L. Mandrich, E. Porzio, M. Rossi, S. Muller, G. Reiter, F. Worek, G. Manco, Improving the promiscuous nerve agent hydrolase activity of a thermostable archaeal lactonase, Bioresour. Technol. 101 (2010) 9204–9212. [199] E.I. Scharff, C. Lucke, G. Fritzsch, J. Koepke, J. Hartleib, S. Dierl, H. Ruterjans, Crystallization and preliminary X-ray crystallographic analysis of DFPase from Loligo vulgaris, Acta Crystallogr D Biol Crystallogr 57 (2001) 148–149. [200] F.C. Hoskin, P. Rosenberg, M. Brzin, Re-examination of the effect of DFP on electrical and cholinesterase activity of squid giant axon, Proc. Natl. Acad. Sci. U. S. A. 55 (1966) 1231–1235. [201] F.C. Hoskin, Diisopropylphosphorofluoridate and Tabun: enzymatic hydrolysis and nerve function, Science 172 (1971) 1243–1245. [202] J. Gab, M. Melzer, K. Kehe, A. Richardt, M.M. Blum, Quantification of hydrolysis of toxic organophosphates and organophosphonates by diisopropyl fluorophosphatase from Loligo vulgaris by in situ Fourier transform infrared spectroscopy, Anal. Biochem. 385 (2009) 187–193. [203] M. Melzer, J.C. Chen, A. Heidenreich, J. Gab, M. Koller, K. Kehe, M.M. Blum, Reversed enantioselectivity of diisopropyl fluorophosphatase against organophosphorus nerve agents by rational design, J. Am. Chem. Soc. 131 (2009) 17226–17232. [204] J. Hartleib, H. Ruterjans, High-yield expression, purification, and characterization of the recombinant diisopropylfluorophosphatase from Loligo vulgaris, Protein Expr. Purif. 21 (2001) 210–219. [205] J. Koepke, E.I. Scharff, C. Lucke, H. Ruterjans, G. Fritzsch, Statistical analysis of crystallographic data obtained from squid ganglion DFPase at 0.85 angstrom resolution, Acta Crystallogr. D 59 (2003) 1744–1754. [206] M.M. Blum, M. Mustyakimov, H. Ruterjans, K. Kehe, B.P. Schoenborn, P. Langan, J.C. Chen, Rapid determination of hydrogen positions and protonation states of diisopropyl fluorophosphatase by joint neutron and X-ray diffraction refinement, Proc. Natl. Acad. Sci. U. S. A. 106 (2009) 713–718. [207] P.L. Turecek, M.J. Bossard, F. Schoetens, I.A. Ivens, PEGylation of biopharmaceuticals: a review of chemistry and nonclinical safety information of approved drugs, J Pharm Sci 105 (2016) 460–475. [208] M. Melzer, A. Heidenreich, F. Dorandeu, J. Gab, K. Kehe, H. Thiermann, T. Letzel, M.M. Blum, In vitro and in vivo efficacy of PEGylated diisopropyl fluorophosphatase (DFPase), Drug Test. Anal. 4 (2012) 262–270. [209] S. Chakraborti, B.J. Bahnson, Crystal structure of human senescence marker protein 30: insights linking structural, enzymatic, and physiological functions, Biochemistry 49 (2010) 3436–3444. [210] S.H. Scott, B.J. Bahnson, Senescence marker protein 30: functional and structural insights to its unknown physiological function, Biomol. Concepts 2 (2011) 469–480. [211] S. Li, X. Chen, W. Lai, M. Hu, X. Zhong, S. Tan, H. Liang, Downregulation of SMP30 in senescent human lens epithelial cells, Mol. Med. Rep. 16 (2017) 4022–4028. [212] J.S. Little, C.A. Broomfield, M.K. Fox-Talbot, L.J. Boucher, B. MacIver, D.E. Lenz, Partial characterization of an enzyme that hydrolyzes sarin, soman, tabun, and diisopropyl phosphorofluoridate (DFP), Biochem. Pharmacol. 38 (1989) 23–29. [213] Y. Kondo, A. Ishigami, S. Kubo, S. Handa, K. Gomi, K. Hirokawa, N. Kajiyama, T. Chiba, K. Shimokado, N. Maruyama, Senescence marker protein-30 is a unique enzyme that hydrolyzes diisopropyl phosphorofluoridate in the liver, FEBS Lett. 570 (2004) 57–62. [214] T. Belinskaya, N. Pattabiraman, R. diTargiani, M. Choi, A. Saxena, Differences in amino acid residues in the binding pockets dictate substrate specificities of mouse senescence marker protein-30, human paraoxonase1, and squid diisopropylfluorophosphatase, Biochim. Biophys. Acta 1824 (2012) 701–710. [215] M.S. Choi, A. Saxena, N. Chilukuri, A strategy for the production of soluble human

