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Bacterial class A acid phosphatases as versatile tools in organic synthesis

van Herk, T.

Publication date2008

Link to publication

Citation for published version (APA):van Herk, T. (2008). Bacterial class A acid phosphatases as versatile tools in organicsynthesis.

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Chapter 1

General introduction

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1. Chemical phosphorylations The role of phosphate esters is vitally important for all cell processes.[1-3] They play an essential part in photosynthesis, carbohydrate and lipid metabolism, the nitrogen cycle, immune response, host-pathogen interactions, transmembrane signaling, activation of metabolites, cellular control by protein phosphorylation and in numerous other biochemical reactions. Further, phosphorus is part of the backbone of both DNA and RNA, and phospholipids are the main structural components of all cellular membranes. A number of essential cofactors or cosubstrates for enzyme-catalyzed reactions of significant synthetic importance involve phosphate esters. For instance nicotinamide adenine dinucleotide phosphate in the oxidized (NADP+) or reduced (NADPH) form are essential cofactors for some enzymatic redox reactions[4] in which glucose-6-phosphate (G6P) can be used to regenerate NADPH.[5,6] Dihydroxyacetone phosphate (DHAP) is needed for enzymatic aldol-reactions using a variety of DHAP dependent aldolases,[7] and adenosine triphosphate (ATP) represents the energy-rich phosphate donor for most biological and synthetic phosphorylation reactions.[8-10] Phosphate prodrugs have been successfully utilized to overcome a variety of drug delivery problems that might otherwise have compromised the therapeutic utilities of the parent drug.[11] The ionic nature of the phosphate group in these prodrugs may significantly improve the solubility and dissolution rate of poorly soluble drugs thereby increasing its bioavailability. In contrast to this, various prodrug approaches involving lipophilic phosphate masking groups have been devised,[12,13] since the high polarity of monophosphate ester precludes their cellular uptake. In addition to their biological significance, phosphate esters are useful synthetic intermediates that can be used as a source of organolithium compounds,[14] be dehydrated to yield alkenes[15] or act as substrates for stereoselective displacement with Grignard reagents.[16,17] Given the importance of this functional group it is not surprising that many methods have been developed for the introduction of phosphate esters into compounds. The methods that currently exist for the introduction of a phosphate group into a substrate molecule largely depend on the substrate itself, since functional group tolerance is the key to facilitating efficient phosphorylation. Chemical phosphorylation The development of phosphorus chemistry began with Hennig Brandt,[18] who isolated elemental phosphorus from urine in 1669. In 1774 Wilhelm Scheele synthesized phosphoric acid by adding nitric acid to phosphorus. It was not until 1809 that two of the most valuable compounds involved in organophosphorus chemistry were isolated: phosphorus trichloride (PCl3) and phosphorus pentachloride (PCl5), first synthesized by John Davy by burning phosphorus in the presence of chlorine gas. Jean Lassaigne first reported the esterification of phosphoric acid with ethyl alcohol in 1820. Another very important compound in many areas of organophosphorus chemistry is phosphorus oxychloride (POCl3), which was first reported by Charles Wurtz in 1846. These discoveries set the stage for much of the phosphorus chemistry that followed. Numerous phosphorylation methods exist, some of them more widely applicable, some very exotic and specific. Some are used to produce monoesters, others for di- and tri-esters. Here an overview of the most general and frequently used methods to prepare phosphomonoesters is given.


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Figure 1. Three classes of phosphorus compounds that are most frequently used in the synthesis of biologically important phosphate esters and their analogues are tetra-coordinated P(V) compounds (oxidation state +5), tri-coordinated P(III) compounds (oxidation state +3), and tetra-coordinated P(III) compounds (oxidation state +3)

To synthesize a phosphorus-containing compound with a given structural feature, chemists have two kinds of phosphorus chemistry at their disposal, referred to as P(V) and P(III) chemistry (Figure 1). The choice between them is often not easy, as both of them have their own strong merits. Use of pentavalent phosphorus reagents in the preparation of phosphomonoesters P(V) compounds (e.g. phosphate esters) have a tetrahedral geometry (Figure 1), and their chemistry is dominated by the presence of a very stable phosphoryl group (P=O) for which formation often is a driving force for reactions. The phosphorus atom is a hard, electrophilic center and is subject to reactions with hard nucleophiles. P(V) compounds are stable during storage and convenient to handle. However, they have disadvantages of typically being less efficient in synthetic transformations, as they react significantly more slowly, even upon activation with condensing agents, compared to tervalent P(III) derivatives. Phosphoric acid. From the perspective of green chemistry, the direct catalytic condensation of equimolar amounts of phosphoric acid (H3PO4) and alcohols is attractive for the synthesis of phosphoric acid monoesters, especially for industrial-scale synthesis, since the reaction produces only water as a by-product. However, the methods described thus far are not very successful. In the pioneering work by Honjo et al.[19] the synthesis of 2’,3’-O-isopropylidene ribonucleoside 5’-monophosphates from the corresponding ribonucleosides and 5 equivalents of H3PO4 in the presence of 10 equivalents of tributylamine under reflux condition in dimethylformamide was reported. A method for synthesizing phosphate monoesters by the dehydrative condensation of equimolar amounts of H3PO4 and alcohols promoted by 2 equivalents of nucleophilic bases such as N-butylimidazole in the presence of 2 equivalents of tributylamine was developed.[20] Sakakura et al.[21] describe the use of oxorhenium(VII) complexes as extremely active catalysts for the direct condensation of H3PO4 with nearly equimolar amounts of alcohols to give the corresponding phosphoric acid monoester.

Pyrophosphates. Free pyrophosphoric acid (phosphoric anhydride) and pyrophosphoric diesters are stable under standard basic conditions and serve as phosphorylating agents only in some limited cases.[22] An example is the trichloroacetronitrile aided phosphorylation using a syn-dialkyl pyrophosphoric diester. Among the tetra-alkyl pyrophosphates,

symmetrical derivatives, e.g. tetra-p-nitrophenyl pyrophosphate, are widely employable as monophosphorylating agents.[23]

Phosphoric acid monoesters. The application of phosphate monoesters as phosphorylating agents can be divided into two groups based upon the nature of the reaction they undergo. The first are those which undergo an ester exchange

Chapter 1

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reaction in which RO- is a suitable leaving group. The most commonly used leaving group is the p-nitrophenoxy group.[24] The second group of phosphomonoesters employed as phosphorylating reagents are alkyl esters in which the alkyl group functions as a blocking group and where the condensation must be performed in the presence of a suitable activating agent, usually a carbodiimide or an arenesulfonyl chloride. A monoester can be prepared after the blocking group is removed from the synthesized diester (see below). A popular reagent of this type appears to be 2-cyanoethyl phosphate[25] used with dicyclohexylcarbodiimide. Once the diester is formed the cyanoethyl blocking group may be removed under mildly alkaline conditions.

5- and 6-membered cyclic phosphorylating agents. Cyclic acyl phosphates and cyclic enediol phosphates are extremely reactive phosphorylating reagents.[26] Derivatives are introduced as alternative reagents for the synthesis of oligonucleotides and other naturally occurring phosphate esters. As early as 1966, o-phenylene

phosphochloridate[27] (2-chloro-2-oxo-1,3,2-benzodioxaphosphole) was introduced. Its pronounced reactivity is demonstrated by the facile phosphorylation of t-butyl alcohol and a variety of sterically hindered alcohols within 10 minutes in the presence of triethylamine, in THF or p-dioxane at 20°C. The intermediate 2-alkoxy derivatives (phosphotriesters) are then quantitatively converted to the o-hydroxyphenyl phosphate esters which are generally stable and can be isolated as crystalline salts. Conversion into the monophosphate ester is accomplished by a) oxidation with excess bromine/water in aqueous barium acetate, b) excess periodic acid in aqueous solution, or c) lead(IV) acetate in dioxane solution, all reactions being performed at room temperature.

