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Phosphorylation by alkaline phosphatase: use of the
enzyme in cascade reactions
Joana Lisboa Vendrell Marques Peralta
Dissertação para obtenção do Grau de Mestre em
Engenharia Química
Júri
Presidente: Prof. José Manuel Madeira Lopes (DEQB, IST)
Orientadores: Prof. Maria Raquel Aires Barros (DEQB, IST)
Prof. Ron Wever (HIMS, UvA)
Vogais: Dr. Pedro de Barros Fernandes (CEQB, IST)
Outubro 2010
i
Acknowledgments I would like to thank my supervisor, Lara Babich, for teaching, supporting and helping me
through the course of this project, as well as the motivation and personal support.
I would also like to thank Professor Ron Wever for the opportunity to accomplish such
gratifying work in a great research group, for the stimulating discussions, helpful suggestions and
the interest demonstrated in the course of my work.
Finally, I would like to thank everyone at the Biocatalysis group at the University of
Amsterdam for the excellent working atmosphere. Not only was their advice and help essential
during the course of the project – a special word to Louis Hartog, Aleksandra Bury and Michael
van der Horst – but I will also keep everything I have learned during the informal meetings, tea
breaks and talks.
ii
Resumo
Visando obter um processo viável para sintetizar hidrocarbonetos não-naturais foram
desenvolvidas duas reacções enzimáticas em cascata; uma partindo de dihidroxiacetona (DHA)
e outra de glicerol. Em ambas as reacções ocorrem etapas de fosforilação e defosforilação,
catalisadas por uma fosfatase ácida. O objectivo deste projecto é verificar se a fosfatase alcalina
(AP,E.C.3.1.3.1) a pode substituir, visto que esta enzima também catalisa reacções de
transferência de fosfato e é de fácil obtenção.
Nestas reacções em cascata dihidroxiacetona fosfato (DHAP), produzida por fosforilação
de DHA ou glicerol (posteriormente oxidado pela catalase), é acoplada a um aldeido por uma
aldolase e o produto fosforilado é defosforilado pela fosfatase já presente na reacção.
A fosforilação da DHA e do glicerol pela AP e o seu uso nas reacções em cascata foi
optimizado. Na fosforilação da DHA obteve-se 2.6 mM de DHAP e na do glicerol 2.2 mM de
glicerofosfato (resultados comparáveis aos obtidos usando fosfatase ácida, PhoN.-Sf), a pH 8 e
30°C. A reacção em cascata partindo de DHA conduziu a 7% de conversão (pH 8, 30°C),
resultado pobre comparado à conversão de 60% obtida usando PhoN-Sf.
AP foi imobilizada em três suportes diferentes – Immobeads 150, Sepabeads EC-EP e
Sepabeads EC-HÁ, para preservar a actividade catalítica e permitir a sua reutilização. Após 4
horas de incubação AP encontra-se totalmente ligada covalentemente a Sepabeads EC-HA,
enquanto para os restantes suportes não ocorre ligação significativa.
As reacções em cascata realizadas usando AP imobilizada resultaram numa conversão
de 35% para a via da DHA e 30% para a via do glicerol (pH 8, 25°C).
.
Palavras chave: reacções enzimáticas em cascata; fosfatase alcalina; fosforilação de DHA e
glicerol; imobilização enzimática.
iii
Abstract
Aiming for a viable process to synthesise non-natural hydrocarbons two enzymatic
cascade reactions were developed in Amsterdam; one starting from dihydroxyacetone (DHA) and
another from glycerol. In both cascades phosphorylation and dephosphorylation steps are
involved, being catalysed by an acid phosphatase. The projects purpose was to investigate
whether it could be substituted by alkaline phosphatase (AP, E.C.3.1.3.1), since this enzyme also
catalyses phosphate transfer reactions and is easily available.
In these cascades dihydroxyacetone phosphate (DHAP), produced by phosphorylation of
DHA or glycerol (subsequently oxidized by catalase), is coupled to an aldehyde by an aldolase
and the phosphorylated aldol product is dephosphorylated by the already present phosphatase.
The phosphorylation of DHA and glycerol using AP and its use in the cascade reactions
was optimized. The DHA phosphorylation led to 2.6 mM of DHAP and the glycerol
phosphorylation to 2.2 mM of glycerophosphate, at pH 8 and 30°C (results comparable to those
obtained using the acid phosphatase, PhoN-Sf). The cascade reaction starting from DHA led to a
7% conversion (pH 8, 30°C), very low results compared to the 60% conversion obtained using
PhoN-Sf.
AP was immobilized on three different supports – Immobeads 150, Sepabeads EC-EP
and Sepabeads EC-HA, to preserve the catalytic activity and allow its repeated use. After four
hours of incubation AP was completely bound to Sepabeads EC-HA, whereas the other supports
didn‟t lead to a significant binding.
The cascade reaction using immobilized AP gave a 35% conversion for the DHA route
and 30% conversion for the glycerol route (pH 8, 25°C).
Key words: enzymatic cascade reactions; alkaline phosphatase; DHA and glycerol
phosphorylation; enzyme immobilization.
iv
Table of Contents
1. Introduction ........................................................................................................... 1
1.1 Motivation and background .................................................................................... 1
1.1.1 Carbohydrate chemistry ................................................................................................. 1
1.1.2. Enzymatic Synthesis of Carbohydrates - Aldolases ..................................................... 3
1.1.3. The IBOS Project .......................................................................................................... 5
1.1.4. The Alkaline phosphatase approach ............................................................................ 9
1.2. Alkaline Phosphatase ......................................................................................... 10
1.2.1. Origin and distribution of AP ....................................................................................... 10
1.2.2. The role of AP in nature .............................................................................................. 12
1.2.3 Protein structure and mechanistic issues .................................................................... 13
1.2.4. Applications of AP ....................................................................................................... 15
1.3. Enzyme immobilization ....................................................................................... 16
1.4. Aim of this Study ................................................................................................. 22
2. Materials and Methods ............................................................................................ 23
2.1. Alkaline Phosphatase ......................................................................................... 23
2.1.1. Calf Intestine Alkaline Phosphatase ........................................................................... 23
2.1.2. Immobilization supports (epoxy and amino groups) ................................................... 23
2.1.3. Reagents ..................................................................................................................... 23
2.1.4. Equipment ................................................................................................................... 25
2.1.5. Analytical Techniques ................................................................................................. 27
2.2. Production and purification of recombinant acid phosphatase from Salmonella
enterica (PhoN-Se) expressed in E. coli BL21 (DE3) with pET23b plasmid ............... 39
2.2.1. Protein expression overview ....................................................................................... 39
2.2.2. Solutions ..................................................................................................................... 41
2.2.3. Protocol for expression and purification...................................................................... 41
2.2.4. Analytical techniques .................................................................................................. 45
3. Results and Discussion .......................................................................................... 47
3.1. Alkaline Phosphatase ......................................................................................... 47
3.1.1. Characterization of the alkaline phosphatase stock solution ...................................... 47
3.1.2. Analytical techniques .................................................................................................. 48
v
3.2. Recombinant acid phosphatase from Salmonella enterica: V78L, V78Y and V78H
.................................................................................................................................. 62
3.2.1. Expression and purification of V78L, V78Y and V78H ............................................... 63
3.2.2. Cascade reaction ........................................................................................................ 64
4. Conclusion ............................................................................................................... 67
4.1. Alkaline Phosphatase ......................................................................................... 67
4.2. Recombinant acid phosphatase from Salmonella enterica: V78L, V78Y and V78H
.................................................................................................................................. 68
4.3. Future prospects ................................................................................................. 69
5. References ............................................................................................................... 71
Appendix A – PPi inhibition ........................................................................................ 78
Appendix B – HPLC chromatogram of DHA cascade using immobilized AP (1st
cycle) ............................................................................................................................ 80
Appendix C – DHA cascade using immobilized AP (2nd cycle) ................................ 82
vi
Abbreviations
AP Alkaline phosphatase
APAAP Alkaline Phosphatase Monoclonal Anti-alkaline Phosphatase
DEA Diethanolamine
DHA Dihydroxyacetone
DHAP Dihydroxyacetone phosphate
ECAP E. coli Alkaline Phosphatase
EDTA Ethylenediamine tetra acetic acid
FDP Fructose 1,6-diphosphate
FPLC Fast Protein Liquid Chromatography
GCAP Germ Cells Alkaline Phosphatase
GPO Glycerophopshate Oxidase
G3PDH Glycerol-3-phosphate Dehydrogenase
HBV Hepatite B Virus
HPLC High Performance Liquid Chromatography
IBOS Integration of Biosynthesis and Organic Synthesis
IAP Intestinal Alkaline Phosphatase
IPTG Isopropyl β-D-1-thiogalactopyranoside
LB Lysogeny Broth or Luria-Bertani broth
MEEC Membrane Enclosed Enzymatic Catalysis
PhoN-Sf/Se Acid Phosphatase from Shigella flexineri or from Salmonella enterica
Pi Phosphate
PLAP Placental Alkaline Phosphatase
pNPP para-Nitrophenylphosphate
PPi Pyrophosphate
RAMA Rabbit Muscle Aldolase
SDS-PAGE Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis
TNAP Tissue Non-specific Alkaline Phosphatase
vii
Figures Index
Figure 1.1 - The two types of aldolase mechanisms: type I Schiff base forming aldolase
and type II zinc enolate aldolase2. 4
Figure 1.2 - The four natural DHAP-dependent aldolases and their respective products
(adapted from 2). 6
Figure 1.3 – Enzymatic one-pot cascade reaction starting from DHA37, 42. 7
Figure 1.4 - Enzymatic one-pot cascade reaction starting from glycerol44. 8
Figure 1.5 - One-pot enzymatic cascade reactions for the synthesis of unnatural carbohydrates using AP. 9
Figure 1.6 – Model of the mammalian alkaline phosphatase based on the PLAP that shows the conserved active sites as sticks, the magnesium ions in green and the zinc ions in magenta. Molecular graphics created with YASARA49
. 11
Figure 1.7 – Overall reaction scheme of the reaction where both hydrolysis and transphosphorylation activities are shown37. 14
Figure 1.8 - Major intermediates in the proposed mechanism of action of alkaline phosphatase50
. 15
Figure 1.9 - Methods of immobilization of biocatalysts. 18
Figure 1.10 – Covalent immobilization of enzyme on the carrier: (A) active amino acid residue; (B) binding functionality of the carrier; (C) carrier; (D) spacer66. 19
Figure 1.11 - Mechanism of immobilization of proteins on epoxy-activated supports. 19
Figure 1.12 - Mechanism of immobilization of proteins on aldehyde-activated supports. 20
Figure 1.13 - Functional groups of Sepabeads® EC-EP and Sepabeads
® EC-HA
supports45
. 21
Figure 2.1 - Sepabeads from Resindion Srl and Immobeads-150 from Sigma-Aldrich. 23
Figure 2.2 – Janke & Kunkel Vibrofix VF1 Electronic. 25
Figure 2.3 – Eppendorf Thermomixer Compact. 26
Figure 2.4 – METTLER Analytical Balance AE 260 Delta Range. 26
Figure 2.5 – AKTA FPLC™ system. 27
Figure 2.6 – Standard BSA curve for protein concentration determination. 30
Figure 2.7 – p-Nitrophenyl phosphate (pNPP) hydrolysis by alkaline phosphatase. 30
viii
Figure 2.8 – Dihydroxyacetone phosphorylation and coupled assay with NADH. 32
Figure 2.9 – Pyrophosphate hydrolysis by alkaline phosphatase. 34
Figure 2.10 – One-pot cascade reaction starting from DHA. 35
Figure 2.11 – Glycerol phosphorylation and coupled assay with NAD+. 37
Figure 2.12 – One-pot cascade reaction starting from glycerol. 39
Figure 2.13 - Gene structure and gene expression in higher organisms73. 40
Figure 3.1 - SDS-PAGE gel of the original alkaline phosphatase solution. Lane (1) contains the molecular markers identified with the corresponding molecular weights in kiloDaltons, lane (2) corresponds to a 2 µg solution of AP while lane (3) corresponds to a 2.5 µg solution. 46
Figure 3.2 – DHAP formation over time using different AP concentrations. Reaction mixture contained DHA (100 mM), PPi (50 mM), and AP (2, 4 and 6 U/ml) in 1ml at pH 9. 48
Figure 3.3 – pH dependency of DHA phosphorylation. Reaction mixtures contained DHA (100 mM), PPi (50 mM), AP (6 U/ml) in 1 ml at pH 7, 7.5, 8, 9 and 10. The pH was set by addition of HCl or NaOH to the PPi, DHA mixture until the desired value was reached. 49
Figure 3.4 – Time course for the DHA cascade reaction using soluble AP. Reaction mixtures contained DHA (500 mM), PPi (100 mM), propanal (100 mM), AP (6 U/ml) and RAMA (6 U/ml),in 1 ml at pH 8. 51
Figure 3.5 – Alkaline phosphatase activity in the presence of different Pi concentrations. Reaction mixtures contained pNPP (100 mM), a DEA (1 M) and MgCl2 (0.5 mM) buffer pH 8, AP (5000x diluted) in 1.01 ml. 53
Figure 3.6 – Alkaline phosphatase activity in the supernatant during immobilization on different supports. Reaction mixture contained pNPP (100 mM), a DEA (1 M) and MgCl2 (0.5 mM) buffer pH 8 and 10 µl of supernatant 5x diluted in immobilization buffer: KPI (30 mM), MgCl2 (0.5 mM) in 1.01 ml. 54
Figure 3.7 – Time course for the formation of product in the DHA cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 56
Figure 3.8 – Time course for the formation of phosphorylated product in the DAH cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 56
Figure 3.9 – Time course for the formation of Pi in the DHA cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 56
ix
Figure 3.10 – Product formation with immobilized AP and PhoN-Sf after the 2 cycles of DHA cascade reaction. Reaction mixtures contains DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 57
Figure 3.11 – Phosphorylation of glycerol by soluble AP. Reaction mixture contains glycerol (100 mM), PPi (50 mM), AP (6 U/ml) in a buffer of glycine (450 mM), hydrazine (274 mM) and EDTA (2.4 mM) pH 9.5, G3P-DH (20 U/ml) and NAD+ (2.5 mM) in 1 ml. 58
Figure 3.12 – Time course for the formation of product in the glycerol cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contained glycerol (500 mM), PPi (250 mM), propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 59