19659–19665. [163] H. Shim, S.B. Hong, F.M. Raushel, Hydrolysis of phosphodiesters through transformation of the bacterial phosphotriesterase, J. Biol. Chem. 273 (1998) 17445–17450. [164] S.B. Hong, F.M. Raushel, Stereochemical constraints on the substrate specificity of phosphotriesterase, Biochemistry 38 (1999) 1159–1165. [165] M. Chen-Goodspeed, M.A. Sogorb, F. Wu, S.B. Hong, F.M. Raushel, Structural determinants of the substrate and stereochemical specificity of phosphotriesterase, Biochemistry 40 (2001) 1325–1331. [166] E. Ghanem, F.M. Raushel, Detoxification of organophosphate nerve agents by bacterial phosphotriesterase, Toxicol. Appl. Pharmacol. 207 (2005) 459–470. [167] L. Briseno-Roa, C.M. Timperley, A.D. Griffiths, A.R. Fersht, Phosphotriesterase variants with high methylphosphonatase activity and strong negative trade-off against phosphotriesters, Protein Eng. Des. Sel. 24 (2011) 151–159. [168] S. Gopal, V. Rastogi, W. Ashman, W. Mulbry, Mutagenesis of organophosphorus hydrolase to enhance hydrolysis of the nerve agent VX, Biochem. Biophys. Res. Commun. 279 (2000) 516–519. [169] T.E. Reeves, M.E. Wales, J.K. Grimsley, P. Li, D.M. Cerasoli, J.R. Wild, Balancing the stability and the catalytic specificities of OP hydrolases with enhanced V-agent activities, Protein Eng. Des. Sel. 21 (2008) 405–412. [170] Y.S. Jeong, J.M. Choi, H.H. Kyeong, J.Y. Choi, E.J. Kim, H.S. Kim, Rational design of organophosphorus hydrolase with high catalytic efficiency for detoxifying a Vtype nerve agent, Biochem. Biophys. Res. Commun. 449 (2014) 263–267. [171] C.M. Cho, A. Mulchandani, W. Chen, Bacterial cell surface display of organophosphorus hydrolase for selective screening of improved hydrolysis of organophosphate nerve agents, Appl. Environ. Microbiol. 68 (2002) 2026–2030. [172] C. Roodveldt, D.S. Tawfik, Directed evolution of phosphotriesterase from Pseudomonas diminuta for heterologous expression in Escherichia coli results in stabilization of the metal-free state, Protein Eng. Des. Sel. 18 (2005) 51–58. [173] S.Y. McLoughlin, C. Jackson, J.W. Liu, D. Ollis, Increased expression of a bacterial phosphotriesterase in Escherichia coli through directed evolution, Protein Expr. Purif. 41 (2005) 433–440. [174] C.M. Hill, W.S. Li, J.B. Thoden, H.M. Holden, F.M. Raushel, Enhanced degradation of chemical warfare agents through molecular engineering of the phosphotriesterase active site, J. Am. Chem. Soc. 125 (2003) 8990–8991. [175] M.M. Benning, H. Shim, F.M. Raushel, H.M. Holden, High resolution X-ray structures of different metal-substituted forms of phosphotriesterase from Pseudomonas diminuta, Biochemistry 40 (2001) 2712–2722. [176] C. Jackson, H.K. Kim, P.D. Carr, J.W. Liu, D.L. Ollis, The structure of an enzymeproduct complex reveals the critical role of a terminal hydroxide nucleophile in the bacterial phosphotriesterase mechanism, Biochim. Biophys. Acta 1752 (2005) 56–64. [177] L. Briseno-Roa, J. Hill, S. Notman, D. Sellers, A.P. Smith, C.M. Timperley, J. Wetherell, N.H. Williams, G.R. Williams, A.R. Fersht, A.D. Griffiths, Analogues