Chlorophosphates. Three types of chlorophosphates, i.e. phosphoryl chloride (POCl3), phosphorodichloridates [(RO)POCl2], and phosphorochloridates [(RO)2POCl], are among the most widely employed

phosphorylating agents for the synthesis of organic phosphoric acid derivatives. Among the chlorophosphates, POCl3 and phosphorochloridates are used to prepare phosphomonoesters. Phosphoryl chloride. The use of POCl3 as a phosphorylating agent was first proposed in 1857. It is one of the most widely used phosphorylating agents for alcohols. The reaction of alcohols with POCl3 in the presence of water and pyridine provides phosphate monoesters along with pyridine hydrochloride as a byproduct. Since POCl3 is very reactive, phosphate diesters and triesters are produced as byproducts when alcohols are reacted with an equimolar amount of POCl3. Therefore, an excess amount of POCl3 is required for the selective synthesis of phosphate monoesters. When first applied to the synthesis of 5’-nucleotides, low yields and a lack of specificity were obtained. Treatment of an unprotected ribonucleoside with partially hydrolyzed POCl3 at 0°C in trimethyl or triethyl phosphate resulted in a selective synthesis of 5’-nucleotides.[28] Using this method, the primary hydroxyl of a number of nucleosides[29] has been phosphorylated. Alternatively, selective phosphorylation of a primary hydroxyl group has been achieved with POCl3 in acetonitrile with added pyridine and water via an active phosphorylating agent, trichloropyrophosphopyridinium chloride, which is an adduct of tetrachloropyrophosphate and pyridinium chloride.[30]


13

Phosphorochloridates. The use of phosphorochloridates as phosphorylating reagents provides a suitably protected, organic soluble phosphate triester intermediate which can be deprotected to result in the desired phosphomonoester. Reaction of a phosphorochloridate with an alcohol is carried out either through a) the formation of an alkoxide[31,32] necessarily limiting its application to those substrates with compatible functional groups or b) by using proton scavengers such as pyridine[33] or triethylamine with or without a nucleophilic catalyst[34-36] or c) by employing Lewis acid catalysts such as TiCl4[37] or Ti(t-BuO)4

[38] or d) using nucleophilic catalysis with DMAP.[39,40] Although widely used, these reaction conditions are not always compatible with base-sensitive functional groups present in the substrates and the stability of the chlorophosphate is sometimes limited. Another major drawback to the phosphorochloridates is their relative low reactivity. This problem was overcome through the use of phosphorochloridites (see below).

N-Phosphoryl oxazolidinones. N-Phosphoryl oxazolidinones can be used as effective phosphate sources in the presence of lithium and magnesium alkoxides.[41] They are developed as an alternative to POCl3-equivalents that function just as effectively as phosphorochloridates, but have improved stability and contain a non-nucleophilic counter-ion. These reagents are attractive since they are easily prepared, have a long shelf life, and are easy to handle. However, given the need to generate highly

basic alkoxides, reactions are limited to substrates with base-tolerant functional groups. Milder reaction conditions to facilitate this process have been found.[42] The use of a Lewis acid catalyst like Cu(OTf)2 turned out to give good phosphorylations with more sensitive substrates. Use of trivalent phosphorus reagents in the preparation of phosphomonoesters P(III) compounds, e.g. phosphite triesters (Figure 1) have the shape of a trigonal pyramid with a lone electron pair located on the phosphorus atom. Due to this, the phosphorus center in these compounds is basic and is a soft nuclophile that may react with various, preferably soft, electrophiles. However, when the phosphorus center is protonated, or when it bears electron-withdrawing substituents, it may also react with nucleophiles. The reactions with electrophiles and nucleophiles are both rapid and make P(III) derivatives attractive phosphorylating agents although they are often difficult to handle due to their high reactivity which makes them susceptible to hydrolysis and spontaneous oxidation upon storage, even by atmospheric oxygen.

Phosphorochloridites. Treatment of alcohol substrates with these highly reactive reagents in the presence of an acid scavenger like pyridine produces phosphite triester intermediates. Subsequent oxidation with iodine in water gives the desired phosphate triester in virtually quantitative yields.[43]

Deprotection results in the phosphomonoester. This method proved quite useful, but the phosphorochloridites are very sensitive to moisture and must be prepared fresh each time, however, diethyl phosphorochloridite is commercially available.

Phosphoramidites. Perhaps the most widely used and most successful of all phosphorylation techniques has been the use of phosphite esters that are usually introduced via reaction of a phosphoramidite with an alcohol.[33,44] When phosphoramidites are activated by weak acids such as N-methylanilinium trifluoroacetate,[45] 1H-tetrazole,[46] 5-methyltetrazole,[44] 5-methylthiotetrazole,

5-(4-nitrophenyl)tetrazole,[47] etc., they react with alcohols, forming phosphites, which can be transferred to the phosphates by oxidation. Under certain circumstances oxidation of the P(III) intermediates to P(V) can be a troublesome step especially for substrates containing

Chapter 1

14

functional groups such as alkenes that are not tolerant to the oxidizing agents. From a synthetic point of view, diethylamidites, diisopropylamidites, and morpholidites are the most efficient reagents. The phosphoramidite reagents are usually easy to prepare, but require care in handling and have limited shelf lives. In general, phosphoramidites are more stable than the corresponding phosphorochloridites and some of the reagents can be purified by silica gel chromatography. The use of phosphoramidites in the chemistry of natural products such as phosphorylated saccharides, nucleotides, phospholipids and other phosphorus-containing natural compounds and their analogues has been reviewed by Nifantiev et al.[48]

H-Phosphonate monoesters. H-phosphonates[49] belong to a special class of P(III) reagents (Figure 1). Due to the presence of a phosphoryl group and the tetrahedral structure, they bear strong resemblance to P(V) derivatives, but the oxidation state +3 clearly relates them to P(III) compounds. However, in

contrast to the latter, they lack a lone electron pair on the phosphorus center. A feature which is unique for compounds of this type is the presence of a P-H bond, usually emphasized in their names (H-phosphonates). Since H-phosphonates are structurally related to P(V) derivatives, one can predict that the phosphorus center should be electrophilic, and the compounds, although harboring the phosphorus atom in the +3 oxidation state, but lacking of a lone pair of electrons, will be less prone to oxidation than P(III) derivatives. H-phosphonate derivatives can be easily converted into various P(V) derivatives using different oxidizing reagents (e.g. iodine/water, elemental sulfur, elemental selenium etc.). However, some of the H-phosphonates need to be activated for example into tervalent bis-trimethylsilyl alkyl phosphite derivatives[50] to react with electrophiles to produce phosphate monoesters.[51,52] A fluoren-9-ylmethyl ester as P-protecting group can also be used to facilitate the oxidation step.[53] This protecting group can be easily removed with piperidine to provide the desired phosphomonoester. Protecting groups of phosphoric acid. Because the preparation of biological molecules often necessitates the synthesis of appreciable quantities for study, it is advantageous to have neutral phosphate esters as intermediates which are amenable to large scale separation techniques employing organic solvents. The major requirement is then a phosphorylating reagent whose blocking groups may be removed without affecting the stability of the remainder of the molecule upon conversion of trialkyl phosphates into the corresponding monoalkyl phosphates.[54] In the preparation of phosphomonoesters two types of protecting groups are possible. In one type the protector is removed via a P-O bond fission and in the other through a C-O bond cleavage. In the former case, it is possible that a P-O bond fission occurs at undesired positions. Use of the latter protector may prevent such a side reaction. Thus the protecting groups belonging to the latter strategy are much more widely employed. Aryl Groups. Aryl groups are widely utilized as protector. The benzyl-, dibenzyl- and diphenyl-protecting groups are removable by hydrogenolysis, using platinum or palladium metal as the catalyst.[55,56] Some kinds of phenyl derivatives (e.g. 4-chloro-2-nitrophenyl or 2-chloromethyl-4-nitrophenyl) can be removed by basic hydrolysis.[57-60] The base hydrolysis under harsh conditions very often brings about undesired cleavage of phosphates. In contrast, removal of an o- or p-chlorophenyl protecting group by N1,N1,N2,N2-tetramethylguanidinium syn-p-nitrobenzaldoxymate or syn-pyridin-2-aldoxymate is achievable at room temperature and no side reactions occur.[61-63] The 8-quinolyl phosphate protecting group is also hydrolyzed under mild conditions with the assistance of a stoichiometric amount of ZnCl2 or CuCl2.[64,65] The 2-t-butylphenyl group,