Figure 3.13 – Time course for the formation of phosphorylated product in the glycerol cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contained glycerol (500 mM), PPi (250 mM), propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 59
Figure 3.14 – Time course for the formation of Pi in the glycerol cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contained glycerol (500 mM), PPi (250 mM), propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 59
Figure 3.15 – Product formation with immobilized AP and PhoN-Sf using different concentrations of glycerol on the glycerol cascade reaction. Reaction mixtures contained glycerol (0.5 and 3 M), PPi (250 mM), propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized phoN-Sf (1 U/ml) pH 6 in 1 ml. 60
Figure 3.16 – pH profile for the DHA and glycerol phosphorylation using the best mutants from PhoN-Se and their comparison with WT PhoN-Se and WT PhoN-Sf. Reactions contain DHA (100mM) or glycerol (100 mM), PPi (50 mM) and
acetate or Tris/acetate buffer (100 mM), together with PhoN (1 M). 61
Figure 3.17 – Amino acid introduced in the V78 position of the enzyme PhoN-Se. 62
Figure 3.18 – Time course of the DHA one-pot cascade reaction using WT PhoN-Se and the mutants V78L and V78Y. Reaction mixtures contained DHA (500 mM), PPi (250 mM), propanal (100 mM), Na-acetate pH 6 (20 mM), RAMA (6 U/ml), PhoN (0.5 and 1 µM) in 0.5 ml at pH 6. 63
Figure 3.19 – Time course of the DHA one-pot cascade reaction using WT PhoN-Se and the mutants V78L and V78Y. Reaction mixtures contained DHA (500 mM), PPi (250 mM), propanal (100 mM), Na-acetate pH 6 (20 mM), RAMA (6 U/ml), PhoN (1 and 2 µM) in 0.5 ml at pH 6. 64
Figure A1 – DHA Phosphorylation adding different concentrations of MgCl2. Reaction mixtures contain DHA (100 mM), PPi (50 mM), AP (6 U/ml) and MgCl2 (1 and 25 mM) at pH 8 and 32°C. 75
x
Figure A2 – PPi hydrolysis adding different concentrations of MgCl2. Reaction mixtures contain PPi (100 mM), AP (6 U/ml) and MgCl2 [100 (1:1), 66 (2:3) and 50 mM (1:2)] at pH 8 and 32°C. 76
Figure B1 – HPLC chromatogram obtained for the DHA cascade reaction using immobilized AP after 0 and 24 hours (black and blue line respectively). The identified peaks are in the UV spectra and correspond to: PPi – pyrophosphate, Phosphorylated product, DHA and Product. All unidentified peaks correspond to impurities originated either from the solutions utilized or the HPLC system. 77
Figure B2 – HPLC chromatogram obtained for the DHA cascade reaction using immobilized AP after 0 and 24 hours (black and blue line respectively). The identified peaks are in the IR spectra and correspond to: PPi – pyrophosphate, Pi – inorganic phosphate, DHA and Propanal. All unidentified peaks correspond to impurities originated either from the solutions utilized or the HPLC system. 78
Figure C1 – Time course for the formation of phosphorylated product in the 2nd
cycle of the DHA cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 79
Figure C2 – Time course for the formation of Pi in the 2nd
cycle of the DHA cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml. 79
Tables Index
Table 1.1 - Advantages and limitations associated to the use of immobilized enzymes (adapted from Cabral et al 65). 17
Table 2.1 - Reagents utilized during the course of the experiments. 24
Table 2.2 – Prestained protein marker, broad range kit. 28
Table 2.3 – Ammonium sulfate added to each mutant solution for precipitation of contaminant proteins. 43
Table 3.1 - Consumption of PPi and formation of Pi, in mM, for different PPi concentrations. 50
Table 3.2 – Comparison of obtained peak areas for the DHA cascade reaction with addition of different concentrations of MgCl2: 100 mM (1:1) and 50mM (1:2). 52
Table 3.3 - Results of expression and purification of the V78 L and V78 Y mutants. 62
1
1. Introduction
1.1 Motivation and background
1.1.1 Carbohydrate chemistry
Since the dawn of civilization humans have made use of carbohydrates in their natural
forms, such as cellulose in cotton, D-glucose in honey and sucrose in cane sugar. The first
documented synthesis of a sugar-like syrup presented in the chemical literature was the
preparation of formose from formaldehyde reported by Boutlerow (1861) and the first enzymatic
transformation of one carbohydrate to another was the oxidation of mannitol to D-fructose by
means of Bacterium aceticum documented by Brown (1886), both cited by Hudickly et al. (1996)1.
These biomolecules, which can be described as polyhydroxyaldehydes and ketones and
their derivatives, perform several roles in living organisms. Polysaccharides have a structure and
storage-related function and monosaccharides are the main providers of fuel for metabolism, as
an energy source or in biosynthesis. Different combinations of multifunctional monosaccharides,
having different stereochemistry, can pull together a vast range of complex structures which have
a vital job in various types of biochemical recognition, such as growth, immune system,
fertilization, metastasis, and several signal transduction events2.Carbohydrates also possess a
vast amount of commercial applications such as sweeteners, non-nutritive fat substitutes,
biodegradable polymers, agents for modifying viscosity and site-specific drug delivery3. This is
why carbohydrate chemistry is an area of particularly great interest to research.
The synthesis of these highly asymmetric and densely functionalized molecules using
classical chemical techniques has been reviewed 5-8
and it bears a number of difficulties such as:
1. Obtaining high yields in glycosidic linkage; the development of reliable chemical
methods for the glycosil activation is still an area of study in progress3.
2. Requirement for high regio- and stereoselectivity for the synthesis of mono- and
oligosaccharides, which frequently demands many sequential selective protection
and deprotection steps.
3. The fact that many carbohydrates are incompatible with non aqueous systems
creates problems for organic chemists, and extensive modification of hydrophilic
2
groups of carbohydrates have to be carried out just to obtain solubility in non-
aqueous media.
Isolation, purification and analysis of carbohydrates structures are also some of the
problems brought by this type of approach3.
A more advantageous approach seems to be the use of enzymes as catalysts for organic
synthesis. The regioselectivity displayed by enzymes, allowing complete chiral control, and the
mild reaction conditions allows the protecting group chemistry to be reduced, therefore reducing
the waste in the reaction2. The fact that most enzymes operate in aqueous solutions at room
temperature with a pH of around 7, makes the use of several enzymes in a one-pot reaction
sequence possible, since their reactions are often compatible with each other. These advantages
added to the great efficiency and environmentally benign nature of this methodology, make
enzymatic catalysis particularly useful for the synthesis of carbohydrates.
There are still some uncertainties in the potential of enzymes for the synthesis of
hydrocarbons, mainly involving the choice of enzymes necessary to perform the major
transformations in this area and their substrate tolerance, stability and inhibition. Other issues of
concern in the use of enzymes are their cost and frequent requirement for expensive cofactors.
Addressing these problems one may refer that many enzymes accept a broad range of
substrates, sometimes being broad enough to allow a wide variety of substrates to be accepted
with an acceptable rate when comparing to the rate of reaction of the natural substrate (which is
normally higher than the rate of unnatural substrates). Among these enzymes are some
esterases and lipases9, and aldolases like rabbit muscle fructose-1,6-diphosphate aldolase10,11.
The setback of cost and inactivation of enzymes can easily be diminished by exploiting
immobilization methods or containment in hollow fiber reactors or utilizing membrane enclosed
enzymatic catalysis, MEEC12, as these techniques allow the reutilization and recovery of the
enzyme and many times improve their stability13. The development of regenerating systems for
certain cofactors required by enzymes, as ATP, has allowed the use of catalytic amounts of
cofactors instead of their stoichiometrically use, which would be extremely expensive3.
The developing methodology of genetic engineering, with its continuing advances on
recombinant DNA technology, has made it possible to express many enzymes, making it possible
to modify their selectivity. Many of these enzymes, including enzymes of use for carbohydrate
synthesis, are commercially available for laboratory use. These advances contribute for a deeper
understanding of carbohydrate recognition and for a greater accessibility of their structures,
bringing new prospects in various fields of science.
3
1.1.2. Enzymatic Synthesis of Carbohydrates - Aldolases
The synthesis of carbohydrates involves two distinct branches: the preparation of
monosaccharides and related compounds, and the coupling of monosaccharides to form
oligosaccharides and oligosaccharide derivatives. The first task depends on the ability to
successfully form carbon-carbon bonds in a stereochemically defined way while the second
demands a high-yielding stereospecific approach to form the glycosidic bonds.
The enzymatic methodologies for both these reactions have been developed14,15; the
synthesis of monosaccharides, via enzymatic aldol addition reaction, catalyzed by aldolases and
their coupling to form oligosaccharides via specific, glycosidic linkages catalyzed by glycosidases
or glycosyltransferases has proved to be very useful2.
This study will focus on the enzymatic synthesis of monosaccharides; therefore a brief
consideration will be given to the enzymatic methods of performing the aldol reaction, one of the
most powerful methods of performing carbon-carbon bonds. The most acknowledged group of
enzymes that perform the aldol type reactions are the aldolases, a group of naturally occurring
enzymes that catalyze the stereospecific construction of C-C bonds16-19. Other types of enzymes,
not classified as aldolases, also catalyze aldol type reactions like Synthetases and Transferases
(transketolases) 3, 20. These enzymes are increasingly important but will not be mentioned since
they have not been used very much in carbohydrate synthesis. Another way to perform the
enzymatic aldol reaction is by the use of catalytic antibodies, compounds developed in recent
years to mimic the aldolases but with improved substrate specificity20, 21.
Aldolases belong to a large group of enzymes called lyases, present in all organisms,
which catalyze the reversible stereospecific addition of a ketone donor to an aldehyde acceptor.
To date over 30 aldolases have been identified and isolated16, 17, 22. Mechanistically two distinct
groups have been recognized (figure 1.1). Type I aldolases, found predominantly in animals and
higher plants, form a Schiff base intermediate in the active site with the donor substrate, which
subsequently adds stereospecifically to the acceptor22, 23. Type II aldolases, found primarily in
bacteria and fungi, contain a Zn2+ cofactor in the active site, which acts as a Lewis acid22, 24. Both
types of enzymes are rather specific for the nucleophilic donor substrates, but generally tolerate a
broad range of acceptor aldehyde components, allowing the synthesis of a variety of unnatural
sugars. The stereoselectivity of the aldol reaction is controlled by the enzyme and does not
depend entirely on the structure or stereochemistry of the substrate; therefore it is possible to
predict the stereochemistry of the products in which the introduction of two chiral centers is
involved25, 26.
4
Figure 1.1 - The two types of aldolase mechanisms: type I Schiff base forming aldolase and type II zinc
enolate aldolase2.
It is possible to divided aldolases in four main groups, on the basis of the donor substrate
accepted by the enzyme. The first group uses dihydroxyacetone phosphate (DHAP) to produce 2-
keto-3,4-dihydroxy adducts after reaction with an aldehyde. The second group uses pyruvate or
phosphoenol pyruvate to form 3-deoxy-2-keto acids. The third group uses acetaldehyde as the
donor to form 3-hydroxyaldehydes and the fourth group uses glycine as the donor to produce
different -substituted amino acids. Apart from these main groups other aldolases have been
known to perform the aldol reaction but their substrate specificity has not been examined in terms
of their use in synthesis.2
The most versatile and widely used aldolases in the synthesis of ketose sugars and
related derivatives have been the DHAP dependent aldolases27-29. These types of enzymes have
the advantage of nearly complete chiral control over the newly formed stereogenic centers2 and
accept a broad range of acceptor substrates16, having more than 100 aldehydes been used as
acceptors to prepare monosaccharides.2, 16, 18 DHAP dependent aldolases also catalyze the
condensation of pentose and hexose phosphates with DHAP, consequently extending the sugar
chain by three carbons while introducing two new stereogenic centers. This provides a route to
novel high-carbon sugars, which are difficult to obtain from either chemical synthesis or natural
sources. Four complementary DHAP dependent aldolases are commercially available2 each of
which produces one of the four possible stereoisomers30.
Given the fact that these aldolases show a very strict dependence for the donor
substrate, the availability of DHAP is an essential issue for the development of their practical use.
DHAP is commercially available, although it is extremely expensive, and several chemical and
5
enzymatic methods for generating this compound have been developed31-33. Among these
methods, several synthetic procedures of DHAP precursors, achieved by chemical
phosphorylation of the dihydroxyacetone dimer, have been developed2, 3, 34-36 but most of these
chemical procedures involve many protection and deprotection steps, give low overall yields,
require complicated multistep purification procedures and involve the use of expensive or toxic
reagents37.
Enzymatic phosphorylation of dihydroxyacetone appears to be more suitable in the sense
that it would require fewer reaction steps and it‟s a cleaner and easier method. The most
convenient method is the in situ generation of 2 equivalents of DHAP from fructose 1,6-
diphosphate (FDP) by the enzyme triosephosphate isomerase or FDP aldolase2, 10, 20, 22 although
in some cases when the reaction does not go to completion, the presence of excess of FDP
complicates the isolation of products. Other method is based on the enzymatic phosphorylation of
dihydroxyacetone using kinases, which need the in situ regeneration of ATP33, 37. L-
Glycerolphosphate oxidation by glycerophosphate oxidase (GPO) also generates DHAP. This
can be achieved by the phosphorylation of glycerol using glycerol kinase37, 38. Recent work has
demonstrated that phosphorylation of dihydroxyacetone is also possible by the use of acid
phosphatase from Shigella flexineri (PhoN-Sf) and cheap pyrophosphate (PPi) as a phosphate
donor37. In this work the DHAP is generated in situ and then coupled to an aldehyde in an
aldolase-coupled reaction using rabbit muscle aldolase (RAMA). The end product was then
dephosphorylated by the already present phosphatase (PhoN-Sf), resulting in a product with new
stereocenters formed with high stereospecificity. As previously mentioned, the fact that enzyme-
catalyzed enzyme reactions are often compatible to each other makes it possible to combine
several enzymes in a one-pot, multistep reaction sequence as described in previous work 37, 39.