with fluorescent leaving groups for screening and selection of enzymes that efficiently hydrolyze organophosphorus nerve agents, J. Med. Chem. 49 (2006) 246–255. [178] M. Goldsmith, S. Eckstein, Y. Ashani, P. Greisen Jr., H. Leader, J.L. Sussman, N. Aggarwal, S. Ovchinnikov, D.S. Tawfik, D. Baker, H. Thiermann, F. Worek, Catalytic efficiencies of directly evolved phosphotriesterase variants with structurally different organophosphorus compounds in vitro, Arch. Toxicol. 90 (2016) 2711–2724. [179] F. Worek, T. Seeger, G. Reiter, M. Goldsmith, Y. Ashani, H. Leader, J.L. Sussman, N. Aggarwal, H. Thiermann, D.S. Tawfik, Post-exposure treatment of VX poisoned Guinea pigs with the engineered phosphotriesterase mutant C23: a proof-of-concept study, Toxicol. Lett. 231 (2014) 45–54. [180] T. Wille, K. Neumaier, M. Koller, C. Ehinger, N. Aggarwal, Y. Ashani, M. Goldsmith, J.L. Sussman, D.S. Tawfik, H. Thiermann, F. Worek, Single treatment of VX poisoned Guinea pigs with the phosphotriesterase mutant C23AL: intraosseous versus intravenous injection, Toxicol. Lett. 258 (2016) 198–206. [181] S.Y. Bae, J.M. Myslinski, L.R. McMahon, J.J. Height, A.N. Bigley, F.M. Raushel, S.P. Harvey, An OPAA enzyme mutant with increased catalytic efficiency on the nerve agents sarin, soman, and GP, Enzym. Microb. Technol. 112 (2018) 65–71. [182] I. Horne, T.D. Sutherland, R.L. Harcourt, R.J. Russell, J.G. Oakeshott, Identification of an opd (organophosphate degradation) gene in an Agrobacterium isolate, Appl. Environ. Microbiol. 68 (2002) 3371–3376. [183] T. Wille, C. Scott, H. Thiermann, F. Worek, Detoxification of G- and V-series nerve agents by the phosphotriesterase OpdA, Biocatal. Biotransform. 30 (2012) 203–208. [184] S.B. Bird, T.D. Sutherland, C. Gresham, J. Oakeshott, C. Scott, M. Eddleston, OpdA, a bacterial organophosphorus hydrolase, prevents lethality in rats after poisoning with highly toxic organophosphorus pesticides, Toxicology 247 (2008) 88–92. [185] C. Gresham, C. Rosenbaum, R.J. Gaspari, C.J. Jackson, S.B. Bird, Kinetics and efficacy of an organophosphorus hydrolase in a rodent model of methyl-parathion poisoning, Acad. Emerg. Med. 17 (2010) 736–740. [186] C.J. Jackson, A. Carville, J. Ward, K. Mansfield, D.L. Ollis, T. Khurana, S.B. Bird, Use of OpdA, an organophosphorus (OP) hydrolase, prevents lethality in an African green monkey model of acute OP poisoning, Toxicology 317 (2014) 1–5. [187] L. Merone, L. Mandrich, M. Rossi, G. Manco, A thermostable phosphotriesterase from the archaeon Sulfolobus solfataricus: cloning, overexpression and properties, Extremophiles 9 (2005) 297–305. [188] E. Porzio, L. Merone, L. Mandrich, M. Rossi, G. Manco, A new phosphotriesterase from Sulfolobus acidocaldarius and its comparison with the homologue from Sulfolobus solfataricus, Biochimie 89 (2007) 625–636. [189] J. Hiblot, G. Gotthard, E. Chabriere, M. Elias, Structural and enzymatic characterization of the lactonase SisLac from Sulfolobus islandicus, PLoS One 7 (2012)