15

which is removed by hydrogenolysis on platinum oxide, can be used in some limited cases.[66] 2-Hydroxyphenyl is a useful protecting group in conversion of phosphodiesters to phosphomonoesters. The deprotection is carried out by oxidative treatment.[27,67] The fluoren-9-ylmethyl P-protecting group can be cleaved under extremely mild conditions which do not affect O-benzyl, O-benzylidene, P-benzoyl or O-acetyl protecting groups.[53] Alkyl groups. Methyl is an easily employable protecting group. Deprotection is effected by thiophenol/triethylamine,[68] metal thiophenoxides,[69,70] dimethyl sulfide/methanesulfonic acid,[71] t-butylamine,[72,73] etc. An allyl group is an excellent protector, removable using a palladium(0) catalyst in the presence of nucleophiles such as primary or secondary amines and their salts or formic acid.[74-77] Removal of the allyl protector is also effected by sodium iodide in hot acetone.[78] Protecting groups that are removed through a �-elimination mechanism are widely used. 2-Cyanoethyl and related protection can be deprotected by treatment with a base such as methanolic ammonia.[72] The 2,2,2-trichloroethyl or 2,2,2-tribromoethyl groups are protectors reductively removed by treatment with a Zn/Cu couple.[79,80] Here, the most widely used protective groups and their deprotections are noted. For more exotic examples see the books by Hayakawa[81] or Green.[54] In large molecules where alternative sites for chemical phosphorylation may exist, various protection and deprotection procedures have to be inserted in the scheme of synthesis. Enzymatic phosphorylations can make synthesis more efficient by eliminating many of these steps. In addition, enzyme-catalyzed introduction of phosphoryl groups can be diastereo-, enantio-, or regioselective. 2. Enzymatic phosphorylations towards phosphomonoesters As described above, the introduction of a phosphate moiety into a polyhydroxy compound by classic chemical methods is tedious since it usually requires a number of protection and deprotection steps of the substrate. Furthermore, the formation of oligophosphate esters as undesired by-products arising from over-phosphorylation is a common problem. Employing enzymes for the regioselective formation of monophosphate esters can eliminate many of these disadvantages. In addition, enzyme-catalyzed introduction of phosphoryl groups can be diastereo-[82] or enantioselective.[83,84] Isolated enzymes that form or cleave P-O bonds are important biocatalysts. Examples are restriction endonucleases, (deoxy)ribonucleases, DNA/RNA-ligases, DNA/RNA-polymerases, reverse transcriptases etc. that are central to modern molecular biology.[85] This part of the thesis gives an overview of research based on different phosphorylating and dephosphorylating enzymes useful in organic synthesis. Hundreds of enzymes potentially useful in synthesis are available in nature. Identifying enzymes useful in phosphorylations and dephosphorylations which have been ambiguously classified[86-88] is difficult for those not familiar with biochemistry. In general little information is available connecting enzymatic activity to synthetic applicability. Firstly, enzymes inserting and removing phosphoryl groups are spread almost over all classes. For example glyceraldehyde-3-phosphate dehydrogenase, which catalyses the oxidative phosphorylation of glyceraldehydes-3-phosphate to 1,3-diphosphoglycerate, is classified under E.C. 1.2.1.12 and 1.2.1.13. Neither the name of the enzyme nor its IUB-classification gives a clue that a phosphorylating step is involved. A second point is that many enzyme-catalyzed reactions are reversible. Some hydrolytic enzymes can be used in enzyme-catalyzed phosphorylations. Alkaline phosphatase for example, was used in the phosphorylation of glycerol with inorganic phosphate.[89,90] A third important point is to choose the right phosphate donor for the enzyme because not all phosphorylated

Chapter 1

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compounds can be used as donors. The free energy of hydrolysis of a phosphorus compound (�G�’hydro) is called its phosphorylating potential and is used to compare the ability of different compounds to effectively transfer a phosphoryl group. Table 1 summarizes the phosphorylating potentials of a number of important biological compounds having phosphoryl donor abilities. By far the most important phosphorylating agent in biological systems is adenosine 5’-triphosphate (ATP) used by kinases. Phosphorylation with low-potential phosphorylating agents are thermodynamically unfavourable. In biological systems, these processes are made possible by coupling them to a thermodynamically more favourable process.

Table 1. Standard free energies of hydrolysis for common metabolites[91] Metabolite �G°’hydro (kJ/mol) Phosphoenolpyruvate -62 Carbamoyl phosphate -51 1,3-Bisphosphoglycerate -49 Acetyl phosphate -43 Phosphocreatine -43 Pyrophosphate (PPi) -33 Phosphoarginine -32 ATP� AMP + PPi -32 Acetyl CoA -32 ATP � ADP + Pi -30

High-energy compounds

Glucose 1-phosphate -21 Glucose 6-phosphate -14 Glycerol 3-phosphate - 9 AMP � Adenosine + Pi - 3

Low-energy compounds

Phosphorylation by kinases In biological systems, phosphate esters are usually produced by phosphorylating enzymes belonging to the class of kinases, which catalyze the transfer of a phosphate moiety (or a di- or triphosphate moiety in certain cases) from ATP to a variety of alcohols[9,10] (Figure 2). Other nucleoside triphosphates have similar phosphorylating potentials but they are rarely used as phosphoryl group donors.[92,93] The kinases discussed below have found application in the synthesis of phosphorylated compounds. Hexokinase (E. C. 2.7.1.1) is an enzyme that is able to phosphorylate D-glucose in a one-step reaction to D-glucose-6-phosphate (G6P), a useful reagent for the regeneration of nicotinamide cofactors.[94,95] The enzyme has a broad substrate specificity since other hexoses and their thio- or aza-analogues are selectively phosphorylated on the primary alcohol moiety located at position 6 as well.[96,97] D-arabinose, a pentose is also a substrate for hexokinase.[98] Ribokinase (E. C. 2.7.1.17) can phosphorylate D-ribose to D-ribose-5-phosphate.[82] Glycerol kinase[99] (E. C. 2.7.1.30) is not only able to accept its natural substrate glycerol to form sn-glycerol-3-phosphate[100] or close analogues such as dihydroxyacetone,[84,101] but it is also able to transform a large variety of prochiral or racemic primary alcohols into chiral phosphates with enantiomeric excesses (ee) > 90-95% and yields of 75-95%.[83,102] Adenylate kinase (E. C. 2.7.4.3) has been used in the synthesis of several nucleoside phosphate analogs. For example, ribavarin triphosphate, a compound with anti-viral properties, was prepared from ribavarin monophosphate with adenylate kinase.[103] Other nucleotide analogues for example ATP-�-S and ATP-�-S have also been synthesized.[104,105] NAD kinase (E. C. 2.7.1.23) has been used in the conversion of nicotinamide adenine dinucleotide (NAD+) into its more expensive phosphate NADP+ with acetylphosphate for ATP regeneration.[106] It was possible to synthesize 8-azido-


17

[2’-32P]NADP(H) as a photoaffinity label for NADP(H)-specific enzymes using [�-32P]ATP.[107] A lot of other kinases exist (E. C. 2.7.1.-),[88] but they are very specific for their substrate and will not be discussed here.