This prevents the retro-aldol reaction from occurring and shifts the equilibrium to completion. It
was also shown37 that the phosphate group hydrolyzed by the enzyme from the
dephosphorylation of the aldol adduct was re used to phosphorylate DHA producing more DHAP
creating higher levels of conversion.
1.1.3. The IBOS Project
The IBOS project, Integration of Biosynthesis and Organic Synthesis, is a long term
research project with the ultimate aim of developing an economically feasible process for the
synthesis of heavily functionalized heterocyclic compounds via enzymatic cascade reactions. The
heterocyclic end products, mainly sugar- and azasugar-like compounds, are of special interest in
the pharmaceutical and fine chemical industry. These sugars are used in medicine as inhibitors of
6
glucosidases in the treatment of diabetes and other diseases connected to the metabolism of
carbohydrates as they mimic the structures of monosaccharides.
These kinds of molecules are hard to synthesize in the traditional chemical way due to
their hydrophilic nature and the high number of stereocenters present. This problem is overcome
by performing the synthesis by enzymatic methods through the aldol condensation reaction
between DHAP and various aldehydes using DHAP dependent aldolases.
The benefits obtained by the employment of DHAP dependent aldolases have been
clarified above: broad substrate specificity for the acceptor aldehydes16
and nearly complete
chiral control over the 2 newly formed stereocenters2. In nature there are four different DHAP
dependent aldolases: D-fructose-1,6-diphosphate aldolase, L-fuculose-1-phopshate aldolase, L-
rhamnulose-1-phosphate aldolase and D-tagatose-1,6-diphosphate aldolase. All four types of
DHAP-dependent aldolases have been explored for synthetic applications and each reaction
generates one unique product whose stereochemistry at C-3 and C-4 is complementary to the
others2 (figure 1.2). The most widely used and best-studied aldolase for aldol reactions is the
fructose-1,6-diphosphate aldolase from rabbit muscle (RAMA)40,41 therefore this is the one mostly
used in this IBOS project.
Figure 1.2 - The four natural DHAP-dependent aldolases and their respective products (adapted from 2).
The synthesis of the necessary DHAP, an intrinsically unstable and very expensive
compound, is performed via enzymatic phosphorylation of DHA. The phosphorylation has been
performed by bacterial non-specific acid phosphatases, PhoN-Sf/Se, which performs the
transphosphorylation reaction using pyrophosphate as a cheap phosphate donor. This enzymatic
method is extremely convenient since it allows the integration of the phosphorylation steps with
7
the aldol reaction, the DHAP is produced in situ and consequently less exposed to degradation37.
Thus an enzymatic one-pot cascade reaction was developed and patented by this
group37, 42
in which starting from DHA and PPi, using the acid phosphatase, a
transphosphorylation reaction occurs to generate DHAP. DHAP is then coupled to an aldehyde
by the aldolase RAMA to give the phosphorylated aldol product that is then dephosphorylated by
the acid phosphatase to give the final product. During catalysis an enzyme-phosphate
intermediate is formed and this can be used to phosphorylate another DHA molecule, thereby
recycling the phosphate group as depicted in figure 1.3:
Figure 1.3 – Enzymatic one-pot cascade reaction starting from DHA37, 42.
The advantages of applying this kind of multi-step reaction rely on the fact that PPi is a
very cheap donor and is the driving force of all the reaction, the fact that there is a
thermodynamic optimization and the in situ generation of DHAP, which is an unstable and highly-
priced compound.
This cascade was found to be very efficient leading to 60% conversion43, however there
were still some limitations such as the difficulties obtained in the separation of the product from
DHA and the fact that the high reactivity of DHA may somehow limit the efficiency of the cascade
in the sense that it can react with the enzymes and after 1 hour the reaction mixture is brownish
and turbid.
Recently an alternative way for the in situ generation of DHAP, other than the one
starting from DHA, was developed. Starting from glycerol, phosphorylation is performed by the
acid phosphatase PhoN-Sf to produce glycerol-3-phosphate in 2 enantiomers D/L and then the
glycerophosphate oxidase, GPO, is used to oxidize the L form to DHAP. H2O2 is formed but it is
reduced by catalase to reform the necessary O2. The D-glycerophosphate can be
8
dephosphorylated by the acid phosphatase and brought back into the cycle (figure 1.4). So
essentially in this scheme a cascade reaction with 4 enzymes is performed, which should work at
the same pH to have a one-pot reaction.
Figure 1.4 - Enzymatic one-pot cascade reaction starting from glycerol44.
A similar experiment was already reported in literature45 using phytase as a
phosphorylating enzyme instead of the phosphatase. This enzyme is only active at pH 4,
therefore it was necessary to switch from pH 4, for the phosphorylation with phytase, to pH 7.5 for
the oxidation with GPO and the aldol reaction with RAMA and then switch again to pH 4 for the
dephosphorylation of the product. Another drawback of using phytase is the requirement for very
high concentrations of glycerol, since this enzyme tends to hydrolyze PPi and not transfer the
phosphate to glycerol.
By using the acid phosphatase PhoN-Sf the whole cascade reaction is performed at pH
6, so there is no need for changes in pH and it is possible to use lower concentrations of glycerol
since PhoN-Sf is very efficient in the transphosphorylation reaction.
Comparing the two cascade reactions (starting from DHA and glycerol) similar
conversions are obtained although the glycerol cascade reaction is slightly slower than the DHA
one which may be due to the oxidation step by GPO. This fact was proved by increasing the
amount of GPO: the reaction was not only much faster but also led to very high yields, up to
90%44
. The reason for better results achieved for the glycerol cascade reaction is probably
because of the absence of DHA which is unstable and reacts with proteins.
9
1.1.4. The Alkaline phosphatase approach
The focus of this study is on the alkaline phosphatase (AP) route.
Given the complexity of the cascade reactions, it is very important to optimize their
conditions in terms of pH, since the pH optimum of the oxidase (GPO) and aldolase (RAMA) is
around 7-8 and that of the acid phosphatase is around 6. The acid phosphatase was successfully
used in these cascades so far but its nature doesn‟t allow performing the reaction at pH above 6.
Thus, the use of an alkaline phosphatase that has a pH optimum around pH 8 to 9 could improve
the conversions and optimize the cascade reaction further leading to higher yields of the non-
natural carbohydrates formed (figure 1.5).
Figure 1.5 - One-pot enzymatic cascade reactions for the synthesis of unnatural carbohydrates using AP.
Previous studies by Pradines et al, 1988 and 199146, 47 have reported the use of alkaline
phosphatase in the enzymatic phosphorylation of polyhydroxylated substrates in reverse
hydrolysis conditions. In the first study different substrates were considered, including
polyfunctional substrates like polyhydroxylated ketones and aldehydes, unsaturated alcohols and
amino-alcohols, leading to appreciable yields of phosphorylation and high regioselectivity.
Various types of phosphate donors were analyzed; phosphate, pyro- and polyphosphates and the
best results, in terms of velocity, were obtained with pyrophosphate46.
10
The second publication disclosed great potential in the usage of the alkaline phosphatase
since it concerned the optimization of this enzymatic phosphorylation for the large-scale
production of glycerol-1-phosphate, a compound of increasing importance in medicine for calcium
and magnesium transport47
. In this paper by Pradines et al, using glycerol as substrate and
phosphate as a phosphate donor (chose over pyrophosphate due to its higher solubility) alkaline
phosphatase is used both in the free and immobilized forms to optimize the synthesis of glycerol-
1-phosphate. The influence of pH, phosphate donor concentration and possible inhibitions by the
substrate and the product were measured in order to achieve the highest yields. The best results
were obtained with the immobilized enzyme on corn grits (EURA-MA, a cellulose based support)
since the inhibition by the substrate and product are severely diminished by the immobilization.
The yields of production of glycerol-1-phosphate (calculated from the initial concentration in
phosphate) obtained were as follows: 41.3% for the immobilized enzyme in a continuous tank
reactor, 18% for the immobilized enzyme in a packed-bed reactor and 12% for the free enzyme.
This demonstrates the feasibility of the usage of alkaline phosphatase in large-scale
production of phosphorylated compounds, especially glycerol-1-phosphate which is extremely
useful in the glycerol cascade reaction.
1.2. Alkaline Phosphatase
1.2.1. Origin and distribution of AP
Alkaline phosphatase (E.C.3.1.3.1) belongs to the class of hydrolases and acts on
phosphate groups. This enzyme catalyzes the hydrolysis of almost every phosphomonoester to
give Pi and the corresponding alcohol, phenol, or sugar, etc., but also catalyzes
transphosphorylation reactions (when in presence of large concentration of a phosphate
acceptor).
It‟s a homodimeric enzyme with a molecular weight of approximately 160 kDa and each
active site region contains three metal ions, two zinc and one magnesium ion, all necessary for
enzymatic activity48.
11
Figure 1.6 – Model of the mammalian alkaline phosphatase based on the PLAP that shows the conserved
active sites as sticks, the magnesium ions in green and the zinc ions in magenta. Molecular graphics
created with YASARA49.
One of the first reports on alkaline phosphatase appeared around 1907 when Suzuki et
al. suggested that phosphatases constituted a separate class of eukaryotic enzymes. The
enzyme we now know as alkaline phosphatase was defined by the work of Grosser & Husler
(1912) and von Euler (1912), who showed that while it was present in a variety of tissues, the
enzyme, which could hydrolyze glycerophosphate and fructose 1 -6 diphosphate, was present in
highest amount in intestinal mucosa50
.
In nature alkaline phosphatases are found in many organisms, both prokaryotes and
eukaryotes. The enzymes are present in bacteria and fungi, are relatively abundant in fish and
mammals although they are absent from higher plants51. In mammals, tissues with high
concentrations of alkaline phosphatase include intestine, kidney, placenta, bone, liver, lung, and
spleen52 even though its distribution within a particular tissue is not homogeneous. In general one
can say that alkaline phosphatase is abundant in the tissues that are concerned with the transport
of nutrients, often being present in secretory organs and developing tissues51. In Homo sapiens,
three out of four AP isozymes are tissue-specific, one is placental (PLAP), the second is from
germ cell (GCAP), and the third is intestinal (IAP). They are 90–98% homologous, and their
genes are clustered on the same chromosome. The fourth is nonspecific and can be found in
bone, liver, and kidney. It is about 50% identical with the other three, and its gene is located on
another chromosome53-55.
Although the main features of the catalytic mechanism are conserved comparing
mammalian and bacterial AP, mammalian AP have higher specific activity and Km values; have a
more alkaline pH optimum; display lower heat stability; are membrane-bound and are inhibited by
12
L-amino acids and peptides through an uncompetitive mechanism48. These properties, however,
differ remarkably among the different mammalian AP isozymes and are likely to reflect very
different in vivo functions.
Finally, while bacterial E. coli alkaline phosphatase, ECAP, is located in the periplasmic
space of the bacteria, mammalian APs are ectoenzymes bound to the plasma membrane via a
glycosyl-phosphatidylinositol anchor56.
Regarding the substrate specificity it was accepted, until recently, that alkaline
phosphatase was strictly specific for phosphomonoesthers (Stadman, 1961; Portmann, 1975),
however it has been shown that not only the E. coli enzyme (Heppel, Harkness & Hilmoe, 1962)
but also the mammalian enzyme (Fernley and Walker, 1966; Russel, 1965; Cox&Griffin, 1965 &
1967; Moss. Eaton, Smith & Whitby, 1966, 1967) have pyrophosphatase activity, hydrolyzing
inorganic pyrophosphate PPi as well as ATP, ADP, AMP and similar compounds48. The enzyme
also hydrolyses oxyphosphate monoesters, a variety of O- and S-phosphorothioates,
phosphoramidates and thiophosphate50
. In presence of high concentrations of acceptor the
enzyme also performs transphosphorylation reactions.46-48.
1.2.2. The role of AP in nature
Although alkaline phosphatase is widely found both in bacteria and in mammals a precise
physiological function has not been assign 51, 57.
In bacteria, even though its role could be the nonspecific hydrolysis of phosphate esters,
it is still reasonable to consider the possibility of other functions such as phosphate transport. In
fact, given that the enzyme readily binds phosphate covalently and non-covalently, especially in
acidic pH57, it could transport and concentrate phosphate from a more acidic medium to the
interior of the cell under conditions of low phosphate57.
In mammals several functions can be recognized according to its distribution throughout
the various tissues. Bone alkaline phosphatase plays an important part in ossification51 with two
possible roles being proposed: 1) the precipitation of calcium phosphate is induced by the
localized production of high concentrations of Pi due to phosphatase activity and 2) the enzyme
allows crystal growth at nucleation sites in the matrix by ensuring the continuous removal of PPi
which is considered a crystal „poison‟51. In tissues with high concentrations of alkaline
phosphatase, like intestine, kidney and placenta, the enzyme location at the absorptive surface
suggest a direct role in the transport of nutrients across the epithelial membrane51. It has been
reported that following ingestion of carbohydrates an increase in plasma phosphatase was
13
observed, also ingestion of fat by rats caused an increase in intestinal alkaline phosphatase
which was then detected in the blood plasma51.
The tissue specificity of the human enzymes has been correlated to an additional
function. In tissue-nonspecific AP, TNSAP, this specificity involves binding to collagen54. In PLAP,
evidence suggests a role in cell division in both normal and transformed cells. This role probably
occurs through its phosphatase activity against phosphorylated proteins53. Furthermore, it has
been suggested that PLAP may be involved in the transfer of maternal immunoglobulin G, IgG to
the fetus53.
1.2.3 Protein structure and mechanistic issues
The overall structure of mammalian alkaline phosphatase (human placental, PLAP) is a
dimmer of identical subunits, each containing 484 residues, four metal atoms, one phosphate ion
and 603 water molecules48. The two monomers are connected by a two-fold crystallography axis.
The surface of PLAP is poorly preserved with that of the E. coli enzyme, with only 8% residues in
common, although the core is preserved.
