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Bacterial class A acid phosphatases as versatile tools in organic synthesis
van Herk, T.
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van Herk, T. (2008). Bacterial class A acid phosphatases as versatile tools in organic synthesis
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Download date: 14 Jun 2017
Chapter 1
General introduction
Chapter 1
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.
10
General introduction
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’-Oisopropylidene 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 syndialkyl 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
11
Chapter 1
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]
12
General introduction
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
13
Chapter 1
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. Hphosphonate 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 bistrimethylsilyl 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-2aldoxymate is achievable at room temperature and no side reactions occur.[61-63] The 8quinolyl 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,
14
General introduction
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 2,2,2tribromoethyl 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 enzymecatalyzed reactions are reversible. Some hydrolytic enzymes can be used in enzymecatalyzed 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
15
Chapter 1
compounds can be used as donors. The free energy of hydrolysis of a phosphorus
compound ('Gq’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
High-energy compounds
Pyrophosphate (PPi)
-33
Phosphoarginine
-32
ATPÆ AMP + PPi
-32
Acetyl CoA
-32
ATP Æ ADP + Pi
-30
Glucose 1-phosphate
-21
Glucose 6-phosphate
-14
Low-energy compounds
Glycerol 3-phosphate
- 9
AMP Æ Adenosine + Pi
- 3
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 dior 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 onestep 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-5phosphate.[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-azido16
General introduction
[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
17
Chapter 1
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 byproduct 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 regiobut 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 Dglucose to D-glucose-6-phosphate.[127,128] Furthermore other hexoses and pentoses, various
18
General introduction
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 bishomogeranyl diphosphate[130] and in sphingoside base 1-phosphate analysis in biological
samples.[131] A regioselective dephosphorylation of 2’-carboxyl-D-arabinitol 1,5bisphosphate 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 DHAPdependent 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-allothreonine 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 O5-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]
19
Chapter 1
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.[159162]
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] (PhoNSf) 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
20
General introduction
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).
His 158
His 158
Gly 157
Gly 157
Arg 191
Arg 191
Ser 156
Ser 156
His 197
His 197
Arg 130
Lys 123
Arg 130
Lys 123
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.
21
22
Table 2. Amino acid sequence comparison of three domains in phosphatases and vandadium peroxidases.
Domain I
Domain II
Domain III
Source
Accession number
Classification
References
133-KEYY-MRIRP-21-SYPSGHT-25-YELGDSRVICGYHWQSDV-212a
Shigella flexneri (PhoN-Sf)
D82966
A1 NSAP
[165]
123-KKYY-MRTRP-21-SYPSGHT-25-WEFGQSRVICGAHWQSDV-202
X59036
A2 NSAP
[176]
Salmonella entericab (PhoN-Se)
123-KKYY-MRTRP-21-SYPSGHT-25-WEFGQSRVICGAHWQSDV-202
Q71EB8
A2 NSAP
[170]
Salmonella entericac
133-KIKY-MRIRP-21-SYPSGHT-25-YELGESRVICGYHWQSDV-212
Q9F1U0
A1 NSAP
[155]
Klebsiella planticola
133-KIKY-MRIRP-21-SYPSGHT-25-YELGESRVICGYHWQSDV-212
CAB59725
A1 NSAP
[155]
Klebsiella pneumoniae
d
133-KIKY-MRIRP-21-SYPSGHT-25-YELGESRVICGYHWQSDV-212
BAB18918
A1 NSAP
Raoutella planticola
133-KIKY-MRIRP-21-SYPSGHT-25-YELGESRVICGYHWQSDV-212