63

Chemico-Biological Interactions 292 (2018) 50–64

M. Goldsmith, Y. Ashani

[216]

[217] [218] [219] [220]

[221]

[222]

[223]

[224]

[225]

[226]

[227]

[228] [229]

[230]

[231]

[232]

[233]

[234]

[235] T.K. Ng, L.R. Gahan, G. Schenk, D.L. Ollis, Altering the substrate specificity of methyl parathion hydrolase with directed evolution, Arch. Biochem. Biophys. 573 (2015) 59–68. [236] J. Xie, Y. Zhao, H. Zhang, Z. Liu, Z. Lu, Improving methyl parathion hydrolase to enhance its chlorpyrifos-hydrolysing efficiency, Lett. Appl. Microbiol. 58 (2014) 53–59. [237] C. Yang, N. Cai, M. Dong, H. Jiang, J. Li, C. Qiao, A. Mulchandani, W. Chen, Surface display of MPH on Pseudomonas putida JS444 using ice nucleation protein and its application in detoxification of organophosphates, Biotechnol. Bioeng. 99 (2008) 30–37. [238] J. Tian, P. Wang, S. Gao, X. Chu, N. Wu, Y. Fan, Enhanced thermostability of methyl parathion hydrolase from Ochrobactrum sp. M231 by rational engineering of a glycine to proline mutation, FEBS J. 277 (2010) 4901–4908. [239] M. Purg, A. Pabis, F. Baier, N. Tokuriki, C. Jackson, S.C.L. Kamerlin, Probing the mechanisms for the selectivity and promiscuity of methyl parathion hydrolase, Philos. Trans. Royal Soc. A: Math. Phys. Eng. Sci. 374 (2016). [240] A. Zanghellini, De novo computational enzyme design, Curr. Opin. Biotechnol. 29 (2014) 132–138. [241] G.B. Akcapinar, O.U. Sezerman, Computational approaches for de novo design and redesign of metal-binding sites on proteins, Biosci. Rep. 37 (2017). [242] S.D. Khare, Y. Kipnis, P. Greisen Jr., R. Takeuchi, Y. Ashani, M. Goldsmith, Y. Song, J.L. Gallaher, I. Silman, H. Leader, J.L. Sussman, B.L. Stoddard, D.S. Tawfik, D. Baker, Computational redesign of a mononuclear zinc metalloenzyme for organophosphate hydrolysis, Nat. Chem. Biol. 8 (2012) 294–300. [243] A. Ozgur, Y. Tutar, Therapeutic proteins: a to Z, Protein Pept. Lett. 20 (2013) 1365–1372. [244] J.H. Lin, Pharmacokinetics of biotech drugs: peptides, proteins and monoclonal antibodies, Curr. Drug Metabol. 