In the active site the residues essential for catalysis are common in all alkaline
phosphatases: the catalytic serine residue and the three metal ion sites, M1 occupied by Zn2+, M2
occupied by another Zn2+
and M3 occupied by Mg2+, as well as their ligands48,53. Half of the
enzyme surface corresponds to three clearly identifiable regions, only present in mammalian
enzymes, the first is a long N-terminal -helix, forming an arm that embraces the other monomer,
second an interfacial flexible loop, „crown domain‟, formed by the intersection of a 60-residue
segment from each monomer and third is a metal binding domain, containing an additional metal
ion, M4. This additional non-catalytic metal-binding site, which appears to be occupied either by
magnesium or calcium 48, 53, is not present in bacteria and its architecture is conserved in all
human and mouse alkaline phosphatases and most probably represents a novel feature common
to all mammalian AP‟s48. Its functional and structural significance, however has not been well
established to date, but probably could be related to the conformational stabilization of the two -
helices that form the peripheral site48.
The availability of the PLAP structure facilitated modeling the human germ cell AP
(GCAP), intestinal AP (IAP) and tissue-nonspecific AP (TNAP, a.k.a. liver/ bone/kidney type AP)
isozymes, revealing that all the novel features discovered in PLAP are conserved in those human
isozymes as well53.
Alkaline phosphatase reactions can fit into the general kinetics presented in figure 1.7:
14
Figure 1.7 – Overall reaction scheme of the reaction where both hydrolysis and transphosphorylation
activities are shown37.
The reaction of phosphate monoesters with alkaline phosphatase is known in detail50, 58-
60. The Mg2+ ion, M3, in the active site activates a bound water molecule making it a better
nucleophile. A Zn2+
ion, M1, coordinates the oxygen of the phosphate monoester, activating the
leaving group. A second oxygen coordinates the second Zn2+ ion, M2, forming a phosphate
bridge between the two metal ions, while the other phosphate oxygens form hydrogen bond with
the guanidine group of Arg166 (figure 1.8). Upon binding of the phosphate monoester the OH
group of the serine residue becomes deprotonated for nucleophilic attack on the phosphorus
center. Serine is the nucleophile in the first half of the reaction and would occupy the position
opposite to the leaving group, RO-, in a five-coordinate intermediate (figure 1.8). Dissociation of
the alcohol group (RO-) and formation of a covalent phosphoserine intermediate takes place. This
intermediate corresponds to the species E-Pi in figure 1.7 (and in figure 1.8). The coordination
site on M1 (Zn1 in figure 1.8) previously occupied by the alcoxide RO- can be occupied by a
water molecule that must dissociate a proton to become Zn-OH.
In the second step the second Zn2+
metal ion, Zn2, activates the leaving group by forming
a coordinate bond with the ester oxygen of the phosphoserine intermediate. The hydroxyl
connected to Zn1 is in position to be the nucleophile for the hydrolysis of the phosphoserine
intermediate and leads to the formation of E.Pi intermediate in which the phosphate is still bound
to the active site. The intermediate E.Pi forms as the phosphate moves away from the serine and
as one of the phosphate oxygen coordinates again a Zn2+ ion, Zn1 to reestablish a phosphate
bridge.
15
Figure 1.8 - Major intermediates in the proposed mechanism of action of alkaline phosphatase60.
As illustrated in figure 1.8 the phosphate group in this intermediate E.Pi is in close
association with the two Zn2+ ions. It bridges both Zn2+ ions, connected by two phosphate oxygen
atoms. The other two phosphate oxygen atoms are tightly held by the amino functions of the
guanidine group of Arg16650, 60.
At acidic pH the hydrolysis of the phospho-enzyme, E-Pi is the rate limiting step while at
alkaline pH the dissociation of the non-covalent enzyme-phosphate complex, E.Pi is the rate
determining step.
1.2.4. Applications of AP
Alkaline phosphatases may potentially be employed as therapeutic agents and
therapeutic targets and show several uses in clinical medicine and in biotechnology.
An example of its use in clinical medicine was described in a technique for labeling
monoclonal antibodies61. The procedure, called the alkaline phosphatase monoclonal anti-alkaline
phosphatase (APAAP) method, gives excellent immunocytochemical labeling of tissue sections
and cell smears, comparable in clarity and intensity to that achieved with immunoperoxidase
labeling. For this reason the APAAP technique is particularly suitable for labeling cell smears (for
both cytoplasmic and surface-membrane antigens) and for detecting low numbers of antigen-
16
bearing cells in a specimen (e.g., carcinoma cells in a malignant effusion).
The method was also applicable to the detection of antigenic molecules following their
electrophoretic transfer from SDS-polyacrylamide gels to nitrocellulose sheets ("immunoblotting")
61.
Another medical use is the application of alkaline phosphatase to label HBV (Hepatitis B
Virus) DNA as probe to detect the HBV DNA in hepatitis serum62. The alkaline phosphatase is
coupled with polyethyleneimine using P-benzoquine as cross-linking reagent. The modified
phosphatase is then covalently linked to single strand DNA and this DNA enzyme complex is
tested for blot hybridization, after hybridization and incubation with a substrate solution.
Sequences complementary to the probe can be visualized directly in only 1 h as opposed to the
32 P labeled probe which takes 1 week62. This experiment certified that the virus DNA detection
method is sensitive, specific, rapid, safe and economical and clinically useful.
A different DNA detection method was developed in which chemiluminescent substrates
are employed63. Chemical or enzymatic removal of a protecting group from stable dioxetanes
produces an unstable aryloxide dioxetane, which decomposes to provide the observed
chemiluminescence. In this research63 it has been discovered a particularly useful
chemiluminescent substrate for alkaline phosphatase, phenylphosphatesubstituted dioxetane.
With this system, the luminescent reaction can be used for ultrasensitive detection of alkaline
phosphatase-linked antibodies and DNA probes63.
Alkaline phosphatases can also be useful for industrial waste treatment: the p-nitrophenyl
phosphate activity assay for this enzyme was modified for use in freshwater sediment64. Studies
indicated that the recovery of purified alkaline phosphatase activity was 100% efficient in sterile
freshwater sediments when optimized incubation and sonication conditions were used.
Significant correlations between phosphatase and total viable cell counts, as well as
adenosine triphosphate biomass, suggested that alkaline phosphatase activity has utility as an
indicator of microbial population density and biomass in freshwater sediments64.
1.3. Enzyme immobilization
Immobilization of biocatalysts ensures the preservation of their catalytic activity and
allows their repeated or continuous use. Immobilization methods have been applied to a wide
array of biocatalysts, ranging from pure enzymatic extracts to whole microbial cells or even
animal and vegetal tissues. When applied to enzymatic extracts in an industrial setting,
immobilization offers several advantages and some restrictions, the most relevant of which are
listed in table 1.1.
17
Table 1.1 - Advantages and limitations associated to the use of immobilized enzymes (adapted from Cabral
et al 65).
Advantages Particular Aspects
Retention of the catalyst inside the reactor Allows reutilization and continuous processes Possibility of operating on high dilution rates
without the risk of wash-out
High catalyst concentrations Allows higher volumetric production rates
Faster conversion, relevant when secondary reactions are an issue
Controlled microenvironment
Allows manipulation of enzymatic activity and specificity
Improves enzyme stability Protects the enzyme against shear stress
Easy product isolation Minimizes product contamination
Precise control of bioconversion time
Limitations Particular Aspects
Loss of catalytic activity May occur during the immobilization process, during the conversion or due to the physical
properties of the immobilization matrix.
Empiric process Specific optimization needed for each particular
application Complex control and modelling
The different types of immobilization of biocatalysts have been the subject of several
classification systems. One of them, adapted from Cabral et al.65, is presented in figure 1.9.
18
Figure 1.9 - Methods of immobilization of biocatalysts.
Alkaline phosphatase has a dimeric structure that dissociates easily leading to
inactivation. Immobilization by covalent bonds may play an important role in avoiding dissociation
of the enzyme by keeping the sub-units together.
Covalent immobilization of enzymes usually provides the strongest linkages between
enzyme and carrier, compared with other types of enzyme immobilization methods such as non-
covalent adsorption-based enzyme immobilization66. Therefore the leakage of enzyme from the
matrix is minimized with covalently bound enzymes66.
Covalent binding of an enzyme to a carrier is generally based on a chemical reaction
between the active amino acid residues located on the enzyme surface and active functionalities
that are attached to the carrier surface (figure 1.10). The covalently immobilized enzyme may be
regarded as a composite consisting of the components carrier, spacer, linkage and enzyme. To
achieve efficient linkage, the functionality of the carrier and/or the enzyme must be activated
before immobilization and often carriers are activated before their use for binding enzymes66. The
physical and chemical nature of the carrier strongly dictates the performance, for instance activity,
selectivity, stability and particularly the application of the immobilized enzymes obtained.
19
Figure 1.10 – Covalent immobilization of enzyme on the carrier: (A) active amino acid residue; (B) binding
functionality of the carrier; (C) carrier; (D) spacer66.
During the immobilization process the enzyme molecules are brought to close contact
with the surface of the carrier and as a consequence the subsequent interactions may disturb the
native forces that maintain the enzyme native structure, leading to modifications on the structure
and function of the enzyme depending on the chemical nature of the carrier used66. However, it
has been found that the enzyme usually adopts a conformation stabilized by the interaction of the
enzyme and the carrier. If the new induced conformation resembles the native structure of the
enzyme the immobilized enzyme is stabilized, otherwise it will be deactivated. Therefore it is
extremely important to find the right carrier for a given enzyme66.
Epoxy- and amino-activated supports are able to form very stable covalent linkages with
different amino acid residues of the enzyme (amino, thiol, and phenolic ones) under very mild
experimental conditions67. In addition, these supports are very stable during storage and also
when suspended in neutral aqueous media. Hence, they can be easily handled before and during
immobilization procedures.
The immobilization of enzymes in epoxy functionalized carrier supports usually follows a
two-step mechanism: first a rapid mild physical adsorption between the enzyme molecules and
the support is produced, and secondly the covalent reaction between adsorbed protein and epoxy
groups occurs (figure 1.11).
Figure 1.11 - Mechanism of immobilization of proteins on epoxy-activated supports.
20
The immobilization of enzymes in amino functionalized carrier supports requires an extra
step: first the carrier needs to be activated; therefore an aldehyde is bound to the amino
functionalized group of the carrier. Then the covalent bonding between the active amino groups
of the enzyme and the aldehyde activated groups of the support occurs (figure 1.12).
Figure 1.12 - Mechanism of immobilization of proteins on aldehyde-activated supports.
Even though other immobilization techniques inside porous supports can increase the
enzyme operational stability by preventing any intermolecular process (proteolysis, aggregation)
and by preserving the enzyme from interactions with external interfaces (air, oxygen, immiscible
organic solvents, etc.), these techniques do not necessarily increase the conformational stability
of the enzyme67. This kind of stability should be achieved if the immobilization of each enzyme
occurs through several residues. This way, all the residues involved in immobilization preserve
their relative positions and the enzyme is unaffected by conformational changes promoted by
heat, organic solvents, or any other distorting agents67. Thus, multipoint covalently immobilized
enzymes should become more stable than their soluble counterparts or than randomly
immobilized derivatives
21
In this study two epoxy: Immobeads 150 and Sepabeads® EC-EP and one amino-
functionalized supports, Sepabeads® EC-HA, were tested for alkaline phosphatase
immobilization. All supports are macroporous, acrylic polymer matrix spherical beads.
Sepabeads® EC-EP is a highly activated support functionalized with short chain epoxy
groups while the Sepabeads® EC-HA supports are functionalized with amino groups on a longer,
more complex, spacer (figure 1.13). Both are very rigid supports that may be used in stirred tanks
or bed reactors. These supports have low swelling tendency in high molar solutions and in
common solvents. Also, they demonstrate outstanding mechano-osmotic stability given by
intense crosslinking. The standard grade beads have a diameter of 150-300 µm with an average
pore diameter of 30-40 nm and a specific gravity of 1.13 g/ml68.
Figure 1.13 - Functional groups of Sepabeads® EC-EP and Sepabeads® EC-HA supports45.
Immobeads 150 is activated similarly to Sepabeads® EC-EP, having on its surface a
dense monolayer of reactive and stable epoxy groups. The particle size is 100-250 micrometre
and the loss on drying is inferior to 10%69. This kind of supports is especially designed to have a
low diffusion limitation that allows for the immobilization of enzymes with high specific activities.
Only a few publications appeared on immobilization of alkaline phosphatase. An
experiment by Taylor et al 70
showed that active AP may be bound to different types of carriers
such as glass, agarose-based carrier (Sepharose) and epoxy-activated acrylate-based supports
(Sepharon HEMA 1000 and Eupergit C). Other kind of supports were also described such as
Sepharose-4B, a macroporous cellulose and chitosan-based carrier69, as well as the already
mentioned paper by Pradines et al 46 which successfully immobilized AP in corn-grits (EURA-MA)
for the large scale production of glycerol-1-phosphate.
22
1.4. Aim of this Study
The main goal of this project is to investigate the uses and limitations of the enzyme
alkaline phosphatase (E.C.3.1.3.1), contributing to the aim of the overall IBOS project of
converting achiral compounds into non-natural carbohydrates via enzymatic cascade reactions.
The information found in literature concerning the use of the enzyme in this kind of reactions is
scarce; bearing that in mind its use in the two cascade reactions, the first starting from DHA and
the second from glycerol were optimized.
In a first stage of the project the phosphorylation of DHA and glycerol by the alkaline
phosphatase was studied at different pH values. The time course of the product formation and the
amounts formed were determined by spectrophotometric coupled-assays. The influence of the
presence of metal ions in the reaction mixture was also studied in terms of the impact on the
enzyme activity.
In a second phase of the project, the cascade reactions using the alkaline phosphatase
instead of the acid phosphatase were studied and the reaction conditions, pH, enzyme
concentrations optimized. The effect of immobilization on the performance of the enzyme in
covalent-binding supports was also investigated in a batch cascade. Different commercially
available epoxy functionalized supports and an aldehyde-activated amino support suitable for
industrial use68, 69 were tested for optimum stability/activity. After selecting the best performing
enzyme preparation, the DHA and glycerol enzymatic cascade reactions were compared.
Finally, even though the main goal of this project is the optimization of the use of alkaline
phosphatase in both cascade reactions for the production of asymmetric heterocyclic compounds,
a secondary objective is to further contribute to the developing IBOS project. This includes, if
possible, the collection of data on the expression, purification and isolation of new mutants of acid
phosphatase from Salmonella enterica ser. Typhimurium (PhoN-Se): V78Y, V78L and V78H and
study their use in the same DHA cascade reaction.