Figure 2. Phosphorylation of alcohols by ATP consuming kinases and enzymatic ATP recycling systems. PEP = phosphoenolpyruvate; AcP = acetyl phosphate; MCP = methoxycarbonyl phosphate; CP = carbamoyl phosphate; PC = creatine phosphate. Enzymes used in the regeneration of ATP The addition of stoichiometric amounts of the cofactor ATP would not only be undesirable from a commercial standpoint[108] but also for thermodynamic reasons. Quite often the accumulation of the inactive form of the consumed cofactor can tip the equilibrium of the reaction in the reverse direction. Furthermore, product isolation would be more difficult. Thus, ATP must be used in catalytic amounts and continuously regenerated[101,109,110] during the course of the reaction by an auxiliary system which usually consists of a second enzyme and a stoichiometric quantity of an ultimate (cheap) phosphate donor as shown in Figure 2. These methods have in common that phosphoryl groups are transferred form a high-energy donor (cf. Table 1) to ADP. For most synthetic applications, either phosphoenolpyruvate (PEP)/pyruvate kinase (PK) or acetyl phosphate (AcP)/acetyl kinase (AcK) are used to regenerate ATP. The PEP/PK (E. C. 2.7.1.40) system is probably the most useful method for the regeneration of nucleoside triphosphates.[111] PEP is not only very stable towards hydrolysis but is also a strong phosphorylating agent. This makes PEP particularly convenient for use in slow and thermodynamically unfavourable phosphorylation reactions. The drawbacks of this system are the more complex synthesis of PEP[112,113] and the fact that PK is inhibited

Chapter 1

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by pyruvate at higher concentrations. So dilute reaction mixtures are used to keep the pyruvate concentration low. AcP/AcK (EC 2.7.2.1) is the most widely used large scale ATP regeneration system because of the ease of preparing AcP[114-117] However, because AcP is modestly stable in aqueous solutions its application is limited to fast phosphorylation reactions where the hydrolysis of AcP is not important. Furthermore, AcK is inhibited by acetate ions, the by-product of the reaction. Propionylphosphate has also been used to regenerate ATP by AcK, but is a poorer substrate than AcP.[117] As for pyruvate kinase, acetate kinase can accept nucleoside diphosphates other than adenosine diphosphate. Carbamoyl phosphate (CP)/carbamate kinase (CK; E.C. 2.7.2.2) is another ATP regeneration method.[118] However, CP spontaneously hydrolyzes in the aqueous reaction medium. Furthermore, the co-product carbamic acid spontaneously decarboxylates to form ammonia and carbon dioxide. Although this latter reaction would drive the phosphorylation to completion, the ammonium ions generated inhibit the kinase by displacing essential Mg2+-ions from the enzyme. Methoxycarbonyl phosphate (MCP) has also been proposed as substrate for acetate and carbamate kinase (but not for pyruvate kinase).[117] It is comparable to PEP in its high phosphorylating strength, but resembles acetyl phosphate in its ease of synthesis. The reaction product after phosphoryl transfer, methyl carbonate, hydrolyzes rapidly to form CO2 and MeOH. Due to the CO2 formation it is easy to drive the reaction to completion. Because of its short half-life (0.3 h, 25°C, pH 7), MCP is only used in a few cases where high phosphorylating potentials are required to push the phosphorylation reaction to the product side. Another very interesting but little-used regeneration method is based on phosphocreatine (PC) and creatine kinase (CrK; E. C. 2.7.3.2).[119] PC is comparable in its phosphorylating potential to AcP, but is more stable in aqueous solutions. CrK is inexpensive and fairly stable. The lack of an efficient and simple laboratory scale synthesis for PC has limited the application of this method to a few phosphorylations of sugars[119] and nucleosides.[120] A number of reactions which consume ATP generate AMP rather than ADP as a product. Still fewer produce adenosine.[121] A simple modification of the above-mentioned recycling systems for ATP from ADP makes the recycling from AMP feasible. The addition of the enzyme adenylate kinase (AdK; E. C. 2.7.4.3) catalyzes the phosphorylation of adenosine to give AMP, which in turn is further transformed to ADP by AdK. Both steps proceed with the consumption of ATP.[122] The above mentioned regeneration systems can be used to form ATP. Phosphorylation by enzymes using other phosphate donors than ATP Alkaline (E.C. 3.1.3.1) and acid phosphatases (EC 3.1.3.2), both non-specific, can be used in phosphorylations using other phosphoryl donors than ATP. Alkaline phosphatase from calf intestine was used in the enzyme-catalyzed phosphorylation of glycerol with inorganic phosphate (Pi) or pyrophosphate (PPi) as phosphate donors, with PPi being the better donor.[89,90] 75 g of glycerol-3-phosphate was isolated in a 41% yield using a 70% (v/v) glycerol solution with 800 mM phosphate donor. The reaction was regio- but not stereoselective. This enzyme was also able to phosphorylate some other simple alcohols, monosaccharides and polyols.[89] Acid phosphatases, especially those belonging to the group of bacterial non-specific acid phosphatases (NSAPs), have been used in phosphorylation reactions more frequently. For example inosine was phosphorylated to inosine monophosphate[123-127] using cheap PPi as phosphate donor. These enzymes were also able to regioselectively phosphorylate D-glucose to D-glucose-6-phosphate.[127,128] Furthermore other hexoses and pentoses, various


19

simple alcohols, among which dihydroxyacetone,[129] polyols and aromatic alcohols are accepted as substrates using PPi as phosphate donor.[128] The use of NSAPs in phosphorylation reactions is described in more detail below. Phosphate hydrolyzing enzymes Three groups of phosphate hydrolyzing enzymes can be distinghuished, alkaline phosphatases, acid phosphatases and the more substrate specific phosphohydrolases. Alkaline phosphatases are used as non-specific phosphatases in for example the hydrolysis of polyprenol phosphates like 6,7-epoxygeranyl diphosphate and 6,7-epoxy bis-homogeranyl diphosphate[130] and in sphingoside base 1-phosphate analysis in biological samples.[131] A regioselective dephosphorylation of 2’-carboxyl-D-arabinitol 1,5-bisphosphate was used in the synthesis of 2’-carboxy-D-arabinitol 1-phosphate, a natural inhibitor of ribulose 1,5-bisphosphate carboxylase.[132] The hydrolysis of the 5-phosphoryl group by alkaline phosphatase gave a 4:1 mixture of the 1- and 5-phosphate derivatives. On the other hand, hydrolysis with acid phosphatase was essentially quantitative yielding exclusively the 1-phosphate derivative. Alkaline phosphatases were also used in the hydrolysis of nucleotides,[133] and aromatic phosphate esters as potential chemiluminescent 1,2-dioxetane based compounds.[134,135] Acid phosphatases have found wider applications. For example, the product from a DHAP-dependent aldolase-catalyzed reaction is a labile 2-oxo-1,3,4-triol, which is phosphorylated at position 1. Dephosphorylation under mild conditions, without isolation of the intermediate phosphate species, by using acid phosphatases is a method frequently used to obtain the chiral polyol products.[136-139] Similarly, hydrolysis of polyprenyl pyrophosphates catalyzed by acid phosphatases readily afforded the corresponding dephosphorylated products in acceptable yields without the side reactions which occur during chemical hydrolysis.[140,141] The hydrolysis of carboxyl esters using lipases have found wide application in the kinetic resolution of chiral alcohols. In contrast to this, enantioselective hydrolyses of phosphate esters have been seldomly reported. As shown by Scollar et al.,[142] rac-threonine was resolved into its D- and L-enantiomers via hydrolysis of the O-phosphate esters using acid phosphatases. The application of acid phosphatases to resolve D-allo-threonine and D-threonine has been described by Kimura et al.[143] We have further explored the potential application of acid phosphatases in this reaction as outlined in chapters 5 and 6.[144] Inorganic pyrophosphate (PPi) may be considered as a particular case of a phosphate monoester. The enzymatic decomposition of PPi by inorganic pyrophosphatase (E. C. 3.6.1.1) can be used to drive a multi-enzyme synthesis towards uridine 5’-monophosphate (UMP).[82] The condensation of 5-phospho-D-ribulose-�-1-pyrophosphate to orotate by O-5-P-pyrophosphatase results in orotidine 5’-monophosphate (O-5-P) and PPi. To drive the reaction to completion the PPi is hydrolyzed by pyrophosphatase. Subsequent decarboxylation by O-5-P-decarboxylase results in UMP. A very good example of a specific enzymatic dephosphorylation is the hydrolysis of (±)-5’-phosphorylated aristeromycin by 5’-ribonucleotide phosphohydrolase (E. C. 3.1.3.5). The (-)-enantiomer of aristeromycin shows cytostatic and antiviral activity, while the (+)-enantiomer is inactive. The racemate (±)-5’-phosphorylated aristeromycin was resolved by selective hydrolysis of the (-)-enantiomer with the hydrolase.[145] The (-)-alcohol and the (+)-5’-phosphate derivative were separated easily on a silica gel column. Subsequent hydrolysis of the (+)-enantiomer with a non-specific alkaline phosphatase yielded pure (+)-alcohol. More phosphate monoester hydrolysing enzymes can be found in the E. C. 3.1.3.- class.[88]