10 (2009) 661–691. [245] K. Rehman, M.S. Hamid Akash, B. Akhtar, M. Tariq, A. Mahmood, M. Ibrahim, Delivery of therapeutic proteins: challenges and strategies, Curr. Drug Targets 17 (2016) 1172–1188. [246] C.J. Roberts, Therapeutic protein aggregation: mechanisms, design, and control, Trends Biotechnol. 32 (2014) 372–380. [247] K.D. Ratanji, J.P. Derrick, R.J. Dearman, I. Kimber, Immunogenicity of therapeutic proteins: influence of aggregation, J. Immunot. 11 (2014) 99–109. [248] D.W. Scott, A.S. De Groot, Can we prevent immunogenicity of human protein drugs? Ann. Rheum. Dis. 69 (Suppl 1) (2010) i72–76. [249] A.S. Rosenberg, A.R. Pariser, B. Diamond, L. Yao, L.A. Turka, E. Lacana, P.S. Kishnani, A role for plasma cell targeting agents in immune tolerance induction in autoimmune disease and antibody responses to therapeutic proteins, Clin. Immunol. 165 (2016) 55–59. [250] H.A. Lagasse, A. Alexaki, V.L. Simhadri, N.H. Katagiri, W. Jankowski, Z.E. Sauna, C. Kimchi-Sarfaty, Recent advances in (therapeutic protein) drug development, F1000Res 6 (2017) 113. [251] J.K. Dozier, M.D. Distefano, Site-specific PEGylation of therapeutic proteins, Int. J. Mol. Sci. 16 (2015) 25831–25864. [252] S.I. Lim, I. Kwon, Bioconjugation of therapeutic proteins and enzymes using the expanded set of genetically encoded amino acids, Crit. Rev. Biotechnol. 36 (2016) 803–815. [253] J. Hall, S. Prabhakar, L. Balaj, C.P. Lai, R.A. Cerione, X.O. Breakefield, Delivery of therapeutic proteins via extracellular vesicles: review and potential treatments for Parkinson's disease, glioma, and schwannoma, Cell. Mol. Neurobiol. 36 (2016) 417–427. [254] A. Bolhassani, B.S. Jafarzade, G. Mardani, In vitro and in vivo delivery of therapeutic proteins using cell penetrating peptides, Peptides 87 (2017) 50–63. [255] C. Kimchi-Sarfaty, T. Schiller, N. Hamasaki-Katagiri, M.A. Khan, C. Yanover, Z.E. Sauna, Building better drugs: developing and regulating engineered therapeutic proteins, Trends Pharmacol. Sci. 34 (2013) 534–548. [256] K. Izutsu, Stabilization of therapeutic proteins in aqueous solutions and freezedried solids: an overview, Meth. Mol. Biol. 1129 (2014) 435–441.