23
2. Materials and Methods
2.1. Alkaline Phosphatase
2.1.1. Calf Intestine Alkaline Phosphatase
The alkaline phosphatase, from bovine calf intestine, utilized during the enzymatic
experiments was supplied by Sigma Aldrich. According to the supplier, the solution has an activity
of 5611 U/ml and is in a 50% glycerol solution containing 5 mM Tris, 5 mM MgCl2 and 0.1 mM
ZnCl2 at pH 7 necessary for enzyme activation.
2.1.2. Immobilization supports (epoxy and amino groups)
Sepabeads® EC-EP and Sepabeads® EC-HA were supplied by Resindion Srl (Mitsubishi
Chemical Corporation). Immobeads-150 was supplied by Sigma-Aldrich.
Figure 2.1 – Sepabeads from Resindion Srl and Immobeads-150 from Sigma-Aldrich.
2.1.3. Reagents
The main chemicals used during this research project, essentially the ones used in the
assays for characterization of the enzyme as well as the ones used for the phosphorylation and
the cascade reactions, are listed in table 2.1.
24
Table 2.1 - Reagents utilized during the course of the experiments.
Chemical Supplier Purity (min)
Acetic acid Pierce 99.8%
Diethanolamine Alfa Aesar 99%
1,3 – Dihydroxyacetone Dimer Sigma-Aldrich 97%
Disodium pyrophosphate Sigma-Aldrich Practical grade
EDTA disodium salt Sigma-Aldrich 99.5%
Glycerol anhydrous Fluka 99.5%
Glycine Sigma-Aldrich 99%
Hydrochloric acid Fluka 25%
Magnesium Chloride hexahydrate Sigma-Aldrich 99%
NAD+ (from yeast) Sigma-Aldrich 96.5%
NADH Sigma-Aldrich 98%
para-nitrophenylphosphate (pNPP) Fluka 97%
Propionaldehyde Acros Organics 97%
Sodium Hydroxide pellets Acros Organics 98%
Tetrasodium pyrophosphate Sigma-Aldrich 95%
Trisma base Sigma-Aldrich 99.9%
Zinc chloride anhydrous Sigma-Aldrich 99.9%
The pyrophosphate (PPi) used in the cascade reactions was prepared according to
literature data, using a 3 ½: 2 ratio of disodium pyrophosphate to tetrasodium pyrophosphate,
both listed above.
25
2.1.4. Equipment
High Performance Liquid Chromatoghraphy
For time course studies 20 µl of the reaction mixture was diluted 10 times (unless
specified otherwise) before injection into the HPLC. Analyses were performed using an Alltech
OA 1000 organic acid column (0.65x30cm) equipped with a DIONEX 580 LPG pump and
DIONEX UVD-340/Shodex RI-101 detector. The column was eluted with 25 mM H2SO4 at a flow
rate of 0.4 mL min-1. The Chromeleon software program (Dionex) was used for HPLC data
acquisition and evaluation.
Rotator (Vortex)
A Janke & Kunkel Vibrofix VF1 Electronic was utilized. This rotator allows the gentle but
efficient mixing from 500 rpm to 2500 rpm, of eppendorf vials and test tubes with different shapes
and sizes. The angle of rotation is adjustable.
Figure 2.2 – Janke & Kunkel Vibrofix VF1 Electronic.
Thermomixer
An Eppendorf® Thermomixer Compact was utilized. This thermomixer is equipped with a
rack that allows the simultaneous heating of up to 24 1.5 ml eppendorf Safe-Lock tubes. It is fully
programmable; capable of heating or cooling samples from 1°C to 99°C and of agitating from 300
rpm to 1500 rpm.
26
Figure 2.3 – Eppendorf Thermomixer Compact.
Spectrophotometer
For spectrophotometer measurements a UV-VIS Cary 50 Bio Varian Array
Spectrophotometer was used. The Simple Reads software program was used for data acquisition
and evaluation.
pH-meter
An AG Herisau Methrom 632 pH-meter was used for pH readings. The pH combination
electrodes with refillable liquid reference system Sentix 51,52 from WTW were used for the pH
measurements.
Analytical balance
In order to weight accurate quantities a METTLER Analytical Balance AE 260 Delta
Range was used. It has a capacity of 200g / 60g delta range, a readability of 1mg / 0.1mg delta
range and a platform size of 80 mm.
Figure 2.4 – METTLER Analytical Balance AE 260 Delta Range.
27
FPLC
An AKTA FPLC™ system was used for FPLC purification. The system is equipped with
UV and conductivity detectors and an automated fraction collector. The system is controlled via
UNICORN software and a Method Wizard can be used for easy design of purification schemes.
A HiTrap SP FF 5 ml column from Amersham Biosciences was used (can bind 25 mg of
protein).
Figure 2.5 – AKTA FPLC™ system.
2.1.5. Analytical Techniques
SDS Page
Concept:
Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS Page) is a method used
to separate proteins according to their electrophoretic mobility, which is a function of the protein‟s
molecular weight. The protein solution to be analyzed is mixed with SDS, an anionic detergent
that denatures secondary and non-sulfide-linked tertiary structures, applying a negative charge to
each protein in proportion to its mass.
An electric field is applied across the gel, causing the negatively charged proteins to
migrate across the gel towards the anode. Depending on their size, each protein will move
differently through the gel matrix: smaller proteins will have traveled farther down the gel, while
larger ones will have remained closer to the point of origin. Therefore, proteins may be separated
approximately according to their size (and therefore, molecular weight).
28
Electrophoresis buffer: 125 mM Tris base (Trisma), 1.25 M glycine and 17.5 mM SDS
(Sodium dodecyl sulfate).
Loading Buffer (Denaturizing solution): 62.5 mM Tris HCl pH 6.8, SDS 4%, 2-
mercaptoethanol 5%, glycerol 20%, bromophenol blue 0.01%.
2 µl of AP were diluted in 180 µl H2O and then 20 µl of the diluted enzyme were added to
5 µl of the loading buffer (denaturating solution), boiled for 5 minutes and centrifuged.
20 μl of denaturated enzyme solution was put on a Bio-Rad Ready Gel 10%
polyacrylamide and run in a Bio-Rad System. To determine the molecular weight, a prestained
protein marker was also put on the SDS gel. It is common to run molecular markers of known
molecular weight in a separate lane in the gel, in order to calibrate the gel and determine the
weight of unknown proteins by comparing the distance traveled relative to the marker. For this
aim, the broad range prestained protein marker kit was used; it contains a mixture of purified
proteins covalently coupled to a blue dye that resolves to 8 bands of even intensity when
electrophoresed (table 2.2).
Table 2.2 – Prestained protein marker, broad range kit.
Protein Source MW (kDa)
MBP-b-galactosidase1 E-Coli 175.0
MBP-paramyosin1 E-Coli 83.0
Glutamic dehydrogenase Bovine liver 62.0
Aldolase Rabbit muscle 47.5
Triosephosphate isomerase Rabbit muscle 32.5
b-Lactoglobulin A Bovine milk 25.0
Lysozyme Chicken egg white 16.5
Aprotinin Bovine lung 6.5
1. MBP- maltose-binding protein MBP-b-galactosidase = fusion of MBP and b-galactosidase MBP-paramyosin = fusion of MBP and paramyosin
After the electrophoresis, the gel was stained with Coomassie Brilliant Blue R-250 for 30
minutes, allowing visualization of the separated proteins.
29
Bio-Rad Assay for determination of Protein concentration
Concept:
The Bio-Rad Protein Assay is based on the method of Bradford, a colorimetric assay
based on an absorbance shift in the dye Coomassie when the red form coomassie dye changes
into coomassie blue due to the binding to protein. The Bio-Rad Protein Assay Kit II was used in
which the acidic dye (Coomassie Brilliant Blue G-250 dye) was added to the protein solution and
absorbance measurements were taken at 595 nm on the Spectrophotometer.
The absorbance maximum for an acidic solution of Coomassie Brilliant Blue G-250 dye
shifts from 465 nm to 595 nm when binding to the protein occurs. The Coomassie blue dye binds
to primarily basic and aromatic amino acid residues, especially arginine.
Binding of the protein stabilizes the blue form of coomassie dye, thus the amount of
complex present in solution is a measure for the protein concentration by use of an absorbance
reading.
The increase of absorbance at 595 nm is proportional to the amount of bound dye, and
thus to the amount (concentration) of protein present in the sample.
Comparison with a standard curve for the bovine serum albumin standard protein, BSA
(1.4 mg/ml) allows the determination of protein concentration. Standard solutions were prepared
using known concentrations of BSA: 0.98, 3.92, 7.84, 11.76, 15.68, 17.64 and 19.60 µg/ml with a
total volume of 1 ml, using 980 µl of diluted Dye reagent (Dye: Water = 1:4).
The samples of unknown concentration of alkaline phosphatase were prepared by adding
20 μl of enzyme (2x and 4x diluted) to separate 1.5 ml eppendorf vials. To each eppendorf vial,
980 μl of diluted Dye Reagent (Dye: Water = 1:4) was added and mixed on a vortex. The samples
were incubated at room temperature for at least 5 minutes, but not more than one hour.
Absorbance was measured at 595 nm and compared with a previously obtained standard curve
from BSA. Each assay was performed in duplicate.
The standard curve was built plotting standards‟ concentration versus absorbance
measurement, as depicted in figure 2.6. After adding a linear trendline, its equation is used to
determine protein concentration according to the formula:
(1)
where:
C is the protein concentration in (g/ml)
Abs is the absorbance read
C(g /ml) Absq
m dilution in cuvette dilution factor
30
qrepresents the origin point of the linear trendline equation
m is the slope of the linear trendline equation
Figure 2.6 – Standard BSA curve for protein concentration determination.
Enzymatic Activity Assay (pNPP assay)
AP activity was checked on a weekly basis. It was determined spectrophotometrically at
405 nm at room temperature, using pNPP (p-Nitrophenyl phosphate) 100 mM as substrate for the
hydrolysis: pNPP is a good substrate for AP and its hydrolysis product, p-nitrophenol (pNP) is a
compound which absorbs at 405nm (figure 2.7).The pNPP solution was prepared fresh before
use and kept in the freezer at -20 °C.
O
NO2
PO-O
O-
Na+
Na+
+ H2O
NO2
OH
+ Pi
AP
pNPP pNP
Figure 2.7 – p-Nitrophenyl phosphate (pNPP) hydrolysis by alkaline phosphatase.
31
The buffer used in the assay was 1 M diethanolamine (DEA, pH 9.8) containing 0.5 mM
MgCl2.
The buffer used for enzyme dilutions and for the blank was 5 mM Tris HCl, pH 7.5, 5 mM
MgCl2 and 0.1 mM ZnCl2.
The reaction was carried in 1.1 mL cuvette containing 900μL buffer, 100µL pNPP solution
and 10μL enzyme against 900 µL of buffer, 100 µl pNPP and 10µl enzyme buffer as blank. The
absorbance was measured on the Spectrophotometer.
The absorbance change was recorded every half minute during 5 minutes, throughout
this time the mixture color turns from clear to yellow due to pNPP hydrolysis.
The alkaline phosphatase activity, using a certain dilution of enzyme, was calculated
according the formula:
Activity (U /ml) Abs
405nm
D
t (2)
where:
Abs is the absorbance read
405nm is the millimolar extinction coefficient of p-nitrophenol at 405 nm, which is 18.5 mM-
1cm
-1
D represents the dilution of the enzyme in the cuvette multiplied by the dilution factor
t is the time of the absorbance measurement
Unit Definition:
One unit will hydrolyze 1 µmole of p-nitrophenyl phosphate per minute at pH 9.8 and 20 °C.
Pi inhibition (pNPP assay)
The same activity assay was performed in the presence of different concentrations of
inorganic phosphate in the reaction mixture to determine if the enzyme is suffering from inhibition
by phosphate. The concentrations of substrate and buffer were 100 mM pNPP and 1 M DEA, 0.5
mM MgCl2 and 10, 25 and 100 mM of sodium phosphate buffer were present. The reactions
were performed at pH 8 and 25 °C in a 1.1 ml cuvette. The absorbance was measured on the
spectrophotometer every half minute during 5 minutes.
The alkaline phosphatase activity, using a certain dilution of enzyme, was calculated
according the formula described above (equation 2).
32
DHA phosphorylation
In order to evaluate the phosphorylating capacity of the enzyme AP a reaction was
carried out using dihydroxyacetone (DHA) as the substrate and pyrophosphate (PPi) as the
phosphate donor. The phosphorylation product is dihydroxyacetone phosphate (DHAP), which
cannot be detected spectrophotometrically. Therefore, a coupled assay was performed, using
reduced NADH in the presence of L-glycerol-3-phosphate dehydrogenase (G3PDH), producing L-
glycerol-3-phosphate (figure 2.8). NADH absorbs at 340 nm and its consumption in the second
step of the reaction equals the amount of phosphorylated product formed in the phosphorylation
(DHAP) using alkaline phosphatase.
The DHAP concentration was spectrophotometrically measured by the decrease of
absorbance of the NADH at 340 nm.
Figure 2.8 – Dihydroxyacetone phosphorylation and coupled assay with NADH.
Reaction mixture: 50 mM PPi, 100 mM DHA and the amount of enzyme ranged from 2,
4 and 6 U/ml. The reaction mixtures were set for different pH (7, 7.5, 8, 9 and 10) to create a pH
profile and determine the optimum pH for this reaction. The different pH values were obtained by
adding the required amount of HCl or NaOH to the PPi and DHA mixture until the desired pH was
reached.
Assay mixture: 0.16 mM NADH (some grains were added to the cuvette until the
absorbance reads 1.2), 100 mM tris/acetate pH 7.5, 1 U/ml G3PDH.
33
The reaction is initiated by adding the enzyme to the pyrophosphate and
dihydroxyacetone mixture and at that point the reaction is started (point zero). The reaction was
carried in the thermomixer at 32 °C. Every 10 minutes 980 µl of assay mixture were pipetted into
a 1.2 ml quartz cuvette and then 20 µl of reaction mixture were added. The mixture was
thoroughly mixed with a pipet (200 µl) and incubated for 3 minutes before reading the
absorbance.