Chapter 1

20

Mechanistic aspects of (enzymatic) P-O bond formations and cleavages have been recently reviewed[146,147] but are outside the scope of this work 3. Phosphatases Most dephosphorylations in vivo are catalyzed by a group of enzymes indicated as phosphatases (EC 3.1.-.-). These enzymes are believed to function essentially in scavenging organic phosphoesters, such as nucleotides, sugar phosphates, phytic acid etc., that cannot cross the cytoplasmic membrane. Inorganic phosphate (Pi) and organic by-products are released, that can be transported across the membrane, thus providing the cell with essential nutrients.[148] Some phosphatases have evolved specialized functions relevant to microbial virulence,[149,150] signal transduction,[151,152] energy conversion and metabolism.[153] Classification of phosphatases was initially based on the biochemical and biophysical properties such as pH optimum, substrate profile, sensitivity to known inhibitors and molecular size. As sequence data became available, it was recognized that phosphatases could be grouped into different molecular families according to similarity at the level of primary structure. Signature sequence patterns specific for each family have been identified.[154] The enzymes used for the work described in this thesis belong to the bacterial non-specific acid phosphatases[155] (NSAPs). These are non-metal soluble periplasmic proteins or membrane-bound lipoproteins. They are able to hydrolyze a broad range of structurally unrelated organic phosphomonoesters and are therefore called non-specific. The optimal pH for this class of enzymes is at acidic to neutral pH values. NSAPs are monomeric or oligomeric proteins containing peptide components with an Mr of 25-30 kDa. On basis of amino acid sequences, three different families of NSAPs were identified: molecular class A,[156] B[157] and C,[158] which are completely unrelated at the sequence level. The NSAPs used in this project belong to class A, therefore further discussion is focused on class A NSAPs. The class A NSAPs possesses a conserved sequence motif, K-(X6)-R-P-(X12–54)-P-S-G-H-(X31–54)-S-R-(X5)-H-(X2)-D.[155] The same domains are found in several lipid phosphatases, mammalian glucose-6-phosphatase and vanadium haloperoxidases.[159-

162] Table 2 shows the amino acid comparison of the three domains that are conserved in these enzymes. Apo-chloroperoxidase, from which the prosthetic group vanadate was removed possesses phosphatase activity, although the turnover with p-nitrophenylphosphate (pNPP) as a substrate is only 1.7 min-1.[159] Similarly, when the acid phosphatase is incubated with vanadate, it shows moderate bromoperoxidase activity.[164] The class A NSAPs are further classified into class A1, A2 and A3 depending on the amino acid sequences, substrate specificities and inhibition effects. The class A1 enzymes exhibit broad substrate specificity. They are able to hydrolyze 5´- and 3´- nucleotide monophosphates (NMPs), hexose and pentose phosphates and aryl phosphates, such as pNPP and phenolphthalein phosphate, but not diesters.[155] The Shigella flexneri[165] (PhoN-Sf) and Escherichia blattae[166] (EB-NSAP) proteins are class A1 NSAPs. The prototype of class A2 NSAPs is the non-specific acid phosphatase from Salmonella enterica ser. typhimurium (PhoN-Se, also indicated as non-specific acid phosphatase I).[167,168] It is active against a very broad array of substrates showing an even wider substrate specificity compared to that of class A1 enzymes. Shigella flexneri apyrase[169] (Apy-Sf) belongs to the class A3 group. Upon sequence comparison, in spite of functional dissimilarity with other NSAPs, it shows striking similarity with other class A enzymes.[155] The enzyme shows a distinctive activity on nucleoside triphosphates (NTPs), which are hydrolyzed to the corresponding nucleoside diphosphates (NDPs). The enzyme is active towards PPi, but has


21

low activity on pNPP. Because of its substrate specificity and its optimum pH (between 7 to 7.5), Apy-Sf can be considered as an ATP diphosphohydrolase or apyrase (EC 3.6.1.5.). Class A1 NSAPs show higher phosphatase activity on 5’-NMPs (primary alcohol) rather than 3’-NMPs (secondary alcohol) whereas class A2 NSAPs are able to hydrolyze both 5’- and 3’-NMPs. Class A3 NSAPs preferably catalyze the hydrolysis of NTPs, but they hardly hydrolyze NMPs. In 2000, X-ray structures of Escherichia blattae NSAP (EB-NSAP) co-crystallised with the transition-state analogues sulfate and molybdate were determined[166] (PDB codes 1EOI, 1D2T). The crystal structure of PhoN from Salmonella enterica ser. typhimurium MD 6001 (PDB code 2A96) has recently been elucidated.[170] A reaction mechanism has been proposed based on structural analysis, homology and mutational analysis.[159,162,163,166,171,172] The conserved active site residues participate in the binding of the phosphate, act as a nucleophile, stabilize the penta-coordinated transition state and play a role in leaving group protonation (Figure 3). Figure 3. Active site of the acid phosphatase from Salmonella enterica ser. typhimurium MD6001 co-crystallized with phosphate.[173,174] The substrate binding site is comprised of Lys 123, Arg 130, Ser 156, Gly 157, His 158, Arg 191 and His 197 residues. The side-chain atoms of Lys 123, Arg 130, Ser 156, Gly 157, and Arg 191 interact with the phosphate oxygen atoms keeping the phosphate group of the substrate close to His 197. The cleavage of the O-P bond of the phosphomonoester is believed to be facilitated in two steps by two histidyl residues. Firstly, deprotonated NE2 of His 197 carries out a nucleophilic attack at the electron deficient phosphorus center of the monophosphate, leading to the formation of a phosphoenzyme intermediate. The stabilization of the organic product is achieved by transfer of a proton from the proximal His 158. In the second step, deprotonated His 158, acting as a general base, activates a water molecule that attacks at the PO3 moiety of the phospho-histidine intermediate, leading to the release of Pi.

Arg 130 Lys 123

His 197

Arg 191

His 158

Gly 157

Ser 156

Arg 130 Lys 123

His 197

Arg 191

His 158

Gly 157

Ser 156

Chapter 1

22

Ref

eren

ces

[165

] [1

76]

[170

] [1

55]

[155

] d [1

55]

[166

] [1

56]

[177

] [1

55]

[169

] [1

68]

d [175

] [1

75]

[175

] [1

75]

[175

] [1

75]

[175

] [1

75]

[175

] [1

75]

[175

] [1

75]

[175

] [1

75]

[179

] [1

80]

[181

]