BAB18917
A1 NSAP
[155]
Enterobacter aerogenes
133-KDHY-MRIRP-21-SYPSGHT-25-YELGQSRVICGYHWQSDV-212
[166]
AB020481
A1 NSAP
Escherichia blattae (EB-NSAP
133-KEHY-MRIRP-21-SYPSGHT-25-YQLGQSRVICGYHWQSDV-212
[156]
X64444
A1 NSAP
Morganella morganii (PhoC-Mm)
135-KDHY-MRVRP-21-SYPSGHT-25-YQMGQSRVICGYHWQSDV-214
AB017537
A3 NSAP
[177]
Prevotella intermedia (PiACP)
133-KEKY-MRIRP-21-SYPSGHT-25-YELGQSRVICGYHWQSDV-212
[155]
Providencia stuartii (PhoN-Ps)
X64820
A1 NSAP
124-KEYY-KRVRP-21-SYPSGHA-25-YEFGESRVICGAHWQSDV-203
[169]
Shigella flexneri apy (Apy-Sf)
U04539
A3 NSAP
132-KNNW-NRKRP-21-SYPSGHT-25-QIFGTSRIVCGAHWFSDV-211
M24141
A NSAP
[168]
Zymomonas mobilis (PhoC-Zm)
d
83-KRIL-KIPRP-15-STPSGHS-48-LLVGFSRVYLGVHYPTDV-179
Treponema denticola
Neutral Pse
L25421
76-KWIL-FGQRP-29-GSPSGHA-43-LNVCLSRIYLAAHFPHQV-181
Homo sapiens
G6Pase
[175]
P35575
76-KWIL-FGQRP-29-GSPSGHA-43-LNVCLSRIYLAAHFPHQV-181
Mus musculus
G6Pase
[175]
P35576
72-KWIL-FGQRP-29-GSPSGHA-43-LNVCLSRIYLAAHFPHQV-177
L37333
Rattus norvegicus
G6Pase
[175]
97-KDKV-QEPRP-52-AFPSGHT-30-TGVMGSRLLLGMHWPRDL-212
PGPase B
[175]
Escherichia coli
P18201
94-KALF-EEPRP-52-SFPSGHT-35-LLMLISRVRLGMHYPIDL-214
PGPase B
[175]
Haemophilus influenzae
P44570
120-KYSI-GRLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228
Rattus norvegicus
PAP2
[175]
U90556
120-KYSI-GRLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228
Homo sapiens
PAP2a
[175]
AB000888
120-KYTI-GSLRP-37-SFYSGHS-38-IYVGLSRVSDYKHHWSDV-228
Mus musculus
LPP-1
[175]
D84376
136-KLII-GNLRP-39-STPSGHS-32-LVVNVSRVIDHRHHWYDV-240
Saccharomyces cerevisiae
LPP-1
[175]
U33057
117-KYMI-GRLRP-37-SFYSGHS-38-LYVGYTRVSDYKHHWSDV-225
Homo sapiens
LPP-2
[175]
AF035959
128-KDYW-CLPRP-18-GAPSSHT-36-MTLVFGRIYCGMHGILDL-215
Saccharomyces cerevisiae
LBP-/YSR1
[175]
Z49410
Saccharomyces cerevisiae
LBP-2/YSR2.1 129-KDYW-CLPRP-18-GAPSSHS-36-LTLVFGRVYCGMHGMLDL-216
[175]
P23501
118-KNWI-GRLRP-37-TTPSGHS-40-ALIALSRTQDYRHHFVDV-228
Saccharomyces cerevisiae
DGPPase
[175]
U51031
149-KVSI-GRLRP-37-SFFSGHA-38-FYTGLSRVSDYKHHPSDV-232
Rattus norvegicus
Dri42
[175]
Y07783
353-KWEF-EFWRP-37-AYPSGHA-78-FENAISRIFLGVHWRFDA-501
Curvularia inaequalis
VCPO
[179]
X85369
341-KWQVHRFARP-62-SYPSGHA-54-VNVAFGRQMLGIHYRFDG-491
Ascophyllum nodosum
VBPO
[180]
P81701
400-KFNIHRRLRP-72-SYGSGHA-52-DNIAIGRNMAGVHYFSDQ-558
Corallina officinalis
VBPO
[181]
AF218810
KXXX-XXXRP
XXXSGHX
XXXXXXRXXXXXHXXXXX
Consensus
a
Numbers shown at the outside of domains I and III refer to the numbering of the first and last amino acid in those domains from primary sequence, respectively. The intervening numbers refer
to the number of amino acids between the domains. Bold letters indicate amino acids that are conserved in the consensus sequence. b ser. typhimurium LT2. c ser. typhimurium MD6001 =
Salmonella enterica ser. typhi (Q934J6). d Sequences listed without reference were deposited directly into databases. The table was modified from references [155] and [175]
Chapter 1
General introduction
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 twostep 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 Kmvalue 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
23
Chapter 1
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 Dglucose-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
24
General introduction
as shown by Klibanov et al.[193] This phosphatase is completely selective towards the transisomer. 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 PhoNSe 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
25
Chapter 1
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
26
General introduction
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.
27
Chapter 1
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]
vR
vS
with
E
(kcat/Km)R[SR]
(kcat/Km)S[Ss]
E
[SR]
[SS]
(1)
(kcat/Km)R
(kcat/Km)S
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.
28
General introduction
E
with
ln[1 - c)(1 - eeS]
ln[1 - c)(1 eeS]
c 1-
SR Ss
S R0 S S0
eeS
ln[1 c(1 eeP)]
ln[1 c(1 eeP)]
SS - SR
SS SR
eeP
(2)
PR - PS
PR PS
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 nonspecific 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 PhoNSe. 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.
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