The reaction was repeated in the presence of different concentrations of MgCl2 to the
reaction mixture in order to verify if its use improves the phosphorylation and to determine the
most advantageous Mg2+/PPi concentration ratio. The reaction was carried in the thermomixer at
32 °C and pH 8. The reaction mixture composition was the same as before with the exception of
1 mM (Mg2+/PPi = 1:50) and 25 mM MgCl2 added (Mg2+/PPi = 1:2), also in a total volume of 1 ml.
The concentration of dihydroxyacetone phosphate formed in the phosphorylation reaction
can be calculated according to the equation 3:
CDHAP Ai A f
NADH ,403nm
dilution in cuvette (3)
where:
Ai is the initial absorbance of the assay mixture before adding the reaction mixture
Af is the absorbance read every ten minutes
NADH,340nm is the millimolar extinction coefficient of NADH at 340 nm, which is 6.22 mM-
1cm
-1
PPi Hydrolysis
Pyrophosphate is the phosphate donor used in the cascade reactions mentioned in this
thesis, for that reason it‟s important to characterize the hydrolysis using the enzyme alkaline
phosphatase (figure 2.9).
-O P O
O
O-
APP
O-
O-
O
-O P
O-
O-
O
2
Figure 2.9 – Pyrophosphate hydrolysis by alkaline phosphatase.
34
The reaction was carried at pH 7 and 8 with various concentrations of pyrophosphate
(250, 100 and 50 mM) and a concentration of enzyme of 6 U/ml and the concentration of PPi was
measured using HPLC. 1 ml of PPi at different concentrations (pH 7 and pH 8 in parallel) was
added to 30 µl of AP 100x diluted and the reaction was carried in the thermomixer at 30 °C.
Every 30 minutes 10x diluted samples were prepared (20 µl reaction mixture in 180 µl
Milli-Q water) and used for HPLC analysis (4x diluted for 100 and 50 mM of PPi). Approximately
20 µl per sample were injected and the column was eluted at 35 °C with a flow rate of 0.4ml/min.
The total time of the run for each sample was 10 minutes. The eluent utilized in the HPLC
analysis was a 25mM H2SO4 solution.
The reaction was repeated adding different concentrations of MgCl2 to test if its use
improves the pyrophosphate hydrolysis and to establish the best Mg2+
/PPi concentration ratio.
The reaction was conducted at pH 8 and carried out during 2 days (~45 hours) in a thermomixer
at 30 °C; samples 4x diluted were taken every 30 minutes to HPLC analysis.
The reaction composition was 100 mM PPi, 6 U/ml AP and various concentrations of
MgCl2: 100 mM (Mg2+/PPi = 1:1), 66 mM (Mg2+/PPi = 2:3) and 50 mM (Mg2+/PPi = 1:2), in a total
volume of 1 ml.
DHA cascade reaction
This one-pot cascade reaction starts with the dihydroxyacetone (DHA) phosphorylation
by AP and it‟s coupling with the aldehyde propanal by the aldolase RAMA. The resulting aldol-
product is then dephosphorylated by the alkaline phosphatase already present in the reaction
leading to the formation of the non-natural carbohydrate, 5,6-dideoxy-D-threo-hex-2-ulose (figure
2.10). The reaction was carried out at pH 7 and pH 8 in the thermomixer at 30 °C. The cascade
mixture was as follows: 100 mM PPi, 500 mM DHA, 100 mM propanal, 6 U/ml of AP and 6 U/ml
of RAMA in a total volume of 1 ml.
The reaction was started by adding the alkaline phosphatase to the cascade mixture and
20 µl samples were taken every hour and diluted 10x with Milli-Q water to HPLC analysis.
Approximately 20 µl per sample were injected and the column was eluted at 35 °C with a flow
rate of 0.4ml/min. The total time of the run for each sample was about 25 minutes.
The reaction was repeated using different concentrations of MgCl2 to attest if its use
improves the reaction rate and to establish the optimal Mg2+
/PPi concentration ratio. The cascade
reaction was conducted at pH 8 and carried out during 24 h in a thermomixer at 30 °C. The
cascade mix composition was the same as before and the amount of MgCl2 added was 100 mM
(Mg2+/PPi = 1:1) and 50 mM (Mg2+/PPi = 1:2), also in a total volume of 1 ml.
35
Figure 2.10 – One-pot cascade reaction starting from DHA.
Immobilization of Alkaline Phosphatase in different supports
Immobilization on Immobeads-150
For the immobilization on Immobeads-150, 500 µl of a 30 mM potassium phosphate pH
8, 0.5 mM MgCl2 buffer were added to a 1.5 ml eppendorf vial containing 10 mg (dry weight) of
support. After gentle mixing, 100 µl of 100x diluted alkaline phosphatase stock solution were
added (~33 U/ml), the eppendorf vial was immediately placed on the rotator at medium rotation
speed and the reaction was started.
The immobilization was carried for 24 hours at 20 °C with 5 μl samples of the supernatant
being collected at 0, 1, 2, 3, 4, 5, 6 and 24 hours. The samples were diluted in 20 µl enzyme
buffer and 10 μl of each sample were immediately utilized for an activity assay.
Immobilization on Sepabeads EC-EP
For the immobilization on Sepabeads EC-EP, 500 µl of a 30 mM potassium phosphate
pH 8, 0.5 mM MgCl2 buffer were added to a 1.5 ml eppendorf vial containing 25 mg (wet beads)
of support. After gentle mixing, 100 µl of 100x diluted alkaline phosphatase stock solution were
added (~33 U/ml), the eppendorf vial was immediately placed on the rotator at medium rotation
speed and the reaction was started.
36
The immobilization was carried for 24 hours at 20 °C with 5 μl samples of the supernatant
being collected at 0, 1, 2, 3, 4, 5, 6 and 24 hours. The samples were diluted in 20 µl enzyme
buffer and 10 μl of each sample were immediately utilized for an activity assay.
Immobilization on Sepabeads EC-HA, Aldehyde activated
For the immobilization on Sepabeads EC-HA, 500 µl of a 30 mM potassium phosphate
pH 8, 0.5 mM MgCl2 buffer were added to a 1.5 ml eppendorf vial containing 25 mg (~50 µl beads
in fluid) of support. After gentle mixing, 100 µl of 100x diluted alkaline phosphatase stock solution
were added (~33 U/ml), the eppendorf vial was immediately placed on the rotator at medium
rotation speed and the reaction was started.
The immobilization was carried for 24 hours at 20 °C with 5 μl samples of the supernatant
being collected at 0, 1, 2, 3, 4, 5, 6 and 24 hours. The samples were diluted in 20 µl enzyme
buffer and 10 μl of each sample were immediately utilized for an activity assay.
DHA cascade reaction using immobilized AP
The DHA cascade reaction was repeated with the immobilized AP in the Sepabeads-EC-
HA support and also with the immobilized PhoN-Sf on Immobeads-150 to serve as a comparison.
The reactions were carried out in the thermomixer at 25 °C at two different pH values: pH
8 with 20 U/ml of immobilized AP (50 µl of Sepabeads EC-HA containing the enzyme) and pH 6
with 1 U/ml of immobilized PhoN-Sf (25 µl of Immobeads-150 containing the enzyme). The
cascade mixture was as follows: 100 mM PPi, 500 mM DHA, 100 mM propanal and 6 U/ml of
RAMA in a total volume of 1 ml.
The reaction was started when the alkaline phosphatase/PhoN-Sf were added to the
cascade mixture and 20 µl samples were taken every hour and diluted 10x with Milli-Q water and
analyzed by HPLC. The total time of the run for each sample was about 25 minutes.
Glycerol phosphorylation
As studied before using DHA, the phosphorylating capacity of the enzyme AP was tested
using this time glycerol as the substrate and pyrophosphate (PPi) as the phosphate donor. The
phosphorylation product is a racemic mixture of L/D glycerol-3-phosphate. None of the
compounds present in this reaction absorbs light in the UV region; therefore, a coupled assay
was performed, adding disodium salt of NAD+ and hydrazine in the presence of L-glycerol-3-
phosphate dehydrogenase (G3PDH), producing L-glycerol-3-phosphate (figure 2.11). The
37
enzyme L-glycerol-3-phosphate dehydrogenase (G3PDH), as the name indicates, only acts on
the L enantiomer of the racemic mixture of glycerophosphate formed, thus only the L enantiomer
is consumed in the coupled assay. NADH absorbs light at 340 nm and its formation in the
coupled assay equals the amount of the L-phosphorylated product formed in the phosphorylation
reaction.
The L-glycerol-3-phosphate concentration was spectrophotometrically measured by the
increase in absorbance of the NADH at 340 nm.
Figure 2.11 – Glycerol phosphorylation and coupled assay with NAD+.
Reaction mixture: 50 mM PPi pH 8, 100 mM glycerol and 6 U/ml of soluble alkaline
phosphatase.
Assay mixture: 450 mM glycine, 274 mM hydrazine, 2.4 mM EDTA pH 9.5, 20 U/ml
G3PDH and 2.5 mM NAD+.
The reaction is initiated when the enzyme is added to the pyrophosphate and glycerol
mixture and at that point the reaction is started (point zero). The reaction was carried in the
thermomixer at 30 °C. Every 10 minutes 980 µl of assay mixture were pipetted into a 1.2 ml
quartz cuvette and then 20 µl of reaction mixture were added. The mixture was thoroughly mixed
with a pipet (200 µl) and incubated for 5 minutes before reading the absorbance.
The concentration of L-glycerol-3-phosphate formed in the phosphorylation reaction can
be calculated according to the equation:
38
CLGlycerol3phosphateAbs
NADH ,403nm
dilution in cuvette (4)
where:
Abs is the absorbance read every ten minutes
NADH,340nm is the millimolar extinction coefficient of NADH at 340 nm, which is 6.22 mM-
1cm
-1
Glycerol cascade reaction
In this one-pot cascade reaction glycerol is phosphorylated using PPi as a phosphate
donor and in the presence of AP and is then oxidized by glycerol-3-phosphate oxidase, GPO, (in
combination with catalase) to form DHAP. DHAP is then coupled to propanal by the aldolase
RAMA and the resulting aldol-product is then dephosphorylated by the alkaline phosphatase
already present in the reaction leading to the formation of a non-natural carbohydrate, 5,6-
dideoxy-D-threo-hex-2-ulose (figure 2.12). The reaction was carried out at pH 8 in the
thermomixer at 25 °C in 20 U/ml of immobilized AP (50 µl of Sepabeads EC-HA containing the
enzyme). The cascade mixture was as follows: 100 mM PPi, 500 mM glycerol, 100 mM propanal,
10 U/ml of catalase, 6 U/ml of RAMA and 50 U/ml GPO in a total volume of 1 ml.
The reaction was started when the cascade mixture was added to the immobilized AP
and 20 µl samples were taken every hour and diluted 10x with Milli-Q water for HPLC analysis.
Approximately 20 µl per sample were injected and the column was eluted at 35 °C with a flow
rate of 0.4ml/min. The total time of the run for each sample was about 25 minutes.
The reaction was performed in parallel with another using an acid phosphatase, PhoN-Sf
to serve as a control. The cascade reaction was conducted at pH 6 and carried out in a
thermomixer at 25 °C. The cascade mix composition was the same as before and was added to
1 U/ml of immobilized PhoN-Sf (25 µl of Immobeads-150 containing the enzyme).
39
Figure 2.12 – One-pot cascade reaction starting from glycerol.
2.2. Production and purification of recombinant acid phosphatase from Salmonella enterica (PhoN-Se) expressed in E. coli BL21 (DE3) with pET23b plasmid
2.2.1. Protein expression overview
The enteric bacterium Escherichia coli is one of the most studied prokaryotic organisms
and is widely used as a shuttle host for genetic manipulations as well as for the industrial
production of proteins of therapeutic or commercial interest. Although chromosomal expression
systems have been described (Peredelchuk and Bennett, 1997; Olson et al., 1998), plasmid-
based expression remains the preferred means of producing recombinant proteins in E. coli71.
As various current expression plasmids require special mutant strains as hosts,
fermentation protocols have been developed for strains of E. coli, which differ considerably from
each other71. PhoN-Se was expressed in expression host E. coli BL21 (DE3) cells, which is a
strain that provides significantly high levels of recombinant protein expression and significantly
reduced levels of background protein levels, and it‟s also well-known and established in
laboratory handling.
40
An expression vector is a plasmid used for genetic engineering and usually contains an
origin of replication (ori), an antibiotic resistance marker and an expression cassette for regulated
transcription and translation of a target gene71.
Gene expression occurs in two major stages (figure 2.13). The first is transcription, in
which the gene is copied to produce an RNA molecule (a primary transcript) with essentially the
same sequence as the gene. Most human genes are divided into exons and introns, and only the
exons carry information required for protein synthesis. Most primary transcripts are therefore
processed by splicing to remove intron sequences and generate a mature transcript or
messenger RNA (mRNA) that only contains exons.
The second stage is protein synthesis. This stage is also known as translation and is so
called because there is no direct correspondence between the nucleotide sequence in DNA (and
RNA) and the sequence of amino acids in the protein. In fact, three nucleotides are required to
specify one amino acid. The chain of amino acids must fold up to generate the final tertiary
structure of the protein72.
Figure 2.13 - Gene structure and gene expression in higher organisms73.
For the expression of PhoN-Se the pET 26b vector was used. This system uses IPTG, Isopropyl β-D-1-thiogalactopyranoside, as an inducer. This compound is used as a molecular
mimic of allolactose, a lactose metabolite that triggers transcription of the lac operon. Unlike
allolactose, the sulfur atom present in IPTG creates a chemical bond, which is non-hydrolysable
41
by the cell, preventing the cell from “eating up”, or degrading the inductant; therefore the IPTG
concentration remains constant. IPTG induces the transcription of the gene coding for beta-
galactosidase, a hydrolase enzyme that catalyzes the hydrolysis of β-galactosides into
monosaccharides. In presence of IPTG, expression of pET23b is turned on while the absence of
inducer produces very low levels of transcription from pET23b.