Domain I Domain II Domain III

133-KEYY-MRIRP-21-SYPSGHT-25-YELGDSRVICGYHWQSDV-212a

123-KKYY-MRTRP-21-SYPSGHT-25-WEFGQSRVICGAHWQSDV-202

123-KKYY-MRTRP-21-SYPSGHT-25-WEFGQSRVICGAHWQSDV-202

133-KIKY-MRIRP-21-SYPSGHT-25-YELGESRVICGYHWQSDV-212




133-KDHY-MRIRP-21-SYPSGHT-25-YELGQSRVICGYHWQSDV-212

133-KEHY-MRIRP-21-SYPSGHT-25-YQLGQSRVICGYHWQSDV-212

135-KDHY-MRVRP-21-SYPSGHT-25-YQMGQSRVICGYHWQSDV-214

133-KEKY-MRIRP-21-SYPSGHT-25-YELGQSRVICGYHWQSDV-212

124-KEYY-KRVRP-21-SYPSGHA-25-YEFGESRVICGAHWQSDV-203

132-KNNW-NRKRP-21-SYPSGHT-25-QIFGTSRIVCGAHWFSDV-211

83-KRIL-KIPRP-15-STPSGHS-48-LLVGFSRVYLGVHYPTDV-179

76-KWIL-FGQRP-29-GSPSGHA-43-LNVCLSRIYLAAHFPHQV-181



97-KDKV-QEPRP-52-AFPSGHT-30-TGVMGSRLLLGMHWPRDL-212

94-KALF-EEPRP-52-SFPSGHT-35-LLMLISRVRLGMHYPIDL-214

120-KYSI-GRLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228

120-KYSI-GRLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228

120-KYTI-GSLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228

136-KLII-GNLRP-39-STPSGHS-32-LVVNVSRVIDHRHHWYDV-240

117-KYMI-GRLRP-37-SFYSGHS-38-LYVGYTRVSDYKHHWSDV-225

128-KDYW-CLPRP-18-GAPSSHT-36-MTLVFGRIYCGMHGILDL-215

129-KDYW-CLPRP-18-GAPSSHS-36-LTLVFGRVYCGMHGMLDL-216

118-KNWI-GRLRP-37-TTPSGHS-40-ALIALSRTQDYRHHFVDV-228

149-KVSI-GRLRP-37-SFFSGHA-38-FYTGLSRVSDYKHHPSDV-232

353-KWEF-EFWRP-37-AYPSGHA-78-FENAISRIFLGVHWRFDA-501

341-KWQVHRFARP-62-SYPSGHA-54-VNVAFGRQMLGIHYRFDG-491

400-KFNIHRRLRP-72-SYGSGHA-52-DNIAIGRNMAGVHYFSDQ-558

KXXX-XXXRP XXXSGHX XXXXXXRXXXXXHXXXXX

Cla

ssifi

catio

n A

1 N

SAP

A2

NSA

P A

2 N

SAP

A1

NSA

P A

1 N

SAP

A1

NSA

P A

1 N

SAP

A1

NSA

P A

1 N

SAP

A3

NSA

P A

1 N

SAP

A3

NSA

P A

NSA

P N

eutra

l Pse

G

6Pas

e G

6Pas

e G

6Pas

e PG

Pase

B

PGPa

se B

PA

P2

PAP2

a LP

P-1

LPP-

1 LP

P-2

LBP-

/YSR

1 LB

P-2/

YSR

2.1

DG

PPas

e D

ri42

VC

PO

VB

PO

VB

PO

Acc

essi

on n

umbe

r D

8296

6 X

5903

6 Q

71EB

8 Q

9F1U

0 C

AB

5972

5 B

AB

1891

8 B

AB

1891

7 A

B02

0481

X

6444

4 A

B01

7537

X

6482

0 U

0453

9 M

2414

1 L2

5421

P3

5575

P3

5576

L3

7333

P1

8201

P4

4570

U

9055

6 A

B00

0888

D

8437

6 U

3305

7 A

F035

959

Z494

10

P235

01

U51

031

Y07

783

X85

369

P817

01

AF2

1881

0

Tab

le 2

. Am

ino

acid

sequ

ence

com

paris

on o

f thr

ee d

omai

ns in

pho

spha

tase

s and

van

dadi

um p

erox

idas

es.

Sour

ce

Shig

ella

flex

neri

(Pho

N-S

f)

Salm

onel

la e

nter

icab (P

hoN

-Se)

Sa

lmon

ella

ent

eric

ac K

lebs

iella

pla

ntic

ola

Kle

bsie

lla p

neum

onia

e Ra

oute

lla p

lant

icol

a En

tero

bact

er a

erog

enes

Es

cher

ichi

a bl

atta

e (E

B-N

SAP

Mor

gane

lla m

orga

nii (

PhoC

-Mm

) Pr

evot

ella

inte

rmed

ia (P

iAC

P)

Prov

iden

cia

stuar

tii (P

hoN

-Ps)

Sh

igel

la fl

exne

ri a

py (A

py-S

f)

Zym

omon

as m

obili

s (Ph

oC-Z

m)

Trep

onem

a de

ntic

ola

Hom

o sa

pien

s M

us m

uscu

lus

Rattu

s nor

vegi

cus

Esch

eric

hia

coli

Hae

mop

hilu

s inf

luen

zae

Rattu

s nor

vegi

cus

Hom

o sa

pien

s M

us m

uscu

lus

Sacc

haro

myc

es c

erev

isia

e H

omo

sapi

ens

Sacc

haro

myc

es c

erev

isia

e Sa

ccha

rom

yces

cer

evis

iae

Sacc

haro

myc

es c

erev

isia

e Ra

ttus n

orve

gicu

s C

urvu

lari

a in

aequ

alis

As

coph

yllu

m n

odos

um

Cor

allin

a of

ficin

alis

C

onse

nsus

a

Num

bers

sho

wn

at th

e ou

tsid

e of

dom

ains

I an

d II

I ref

er to

the

num

berin

g of

the

first

and

last

am

ino

acid

in th

ose

dom

ains

from

prim

ary

sequ

ence

, res

pect

ivel

y. T

he in

terv

enin

g nu

mbe

rs re

fer

to th

e nu

mbe

r of

am

ino

acid

s be

twee

n th

e do

mai

ns. B

old

lette

rs in

dica

te a

min

o ac

ids

that

are

con

serv

ed in

the

cons

ensu

s se

quen

ce. b s

er. t

yphi

mur

ium

LT

2. c s

er. t

yphi

mur

ium

MD

6001

=

Salm

onel

la e

nter

ica

ser.

typh

i (Q

934J

6). d S

eque

nces

list

ed w

ithou

t ref

eren

ce w

ere

depo

site

d di

rect

ly in

to d

atab

ases

. The

tabl

e w

as m

odifi

ed fr

om re

fere

nces

[155

] and

[175

]


23

Phosphorylation by Class A Bacterial Non-Specific Acid Phosphatases It is known already for some time that these NSAPs are able to carry out transphosphorylation reactions,[182] that is the transfer of phosphate from one molecule (donor phosphate e. g. phosphomonoesters or pyrophosphate (PPi)) to another different molecule (acceptor alcohol). Phosphorylation of alcoholic substrates is thought to be a two-step reaction (Scheme 1).[127] First the enzyme binds to phosphate donor to form a phosphoryl intermediate. In the second step the phosphoryl intermediate is either attacked by water (hydrolysis) or by an alcoholic acceptor resulting in phosphorylation. The Km-value for the alcohol, therefore, is a very important factor that determines whether an effective phosphorylation occurs. When the affinity for the substrate is low the phosphorylated intermediate prefers to react with water resulting in hydrolysis of the phosphate monoesters.

Scheme 1. Overall mechanism of phosphorylation and dephosphorylation catalyzed by acid phosphatases. The enzyme reacts with PPi to produce a binary PPi-enzyme complex (1). This complex dissociates (2) to yield an activated phosphorylated enzyme intermediate (E.Pi). A reaction (3) with water may occur resulting in dissociation of the intermediate as well as hydrolysis of PPi. The intermediate may also transfer (4) the phosphate to a bound acceptor (R-OH), which dissociates (5) to form a phosphomonoester and the free enzyme. Hydrolysis of phosphomonoesters proceeds also via the E.Pi intermediate. The group headed by Asano showed in pioneering studies that NSAPs transfer a phosphate group from PPi to nucleosides.[182] PPi is a safe compound which is also used as a food additive. It can be simply produced from phosphate at low costs.[183] However, PPi has a chelating effect and binds multivalent metals such as Ca2+, Mg2+, and Fe2+. Therefore PPi can not be used in combination with phosphatases that require metal ions[184] because PPi will inhibit the activity. Class A acid phosphatases do not require metal ions, and most of the enzymes in this class are able to hydrolyze PPi to form a phosphoryl intermediate that may react with a suitable alcoholic function. Nucleotides are often used as food additives and as pharmaceutical intermediates. Their biological activity is related to the position of the phosphate group. Inosine 5’-monophosphate (5’-IMP) or guanosine 5’-monophosphate (5’-GMP) are used as a flavour potentiator (umami) in various foods whereas the 2’-monophosphates are tasteless.[182] An enzymatic procedure based upon the use of inosine kinase from Escherichia coli as a phosphorylating enzyme is known.[185] The kinase requires ATP, which needs to be