senescence marker protein-30 in Escherichia coli, Biochem. Biophys. Res. Commun. 393 (2010) 509–513. N.K. Vyas, A. Nickitenko, V.K. Rastogi, S.S. Shah, F.A. Quiocho, Structural insights into the dual activities of the nerve agent degrading organophosphate anhydrolase/prolidase, Biochemistry 49 (2010) 547–559. T.C. Cheng, J.J. DeFrank, V.K. Rastogi, Alteromonas prolidase for organophosphorus G-agent decontamination, Chem. Biol. Interact. 119–120 (1999) 455–462. J.J. DeFrank, T.C. Cheng, Purification and properties of an organophosphorus acid anhydrase from a halophilic bacterial isolate, J. Bacteriol. 173 (1991) 1938–1943. B. Zwanenburg, M. Mikołajczyk, P. Kiełbasiński, Enzymes in Action: green Solutions for Chemical Problems, Kluwer Academic Publishers, Boston, 2000. C.M. Hill, F. Wu, T.C. Cheng, J.J. DeFrank, F.M. Raushel, Substrate and stereochemical specificity of the organophosphorus acid anhydrolase from Alteromonas sp. JD6.5 toward p-nitrophenyl phosphotriesters, Bioorg. Med. Chem. Lett 10 (2000) 1285–1288. I. Petrikovics, W.D. McGuinn, D. Sylvester, P. Yuzapavik, J. Jiang, J.L. Way, D. Papahadjopoulos, K. Hong, R. Yin, T.C. Cheng, J.J. DeFrank, In vitro studies on sterically stabilized liposomes (SL) as enzyme carriers in organophosphorus (OP) antagonism, Drug Deliv. 7 (2000) 83–89. C.M. Daczkowski, S.D. Pegan, S.P. Harvey, Engineering the organophosphorus acid anhydrolase enzyme for increased catalytic efficiency and broadened stereospecificity on Russian VX, Biochemistry 54 (2015) 6423–6433. A. Lupi, R. Tenni, A. Rossi, G. Cetta, A. Forlino, Human prolidase and prolidase deficiency: an overview on the characterization of the enzyme involved in proline recycling and on the effects of its mutations, Amino Acids 35 (2008) 739–752. P. Wilk, M. Uehlein, J. Kalms, H. Dobbek, U. Mueller, M.S. Weiss, Substrate specificity and reaction mechanism of human prolidase, FEBS J. 284 (2017) 2870–2885. A. Stepankova, J. Duskova, T. Skalova, J. Hasek, T. Koval, L.H. Ostergaard, J. Dohnalek, Organophosphorus acid anhydrolase from Alteromonas macleodii: structural study and functional relationship to prolidases, Acta Crystallogr Sect F Struct Biol Cryst Commun 69 (2013) 346–354. Q. Wang, M. Sun, H. Zhang, C. Huang, Purification and properties of soman-hydrolyzing enzyme from human liver, J. Biochem. Mol. Toxicol. 12 (1998) 213–217. M. Costante, L. Biggemann, Y. Alamneh, I. Soojhawon, R. Short, S. Nigam, G. Garcia, B.P. Doctor, M. Valiyaveettil, M.P. Nambiar, Hydrolysis potential of recombinant human skin and kidney prolidase against diisopropylfluorophosphate and sarin by in vitro analysis, Toxicol. Vitro 26 (2012) 182–188. S.H. Wang, Q.W. Zhi, M.J. Sun, Dual activities of human prolidase, Toxicol. Vitro 20 (2006) 71–77. V. Aleti, G.B. Reddy, K. Parikh, P. Arun, N. Chilukuri, Persistent and high-level expression of human liver prolidase in vivo in mice using adenovirus, Chem. Biol. Interact. 203 (2013) 191–195. P.E. Rezk, P. Zdenka, P. Sabnekar, T. Kajih, D.G. Mata, C. Wrobel, D.M. Cerasoli, N. Chilukuri, An in vitro and in vivo evaluation of the efficacy of recombinant human liver prolidase as a catalytic bioscavenger of chemical warfare nerve agents, Drug Chem. Toxicol. 38 (2015) 37–43. C. Zhongli, L. Shunpeng, F. Guoping, Isolation of methyl parathion-degrading strain M6 and cloning of the methyl parathion hydrolase gene, Appl. Environ. Microbiol. 67 (2001) 4922–4925. H. Liu, J.J. Zhang, S.J. Wang, X.E. Zhang, N.Y. Zhou, Plasmid-borne catabolism of methyl parathion and p-nitrophenol in Pseudomonas sp. strain WBC-3, Biochem. Biophys. Res. Commun. 334 (2005) 1107–1114. R. Zhang, Z. Cui, X. Zhang, J. Jiang, J.-D. Gu, S. Li, Cloning of the organophosphorus pesticide hydrolase gene clusters of seven degradative bacteria isolated from a methyl parathion contaminated site and evidence of their horizontal gene transfer, Biodegradation 17 (2006) 465–472. Y.-J. Dong, M. Bartlam, L. Sun, Y.-F. Zhou, Z.-P. Zhang, C.-G. Zhang, Z. Rao, X.E. Zhang, Crystal structure of methyl parathion hydrolase from Pseudomonas sp. WBC-3, J. Mol. Biol. 353 (2005) 655–663.

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