The stable maintenance of plasmids is usually achieved by supplementing the growth
medium with antibiotics that are inactivated by plasmid-encoded resistance genes. The most
common markers are ampicillin, chloramphenicol, kanamycin and spectinomycin resistances71.
This plasmid contains a kanamycin resistance gene, which allows the selection of the plasmid in
E.coli.
For this project the expression, purification and isolation of new mutants of acid
phosphatase from Salmonella enterica ser. Typhimurium: V78Y, V78L and V78H were performed
as subsequently described.
2.2.2. Solutions
50 mg/ml kanamycin 1000x stock, filtered sterile and stored at -20 °C; final concentration
for induction is 50 µg/ml.
LB medium: 10 g/L peptone-tryptone, 5 g/l yeast extract, 10 g/l NaCl; kanamycin was
added right before starting the growth; autoclaving at 121 °C for 20 minutes.
1 M IPTG, Isopropyl β-D-1-thiogalactopyranoside, stored at -20 °C.
1 M Na-acetate buffer pH 6.
FPLC buffer A: 20 mM Na-acetate pH 6.
FPLC buffer B: 20 mM Na-acetate pH6, 1 M NaCl.
FPLC buffers should be filtered and prepared with Milli-Q water.
2.2.3. Protocol for expression and purification
1st Day
The inoculation was started at the end of the day. The -80 °C glycerol stocks of BL21
(DE3) pET26b-PhoN-Se cells for each mutant were taken of the freezer with the concern of not
liquefying the cells. The rest of the tasks for this day were performed in a laminar flow hood to
provide the required biological safety. With an inoculation loop, heated in a flame and pre cooled
42
in ethanol, some material from the glycerol stock were scraped and then spread across one
quadrant of the agar plate containing a growth medium which has been sterilized in an autoclave.
This introduces a solution of the bacteria to a substrate, which provides them nutrients. The loop
was re-sterilized and dragged across the inoculated quadrant of the streak plate. This is done to
collect some bacteria on the loop. The loop is spread around another fourth of the plate as done
for the previous step. The loop is sterilized and the procedure is repeated. Each time the loop
gathers fewer and fewer bacteria until it gather just one single bacterial cell that can grow into a
colony.
The agar plate contained kanamycin, an antibiotic used as a selective agent to isolate
bacteria that have taken up genes coupled to a gene coding for kanamycin resistance. Bacteria
that have been transformed with a plasmid containing the kanamycin resistance gene are plated
on kanamycin containing agar and only the bacteria that have successfully taken up the
kanamycin resistance gene become resistant and will grow under these conditions.
The plates were incubated a 37 °C overnight.
2nd Day
The plates with single colonies for each mutant were stored at 4 °C (maximum storage
time of the plates at this temperature is 2 weeks).
At the end of the day, also working under the laminar flow cabinet, a single colony was
chosen from each plate and put in 10 ml of LB containing 50 µg/ml of kanamycin in a 50 ml
Greiner tube for better aeration (two Greiner tubes were prepared for each mutant). The Greiner
tubes were incubated with vigorous shaking at 37 °C overnight.
Erlenmeyer flasks were sterilized in the autoclave at 121°C for 20 minutes. One 2.5 L
flask is used for 0.5 L of LB for good aeration. Since for each mutant 2 L of LB were prepared, the
total of Erlenmeyer flasks was 4 for each mutant, therefore 12 flasks to sterilize.
Autoclaving at 121°C for 20 minutes was also performed for the LB medium. 0.5 L in a 1
L bottle. Since 2 L of LB were prepared for each mutant, 6 L were necessary, therefore 12 1L
bottles were sterilized.
3rd Day
5 ml from the overnight culture were inoculated into 0.5 L of LB with 50 mg/ml kanamycin
in a 2.5 L Erlenmeyer flask. The flasks were incubated with vigorous shaking at 37 °C until the
optical density OD600>0.4 (it takes around 3-4 hours). When OD600 reaches 0.4, 1 mM IPTG was
added to each Erlenmeyer flask to induce PhoN-Se expression. The shakers were cooled down
to 25 °C and the flasks were incubated overnight.
43
4th Day
The cells were collected by centrifuging the content of each Erlenmeyer at 6000 rpm for
15 minutes at 5 °C. The pellet obtained after centrifuging was resuspended in 20 mM Na acetate
pH 6 buffer (pellet from 0.5 L LB can be resuspended in 5 mL of the buffer) and kept on ice.
The resuspended pellets were sonicated to break the cells membranes and free the
protein into the solution. For the sonication the sonicator probe was cleaned with 20% ethanol.
Sonication during 45 seconds and 15 seconds break was performed 4 times. During the 15
second break the solution was kept on ice.
The sonicated solutions were centrifuged at 13000 rpm for 30 minutes at 5 °C and the
supernatant was decanted into a clean tube and checked for activity with a pNPP activity assay in
a micro titer plate.
Ammonium sulfate precipitation was performed to each mutant solution. This is a method
used to purify proteins by altering their solubility since at high salt concentrations (high ionic
strength) the solubility of the protein begins to decrease and at sufficient high ionic strength the
protein will be almost completely precipitated from the solution – salting out. Since proteins differ
noticeably in their solubility at high ionic strength, salting-out is a very useful procedure to assist
in the purification of a given protein. The amount of ammonium sulfate to add was determined
from a published nomogram. The final concentration of ammonium sulfate in the solution was of
20%: 10.6 g in 100 ml. According to the volumes of each mutant solution different amounts of
ammonium sulfate were added as shown in table 2.3.
Table 2.3 – Ammonium sulfate added to each mutant solution for precipitation of contaminant proteins.
Mutants Sample volume (ml) Ammonium sulfate added (g)
Y 20 2.12
L 19 2.01
H 15 1.59
The ammonium sulfate was added to the mutant solution in 50 ml beakers and stirred for
1 hour on ice.
After ammonium sulfate precipitation the solutions were centrifuged at 13000 rpm for 30
minutes at 5 °C and the supernatant was dialyzed with 20 mM Na acetate pH 6 overnight at 4 °C.
44
5th Day
The dialysis buffer was changed and left for one more hour. The protein solutions were
centrifuged at 13000 rpm for 30 minutes at 5 °C.
Further purification was done on a FPLC using a HiTrap SP FF 5ml 30B column. The
chromatographic bed is composed by gel beads inside the column and the sample is introduced
into the superloop and carried into the column by the flowing solvent. As a result of different
components adhering to or diffusing into the gel, the sample mixture gets separated. This HiTrap
SP FF 5ml column is an ionic exchange column therefore the macromolecules are separated
based on their charge distribution.
The pump system and the column were washed to get rid of 20% ethanol and to change
to the correct buffers. The protein solutions were syringe-filtered with a MDI-syringe-filter type
SY25PG-s/0.2µm/25mm and loaded into the FPLC superloop. The superloop with protein
solution was connected to the column. First the column was equilibrated with 25 mL of Buffer A
to get rid of salt or ethanol. Then the superloop was loaded into the column with buffer A, where
the protein was bound to the chromatographic bed. The column was eluted with buffer B with a
gradient of 1 M NaCl until all proteins were unbound. After using the system it should be stored in
20% Ethanol, therefore the pumps and the column must be washed with ethanol 20% and left like
this.
When the run was finished, the fractions were tested for PhoN-Se activity in a microtiter
plate and the active fractions were collected and dialyzed against 20 mM Na-acetate pH 6 to get
rid of the NaCl.
6th Day
The samples were concentrated in Amicon Centriplus tubes from Millipore with a 10 kDa
filter (the molecular weight of PhoN-Se is 27 kDa, so the protein doesn‟t pass in the filter). The
tubes were centrifuged at 3000 rcf for 2-3 hours at 5 °C.
Protein concentration was determined by the Bio-Rad assay described above.
Phosphatase activity was determined through an assay for acid phosphatase.
45
2.2.4. Analytical techniques
Assay for acid phosphatase
Assay mixture: 100 mM Na-acetate pH 6, 10 mM pNPP.
The reaction is started by addition of enzyme to the assay mixture (10 µl of enzyme in
490 µl of assay mix). The reaction is incubated for 3 minutes, or less depending on how fast the
solution turns yellow, and then quenched by addition of 0.5 M of NaOH to bring to pH 12. The
absorbance is monitored at 410 nm using the extinction coefficient for pNPP of 16.6 mM-1
cm-1
.
The mutants‟ activity, using a certain dilution of enzyme, was calculated according the
equation 2, described in section 2.1.5 – Enzymatic activity assay.
DHA cascade reaction
The reaction was carried out at pH 6 in the thermomixer at 30 °C. The cascade mixture
was as follows: 250 mM PPi, 500 mM DHA, 100 mM propanal, 20 mM Na-acetate pH 6 and 6
U/ml of RAMA in a total volume of 0.5 ml. To this mixture 1 µM of PhoN-Se WT, V78 L and V78 Y
were added separately.
The reaction was started when the enzyme was added to the cascade mixture and 20 µl
samples were taken every hour and diluted 10x with Milli-Q water to HPLC analysis.
Approximately 20 µl per sample were injected and the column was eluted at 35 °C with a flow
rate of 0.4ml/min. The total time of the run for each sample was about 25 minutes.
The reactions were repeated for the V78 L and V78 Y mutants adding different
concentrations of enzyme: 0.5 µM and 2 µM (also performed for the WT enzyme).
46
47
3. Results and Discussion
3.1. Alkaline Phosphatase
3.1.1. Characterization of the alkaline phosphatase stock solution
The precise protein concentration and activity in the calf intestine alkaline phosphatase
stock solution were determined by a Bio-Rad and activity assays respectively. The Bio-Rad assay
gave a protein concentration of 15 mg/ml. This value together with the soluble enzymatic activity
assay, which reported an activity of 20000 U/ml, gives a specific activity of 1333 U/mg-protein in
the alkaline phosphatase stock solution. One unit is defined as one µmol of p-nitrophenyl
phosphate hydrolysed per minute at pH 9.8 and 20 °C.
According to the values provided by the supplier; protein concentration of 19.5 mg/ml and
enzyme activity of 109414 U/ml, the specific activity is of 5611 U/mg. The specific activity
obtained here is 24% of the one provided by the supplier. A possible explanation for this low
value can be that the enzyme has suffered from denaturation due to a long storage period. An
SDS-PAGE gel of the original alkaline phosphatase solution was prepared, as described above,
in order to verify its purity.
Figure 3.1 - SDS-PAGE gel of the original alkaline phosphatase solution. Lane (1) contains the molecular
markers identified with the corresponding molecular weights in kiloDaltons, lane (2) corresponds to a 2 µg
solution of AP while lane (3) corresponds to a 2.5 µg solution.
(1) (2) (3)
175.0
83.0 62.0 47.5
32.5 25.0
16.5
6.5
48
Figure 3.1 shows a strong band with a molecular mass slightly below 83 kDa. This band
should correspond to the monomeric unit of alkaline phosphatase because of the presence of the
denaturizing agent that breaks the secondary and non-sulfide-linked tertiary structure of the
protein. The value obtained is in accordance with the range of values reported for the molecular
weight of monomers of other mammalian alkaline phosphatases, 72-150 kDa51, 52. The fact that
no other bands are found on the AP‟s lanes demonstrates that the enzyme is fully dissociated in
the presence of the reducing agent.
It is then possible to conclude that the supplied alkaline phosphatase solution is highly
pure, having a monomeric molecular weight of approximately 80 kDa and a total molecular weight
of around 160 kDa.
3.1.2. Analytical techniques
DHA Phosphorylation using soluble AP
In order to optimize the DHAP formation two parameters were analysed: the alkaline
phosphatase concentration and dependency on pH.
The dihydroxyacetone phosphorylation, using 50 mM PPi as a phosphate donor and 100
mM of DHA as substrate, was performed using different concentrations of soluble alkaline
phosphatase: 2, 4 and 6 U/ml and the reaction was followed for about 140 minutes at pH 9 and
32°C. Time-dependent optima are observed since, after the consumption of the PPi, DHAP will
be dephosphorylated by the phosphatase.
In figure 3.2 the effects of the phosphatase concentration in the formation of DHAP are
presented.
49
Figure 3.2 – DHAP formation over time using different AP concentrations. Reaction mixture contained DHA
(100 mM), PPi (50 mM), and AP (2, 4 and 6 U/ml) in 1ml at pH 9.
As expected an increase in the amount of enzyme positively affects the conversion of
DHA in DHAP, although after 40 minutes, the results obtained using 4 U/ml of alkaline
phosphatase seem to be more favourable than the ones obtained using 6 U/ml. This lower
product formation obtained with 6 U/ml could be due to the fact that this experiment was
performed in a different day, whereas the other two reactions (2 and 4 U/ml) were performed in
parallel. This can cause some disparity in the composition of the assay mixtures and/or the
reaction mixtures, therefore leading to results that cannot be compared directly.
In any case, for the subsequently reactions performed, the amount of alkaline
phosphatase used was 6 U/ml since no great difference is noticed in the concentrations of DHAP
formed: 2.2 mM using 4 U/ml and 1.9 mM using 6 U/ml after 130 minutes of incubation (figure
3.2).
Figure 3.3 shows the pH dependence of the alkaline phosphatase phosphorylating
activity.
50
Figure 3.3 – pH dependency of DHA phosphorylation. Reaction mixtures contained DHA (100 mM), PPi (50
mM), AP (6 U/ml) in 1 ml at pH 7, 7.5, 8, 9 and 10. The pH was set by addition of HCl or NaOH to the PPi,
DHA mixture until the desired value was reached.
As depicted in the figure above, the highest DHAP concentration obtained, 2.8 mM, was
reached at pH 7.5 after 110 min of incubation. Increasing the pH increased the initial rate of
DHAP formation, but the amount of DHAP formed at pH 9 and 10 was considerably lower. The
pH optimum for the DHA phosphorylation is clearly between pH 7.5 and pH 8. Combining yield
and reaction time (2.6 mM of DHAP after 80 minutes), pH 8 was taken as a starting point for
further optimization.
The results obtained from these optimizations are in accordance to previous studies
performed with acid phosphatases, PhoN-Se/Sf 37, 43 in which approximately 3 mM of DHAP were
obtained at pH 4, using the same concentrations of PPi and DHA.