Chapter 1

24

regenerated, making the process more complex. It is also possible to obtain 5’-nucleotides by a chemical method[28] but this is not acceptable due to toxicity and complexity because two reactors are needed, one for fermentation of inosine and one for the chemical phosphorylation. Asano et al. discovered that NSAPs are able to regioselectively phosphorylate nucleosides by using PPi as a phosphate donor. In particular the phosphorylation of inosine was studied.[182,186-188] The advantages of this new method are simplicity, low cost and mild reaction conditions. However, there were a number of problems to be solved. Firstly, the solubility of nucleosides is often below the Km-value of the enzyme.[187] Secondly, all of the synthesized 5’-NMP is rehydrolyzed to the nucleoside as the reaction time is prolonged and the PPi is consumed. To solve these problems random mutagenesis was carried out on the Morganella morganii acid phosphatase (PhoC-Mm) gene resulting in the variant I171T/G92D showing a significant enhancement of inosine phosphorylation.[187] Corresponding mutations into EB-NSAP also show the decrease in the Km-value for inosine, resulting in an increased yield of 5’-IMP.[189] The phosphorylation of inosine to 5’-IMP using PPi by PhoN-Sf and PhoN-Se was shown in our group by Tanaka et al.[127] PhoN-Sf catalyzes the phosphorylation of inosine to 5’-IMP, whereas PhoN-Se synthesizes both 5’-IMP and 3’-IMP. Since the acid phosphatases were able to (regioselectively) phosphorylate the ribose group in inosine, it came as no surprise that more simple compounds were phosphorylated as well by PhoN-Sf and PhoN-Se. Both phosphatases are able to phosphorylate D-glucose to D-glucose-6-phosphate using PPi as phosphate donor in a very efficient manner.[127,128] Several different compounds containing alcoholic functions were also phosphorylated showing the broad substrate specificity of the enzymes. Among these substrates was dihydroxyacetone which was phosphorylated by both PhoN-Sf and PhoN-Se to dihydroxyacetonephosphate (DHAP).[129] DHAP is an important compound which is used in aldol condensations using DHAP dependent aldolases, resulting in a C-C coupled product with two new stereocenters with high stereoselectivity.[6] It was shown by us that the in situ generated DHAP can be coupled to an aldehyde in an aldolase-catalyzed condensation using rabbit muscle aldolase (RAMA) in a one-pot cascade reaction.[129] Advantage is taken of the two-way action of the phosphatase. First it catalyzes the simple phosphorylation of DHA avoiding the problems with chemical phosphorylation and second, it dephosphorylates the aldol adduct, avoiding non-enzymatic dephosphorylation which, in general, may cause decomposition of the products. Further details are given in chapter 3 of this thesis. Dephosphorylation by class A bacterial non-specific acid phosphatases Enzymes are known for their possible enantioselectivity and lipases are a well known example.[190] NSAP’s have not been used before as a tool for deracemization of racemic mixtures. However, the acid phosphatase from wheat germ, belonging to another enzyme class, was able to discriminate between D- and L-O-phosphothreonine.[142] It was shown by us that PhoN-Se was also able to discriminate between O-phospho-D- and L-threonine in a highly selective manner.[144] In contrast to the high selectivity with O-phospho-threonine, PhoN-Se was not able to resolve O-phospho-DL-serine. However, by random mutagenesis and screening, a mutant was selected that showed an E-value of 18.1 compared to 3.4 for the wild type enzyme.[144] Not only stereoselectivity is a major issue in enzyme catalysis, but the separation of geometric isomers by enzymes is also possible. Geometric isomers such as 2-methylcyclohexanol have been subjected to separation by esterases showing preference for the trans-isomers but practical separation of the geometric (cis/trans) isomers has not been achieved[191,192] in contrast to separation of the stereoisomers. Separation of the geometric isomers has been achieved with wheat germ acid phosphatase


25

as shown by Klibanov et al.[193] This phosphatase is completely selective towards the trans-isomer. PhoN-Sf is shown to have a two times higher preference for the cis-isomer compared to the trans-isomer (this thesis, chapter 5). NSAPs in other functions than phosphorylation and dephosphorylation The class A NSAPs show similarity in the active site with vanadium haloperoxidases.[159] When vanadate is bound to the active site of the acid phosphatase, it shows vanadium haloperoxidase activity and it also performs enantioselective sulfoxidations. The WT PhoN-Se catalyzes the sulfoxidation of thioanisole towards the (S)-enantiomer with a selectivity of 36%.[164] Futhermore, NSAPs can be used in the bioremediation of heavy-metal waste.[194,195] The Pi released in the periplasmic space during hydrolysis of phosphomonoesters by NSAP’s promotes precipitation of heavy metal ions. The efficiency of these systems is greater than that of traditional ion-exchange sorption/desorption processes. 4. Enzyme engineering in biocatalysis Biocatalysis is now an important tool in the (industrial) synthesis of bulk chemicals, pharmaceuticals, agrochemicals and food ingredients.[196-198] Despite the successful development of biocatalysts for a variety of important transformations, industry demands different properties from enzymes than nature does. In nature most reactions occur at moderate temperatures in aqueous media, while an enzyme in an industrial process usually needs to be as stable as possible in an environment of higher temperatures, high substrate concentrations, sheering forces and organic solvents. Furthermore, low specific activity, inadequate substrate scope, and low or undesired enantiomer selectivity have limited the number and diversity of industrial enzyme applications. In cases where a potential biocatalytic route is not yet efficient enough, process engineering could be used for improvements.[199] Many different strategies are used including changing substrate properties, solvent engineering (e.g. polarity, hydrophobicity, ionic strenght), changing reaction conditions (e.g. temperature, pH, pressure) or using immobilized enzymes. All these approaches have in common that the biocatalyst itself is not modified. These modifications result in altered enzyme-substrate interactions, which could sometimes also alter the active-site geometry of the enzyme. By contrast, genetic engineering methods directly alter at least the primary structure of an enzyme, and often alter the secondary and tertiary structure as well. The improvement of enzyme performance by mutagenesis can be done by rational and random techniques. Rational redesign[200] of proteins involves the introduction of mutations at specific places by site-directed mutagenesis. The targets for such mutations are chosen based on knowledge of crystal structure and enzyme mechanism and are mostly focused on amino acids close to the active site keeping Emil Fischer’s principle of ‘lock and key’ or Koshland’s improved model based on ‘induced fit’ in mind. Rational redesign has its problems, most notably the amount of data that has to be accumulated on each enzyme under study, because even now, our understanding of the relationship between enzyme structure and function is limited. In addition, the prediction of the effect of mutations is complicated by the growing realization that enzyme molecules exist in solution as a mixture of structural conformers, and that dynamics play an important role in enzyme function. Many enzymes are not crystallized yet, which makes it difficult to use rational design. Fortunately, this gap is filled by random mutagenesis techniques[201,202] such as error prone polymerase chain reaction[203] (epPCR) where knowledge of the 3D-structure or mechanism

Chapter 1

26

of the enzyme is not required. In nature, evolution and creation of new functionalities is achieved by mutagenesis, recombination and survival of the fittest. This process can be mimicked in the laboratory where it is called directed evolution and follows iterative cycles of producing mutants through random mutagenesis and (high-throughput) screening or selection of the mutant with the desired properties (Figure 4). The extent to which directed evolution succeeds depends critically on the delicate interplay between the quality of biological diversity present in the library, the size of the library, and the ability of an assay to meaningfully detect improvements in the desired activity. Directed evolution in artificial conditions may not necessarily result in the desired enzyme for a real biocatalytic process, as it is well known that “you get what you screen for”. Creating biological diversity is very important in directed evolution. epPCR typically generates only one base-pair substitution per codon, so instead of generating 63 new codons encoding the full range of amino acids, only 9 new codons encoding 5 to 6 amino acids in average are generated.[204,205] Due to this degeneracy of the genetic code, some positive amino acid changes or important amino acid residues will be missed. By using a combination of mutagenesis techniques more diversity can be introduced. Important positions that are found after screening of an epPCR generated library can be subjected to site-saturation mutagenesis[206-208] in order to reveal which amino acid is optimal for that position. The size of a library depends on the average number of mutations per gene that are introduced. Only a small number of random mutations can be made at a time, as each new mutation typically inactivates between 30 and 40% of the remaining active proteins.[209] A low mutational rate (one to three mutations per gene, or in case of a large gene per 1000 bps) results in many functional sequences, but only a small number are unique. By contrast, very high mutation rates (15 to 30 mutations per gene) produce mostly unique sequences, but few retain function.[210] The success of a directed evolution experiment highly depends on the method that is used to select the best mutant enzyme from a large mutant library in which usually only a small percentage shows the desired properties. The big challenge is making the improved function quantifiable. Enzymatic assays have to be sufficiently sensitive and specific to identify positive mutants. Therefore, many studies have been devoted to the development of automated high-throughput screening methods[211] that make it possible to test thousands of variants per day. Directed evolution methods like random mutagenesis have been used to improve the biocatalytic performance of various enzymes. Especially substrate specificity, catalytic activity, thermal and oxidative stability, solvent tolerance, pH optimum, substrate/product inhibition and enantioselectivity have been successfully improved with directed evolution (as reviewed by [205,212-215]). These properties are often difficult to improve by rational design.[199] Furthermore, it is usually possible to screen a random library for multiple properties simultaneously, such as improved thermostability, solvent tolerance and activity at room temperature.[216] When using rational design, each enzyme characteristic that requires improvement is dealt with separately. What took nature millions of years to develop for a specific purpose can now in principle be performed in the test tube within weeks or months, namely the creation of an optimal catalyst for a reaction of interest to the organic chemist. However, to achieve this a number of stringent conditions have to be met. A laboratory that is equipped for doing molecular biology experiments meeting the safety requirements is necessary. A recombinant enzyme should be present that is easily expressed in a convenient host. Furthermore molecular biology and enzymology expertise should be available because an ensemble of technologies for first constructing a diversity of mutant genes and then sorting them based on their


27

corresponding phenotype has to be carried out. Mutations associated with improvements are a rare event, and detecting these positive mutants is not an easy task since the protein expression rate may vary from mutant to mutant in a given library, leading to different amounts of enzyme from well to well of microtiter plates.

Figure 4. The directed evolution process. The starting point is a wild-type (WT) enzyme which catalyzes a given reaction of interest but with suboptimal conversion. The gene that encodes for the WT enzyme is first subjected to random mutagenesis using e.g. epPCR, cassette mutagenesis and/or DNA shuffling to create a library of mutated genes. In each cycle the gene-library is first inserted in a standard bacterial host such as E. coli. Then bacterial colonies are plated on agar plates and harvested individually by a colony picker. Each colony is placed in a separate well of a microtiter plate containing nutrient broth, so that the bacteria grow and produce the protein of interest. Each colony originates from a single cell and thus produces only one mutant enzyme (provided that there is no undesired cross-contamination). A portion of each mutant is then robotically placed on a different microtiter plate, where the reaction of interest is carried out. Because the enzyme variants and the corresponding mutant genes are spatially addressable, the genotype/phenotype relation is maintained, and tedious deconvolution is not necessary. The best mutant (positive hit) is then the starting point for the next cycle of gene mutagenesis, expression, and screening. Because the inferior mutants are discarded, an evolutionary character of the overall process is simulated, leading to the formation and identification of a better enzyme. Since the process can be repeated as often as needed, a type of ‘Darwinistic’ principle holds.

Chapter 1

28

5. Enantioselective biocatalysis The demand for enantiopure chemicals has increased considerably in recent years. Many pharmaceuticals, but also food additives, fragrances and agrochemicals are nowadays applied as enantiopure products. One enantiomeric form may have the required effect whereas the other is ineffective and even may be toxic. The FDA has become increasingly reluctant to permit the introduction of racemic drugs, as these drugs are by definition saddled with 50% of chemical ballast.[217] To meet the growing demand for enantiopure chemicals, a significant amount of research has been devoted to the development of methods for producing optically pure intermediates and building blocks.[218-220] One of the most useful and practical ways to prepare compounds of high optical purity is catalysis by enantioselective enzymes.[221-223] Enzymes are renewable environmentally benign catalysts, whereas many chemical catalysts contain toxic and expensive heavy metals that are difficult to remove from the product. Enzyme catalysis can be carried out at ambient temperature and atmospheric pressure, avoiding the use of more extreme conditions, which could cause problems with isomerization, racemization, epimerization and rearrangement. This can be very important for large-scale applications in which energy costs of the process are also a factor. Enantiopure compounds can be produced by enantioselective catalysis in three different ways. In a kinetic resolution,[224,225] only one of the enantiomers of a racemic mixture (SR + SS) is converted into product (PR), leaving the other enantiomer behind in optically pure form (SS). In an enantioconvergent reaction,[226] both enantiomers of a racemic mixture (SR + SS) are converted to a single optically pure product (PR) making use of two independent reactions. In an asymmetric synthesis reaction,[227] a prochiral substrate (S) is converted to an optically enriched product (PR). Kinetic resolution of a racemic starting compound is one of the most commonly used techniques to obtain enantiopure chemicals.[224,225] In an ideal case, a catalyst converts only one of the two enantiomers, yielding both substrate and product in optically pure form with a yield of 50%, which is the theoretical maximum. The enantioselectivity of a reaction can be described by the enantiomeric ratio or E-value. The E-value is the ratio between the initial rates towards the different enantiomers at equal substrate concentrations. For an enzyme-catalyzed kinetic resolution, the E-value is an intrinsic parameter of the enzyme that describes the ratio between the specificity constants of both enantiomers (equation 1).[228]

][S][SE

][S)/K(k][S)/K(k

vv

S

R

S

RR

S

R

smcat

mcat�� (1)

with S

R

)/K(k)/K(kE

mcat

mcat�

The E-value can be calculated from a kinetic resolution experiment from the degree of conversion (c) and the enantiomeric excess of the substrate (eeS) or product (eeP) at different time points (equation 2). The E-value can be used to calculate the yield (1-c) of the remaining substrate at a certain ee and the ee of the product at that point.


29

)]eec(1ln[1)]eec(1ln[1

]eec)(1-ln[1]ee-c)(1-ln[1E

P

P

S

S

��

��

� (2)

with The major drawback of a kinetic resolution is the fact that the maximum yield of product and that of the remaining substrate can never exceed 50%. This problem can be circumvented by using a dynamic kinetic resolution,[229,230] in which constant in situ racemization[231] of the remaining enantiomer causes a racemic mixture to be converted to a single product enantiomer with a theoretically 100% maximum yield. Chapter 5 and 6 of this thesis describe the kinetic resolution of O-phospho-DL-threonine and -serine by the acid phosphatase from Salmonella enterica ser. typhimurium LT2.[144]

6. Outline of This Thesis The object of the research described in this thesis was to explore the possibility to use acid phosphatases in organic synthesis. The project was carried out using the recombinant non-specific acid phosphatases from Shigella flexneri (PhoN-Sf) and Salmonella enterica ser. typhimurium (PhoN-Se). In Chapter 2, the regioselectivity of the phosphatases in phosphorylation of various alcoholic compounds was investigated using PPi as a donor. It was shown that the enzymes phosphorylate a wide range of alcoholic substrates and the broad substrate specificity of PhoN-Sf was demonstrated. In Chapter 3, the phosphorylation of dihydroxyacetone (DHA) to dihydroxyacetone phosphate (DHAP) by the NSAPs was studied. DHAP is a useful compound for the enzymatic preparation of a variety of sugars by DHAP dependent aldolases. Phosphorylation was demonstrated, as well as the coupling of DHAP to an aldehyde via an aldolase mediated reaction and subsequent hydrolysis of the sugarphosphate to the final dephosphorylated product. It was also shown that the phosphate cycles through several rounds of DHA phosphorylation, thereby increasing the efficiency of the reaction. In Chapter 4, directed evolution is used to optimize the phosphorylation of DHA by PhoN-Se. Several variants were shown to be more efficient in the phosphorylation of DHA by PPi. The stereoselectivity of the NSAPs has been described in Chapter 5. Dephosphorylation of O-phospho-DL-threonine by PhoN-Se was shown to proceed in a stereoselective way resulting in L-threonine. PhoN-Se dephosphorylates O-phospho-DL-serine with a minor selectivity towards D-serine which is not sufficient for practical purposes. Chapter 6 describes a directed evolution method to optimize the stereoselectivity of the hydrolysis of O-phospho-DL-serine. Two variants were shown to be more stereoselective than the wild-type enzyme from PhoN-Se. These variants showed mutations close to the active site. References 1. F. H. Westheimer, Science 1987, 235, 1173-1178. 2. M. J. Berridge, R. F. Irvine, Nature 1984, 312, 315-321. 3. P. P. Dzeja, A. Terzic, J. Exp. Biol. 2003, 206, 2039-2047. 4. W. Kroutil, H. Mang, K. Edegger, K. Faber, Adv. Synth. Catal. 2004, 346, 125-142.

RS

RSS

SSS-See�

�SR

SRP

PPP-Pee�

�0S0S

SS-1cSR

R s

��

�

Chapter 1

30

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31

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