Pyrophosphate (PPi) hydrolysis using soluble AP
It‟s very important to establish the optimum conditions for the hydrolysis of the phosphate
donor, in this case PPi, since it‟s an essential step for the subsequent reactions involved in the
cascades. The PPi hydrolysis was performed using different concentrations of PPi: 50, 100 and
250 mM, and the reaction was carried out for about 4 hours at pH 7 and pH 8 and 30°C. Samples
were taken every half an hour for HPLC analysis.
A reliable way of analysing the results is determining the decrease in the peak area of
PPi, as well as the increase in the concentration of Pi, through the course of the reaction since
51
the peak areas obtained in the chromatogram are proportional to the concentration of the
compounds.
Table 3.1 - Consumption of PPi and formation of Pi in mM for different PPi concentrations.
PPi left (mM) Pi formed (mM)
PPi (mM) pH 7 pH 8 pH 7 pH 8
50 0.8 0,5 49,2 49,5
100 67 35 33 65
250 230 200 20 50
According to our measurements the correlation between the peak area values and the
concentrations of PPi is of one to one, taking into account the dilution of the injected samples. In
this case for the hydrolysis of 250 mM of PPi the samples were 10x diluted whereas for 100 and
50 mM PPi the samples were 4x diluted.
As observed in table 3.1 the best results were obtained using lower concentrations of
PPi: with 50 mM PPi the reaction is practically complete after the 4 hours and there are no
differences in the choice of pH. When 100 mM PPi is used as initial concentration it is clear that
better results are obtained at pH 8 compared to pH 7 although the reaction does not lead to
completion in either cases. The rates of reaction seem to be slower than with 50 mM PPi, which
is understandable due to the fact that there is more reagent to hydrolyse. When 250 mM PPi is
used (same amount as utilized in the cascade reactions using acid phosphatases) the reaction is
extremely slow and very little PPi is hydrolysed and the reaction tends to reach a steady state,
with no further hydrolysis. These results disagree with the ones observed when acid
phosphatases are used in the phosphorylation of DHA37, 43. In that case the increase of PPi
concentration caused a further increase in DHAP formation, suggesting that the hydrolysis of PPi
was also improved by this increase.
The choice of PPi concentration for further studies was 100 mM since that‟s a convenient
amount to use in the cascade reactions; 50 mM gave the best results but its concentration is too
low to obtain the desired product concentrations, and 250 mM of PPi seemed to inhibit the
enzyme´s activity to some extent.
52
DHA cascade reaction using soluble AP
The reaction mixture for the dihydroxyacetone cascade consisted of 100 mM PPi, 500
mM DHA, 100 mM propanal, 6U/ml AP 6nd U/ml RAMA and was carried in 1ml eppendorfs vials
at pH 8 and 30 °C.
Samples were taken every hour and injected in the HPLC for peak area readings. Figure
3.4 illustrates the time course of the reaction for 24 hours, showing the product formation, and the
phosphorylated product and phosphate present in the reaction mixture, in terms of peak areas. It
was not possible to quantify the peak areas since for the native compounds no reference was
available.
Figure 3.4 – Time course for the DHA cascade reaction using soluble AP. Reaction mixtures contained
DHA (500 mM), PPi (100 mM), propanal (100 mM), AP (6 U/ml) and RAMA (6 U/ml),in 1 ml at pH 8.
After 24 hours the peak area of the final product was around 4, which by our estimations
corresponds to around 7 mM (7% conversion, calculated from the initial concentration of
propanal). However still a considerable amount of the phosphorylated product is present which
does not decrease. The phosphate peak area increases for the first 4-5 hours but then seems to
stagnate at a peak area of around 2.5.
These results are not very satisfactory since the reaction rate is very slow. Also the
product formation is poor (with acid phosphatase PhoN-Sf the final product concentration is
around 60 mM) 43 and the presence of phosphorylated product indicates that the reaction isn‟t
finished, as is also indicated by the very low Pi peak area. At the end of the reaction, starting from
100 mM PPi and using 10x-diluted samples, the peak areas should be around 10 for Pi and 0 for
PPi, showing that the hydrolysis was complete. In this case the reaction stops after only 25% of
53
PPi has been consumed.
PPi inhibition
A possible explanation for these unsatisfactory results is that the enzyme is suffering from
some type of inhibition. The first assumption was that AP was being inhibited by the substrate
PPi. Several publications supported this fact74, 75 and have indicated that the addition of
magnesium ions activates the pyrophosphatase activity of the enzyme, therefore possibly
overcoming the inhibitions problems caused by excess of PPi and enhancing the enzyme activity
in the cascade reaction.
For this purpose the reactions described above: DHA phosphorylation, PPi hydrolysis
and the DHA cascade reaction were performed in the same optimized conditions but with
different Mg2+
/PPi concentration ratios. The results obtained from each experiment were very
similar and just from a demonstrative point of view table 3.2 shows the results obtained in the
DHA cascade reaction with and without addition of MgCl2.
Table 3.2 – Comparison of obtained peak areas for the DHA cascade reaction with addition of different
concentrations of MgCl2: 100 mM (1:1) and 50 mM (1:2).
Peak Area
No Mg2+ Mg2+/PPi, 1:1 Mg2+/PPi, 1:2
Product 4.08 0.42 2.83
Phosphorylated Product 2.19 1.55 3.98
Phosphate 2.69 0.74 1.99
As observed in table 3.2 and confirmed by the other experiments, DHA phosphorylation
and PPi hydrolysis (see Appendix A) the addition of Mg2+
didn‟t improve the reaction in any of the
concentration ratios employed, the reaction without magnesium even leads to higher peak areas
of the product. When the concentration ratio used was Mg2+
/PPi = 1:1 the enzyme is clearly
affected by the excess of magnesium and the PPi hydrolysis is practically non-existing.
According to previous studies74, 75 the magnesium forms complex ions with PPi: MgPPi2-
,
and Mg2PPi and it appears that the enzyme is more active towards the complex ion MgPPi2-.
Therefore the presence of magnesium is necessary for pyrophosphatase activity but at high Mg2+
concentrations the complex ion Mg2PPi is formed which inhibits the enzyme due to depletion of
the true substrate.
54
These results show that the presence of magnesium on the enzyme buffer appears to be
sufficient to activate the pyrophosphatase activity and there is no need to add further amounts of
this ion since it inhibits PPi hydrolysis. However the low activity displayed by the enzyme when
high concentrations of PPi were used is still to be solved, as apparently the problem does not lie
in PPi inhibition.
Pi inhibition
The second hypothesis was that AP was being inhibited by inorganic phosphate, also a
known inhibitor for this enzyme as described in several articles76. For that matter a normal pNPP
activity assay was carried (100 mM pNPP, 1 M diethanolamine, DEA, 0.5 mM MgCl2 buffer, 6
U/ml AP) and different concentrations of phosphate: 10, 25 and 100 mM were prepared. Activity
calculations were performed as described above in section 2.1.5 – Enzymatic activity assay
(equation 2) and the results are shown in figure 3.5.
Figure 3.5 – Alkaline phosphatase activity in the presence of different Pi concentrations. Reaction mixtures
contained pNPP (100 mM), a DEA (1 M) and MgCl2 (0.5 mM) buffer pH 8, AP (5000x diluted) in 1.01 ml.
Figure 3.5 clearly shows that the activity of the enzyme is strongly affected by the
addition of Pi. The addition of only 10 mM of Pi decreases the enzyme activity almost 15%, this
loss in activity increasing to 60% when 25 mM Pi is used and the enzyme becomes completely
inhibited by 100 mM Pi.
This fact explains the poor results obtained in the cascade reaction and in the PPi
hydrolysis using high concentrations of PPi. As PPi is being hydrolyzed the accumulation of
55
phosphate in the reaction mixture causes inhibition of the alkaline phosphatase and thus the
cascade reaction does not lead to completion.
Immobilization of alkaline phosphatase
In an attempt to overcome the inhibition problems caused by Pi as well as to ensure the
preservation of the catalytic activity and allow its repeated or continuous use, the next step was to
immobilize the enzyme.
Three supports were tested for AP immobilization: Immobeads-150, Sepabeads EC-EP
and Sepabeads EC-HA. The first two have an epoxy functional group and the Sepabeads EC-HA
have an amino functional group which is aldehyde activated. The covalent binding of the enzyme
to the supports was followed for 24 hours by an activity assay on the supernatant (figure 3.6).
When the activity of the supernatant reaches zero it means the enzyme is completely bound to
the beads.
Figure 3.6 – Alkaline phosphatase activity in the supernatant during immobilization on different supports.
Reaction mixture contained pNPP (100 mM), a DEA (1 M) and MgCl2 (0.5 mM) buffer pH 8 and 10 µl of
supernatant 5x diluted in immobilization buffer: KPI (30 mM), MgCl2 (0.5 mM) in 1.01 ml.
The immobilization of AP in the three different supports differed. Only the Sepabeads EC-
HA (figure 3.6) showed no AP activity in the supernatant after 4 hours indicating that all enzyme
is bound to the support. Both Immobeads-150 and Sepabeads EC-EP showed very high values
of remaining activity in the supernatant even after 24 hours of incubation. This may imply that the
amino groups of the enzyme have a better interaction with the aldehyde activated support instead
56
of the epoxy groups of the other supports. Therefore Sepabeads EC-HA was the support chosen
for the following reactions.
The fact that only the Sepabeads EC-HA support gave a successful immobilization of AP
may be explained by it being the only carrier with a relatively long and branched spacer. The
properties of spacers, such as length, hydrophobicity/hydrophilicity and charged/neutral character
can exert a striking influence on binding capability, retention of activity, stability and catalytic
performance because of the higher molecular mobility in the presence of spacers between the
enzyme and the carrier.
Previous studies70 showed the effect of binding functionality on retention of the activity,
using several types of carriers such as glass, agarose-based carrier, like Sepharose, and
synthetic polymeric carriers such as Eupergit-C (epoxy synthetic carriers), which had different
activating agents, for the immobilization of AP70. On the basis of activity retention the best
supports for AP immobilization were Eupergit-C and CNBr-Sepharose (95% and 92%,
respectively at pH 9). Other kind of supports were also described: corn grits (EURA-MA) 47 the
main component of which is cellulose and Sepharose-4B, a macroporous cellulose and chitosan-
based carrier77
, leadind to 30% yield of immobilization for EURA-MA and around 60% for
Sepharose-4B.
Finally, it should be noted that both Sepabeads and Immobeads supports were easily
handled, leading to minimal support loss during the immobilization of AP in these supports.
DHA cascade reaction on immobilized AP
After successful immobilization of alkaline phosphatase on Sepabeads EC-HA the DHA
cascade was repeated. The cascade mixture was the same as before and the reaction was
carried at 25°C pH 6 and 8 in a batch system. For pH 8 20 U/ml of immobilized AP (~50 µl) were
used and for pH 6, 1 U/ml of acid phosphatase PhoN-Sf, immobilized on Immobeads-150 (~25 µl)
was used to serve as a control. Figures 3.7, 3.8 and 3.9 show the time courses for the formation
of product, phosphorylated product and inorganic phosphate, Pi, for both immobilized alkaline
phosphatase and immobilized PhoN-Sf.
57
Figure 3.7, 3.8 and 3.9 – Time course for the formation of product, phosphorylated product and Pi in the
DHA cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi
(100 mM), propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1
U/ml) pH 6 in 1 ml.
Observing the figures above it is clear that the AP inhibition problems caused by the
presence of phosphate in the reaction mixture are overcome: the PPi hydrolysis seems to be
complete since the peak area of phosphate reaches 10 after 24 hours (figure 3.9). There is no
presence of phosphorylated product in the reaction mixture after 24 hours indicating that the
reaction is complete (figure 3.8) and the formation of product reaches much higher values than
the ones obtained with the soluble enzyme. The concentration of product at the end of the
reaction is approximately 35 mM (figure 3.7) using immobilized AP (35 % conversion, calculated
from the initial concentration of propanal). The obtained results were very similar to the ones
obtained with PhoN-Sf using the same reaction composition meaning that also AP can be used in
the cascade reaction. The good results obtained using the immobilized AP in the DHA cascade
reaction are supported by the complex HPLC chromatograms shown in Appendix B.
After the reaction was completed (no more aldehyde or phosphorylated product on the
reaction mixture) the supernatant was removed from the supports and the beads were washed
with milli-Q water 2-3 times. A new cascade mixture was prepared (1 ml with the same
58
concentrations as previously mentioned) and added to the same immobilized alkaline
phosphatase and PhoN-Sf beads for a 2nd
cycle of the cascade reaction (cycles of 24 hours).
Figure 3.10 shows the product formation after the two cycles for the AP and PhoN-Sf
beads.
Figure 3.10 – Product formation with immobilized AP and PhoN-Sf after the 2 cycles of DHA cascade
reaction. Reaction mixtures contains DHA (500 mM), PPi (100 mM), propanal (100 mM), RAMA (6 U/ml),
immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml.
As shown in figure 3.10 when the enzymes are reutilized for a 2nd
cycle (refresh the
cascade mix but using the same beads) the cascade reaction performed on the AP beads leads
to a large decrease in product formation when compared to the PhoN-Sf beads. This means that
the beads are clearly deactivated already after only one cycle. Hardly any product is formed and
nearly no PPi hydrolysis occurs (See Appendix C).
Glycerol phosphorylation on soluble AP
Concerning the glycerol route for the cascade reaction it‟s important to study and
optimize the phosphorylation of glycerol (as done previously for DHA). The reaction was carried
using 50 mM pyrophosphate as phosphate donor, 100 mM glycerol and 6 U/ml of soluble AP at
pH 8 and 30°C. After incubation of the reaction mixture, the concentration of glycerophosphate
59
formed was measured using a coupled assay in a buffer of 450 mM glycine, containing 274 mM
hydrazine and 2.4 mM EDTA pH 9.5, 20 U/ml G3P-DH and 2.5 mM NAD+.
Figure 3.11 – Phosphorylation of glycerol by soluble AP. Reaction mixture contains glycerol (100 mM), PPi
(50 mM), AP (6 U/ml) in a buffer of glycine (450 mM), hydrazine (274 mM) and EDTA (2.4 mM) pH 9.5, G3P-
DH ( 20 U/ml) and NAD+ (2.5 mM) in 1 ml.
By this procedure it was possible to obtain a time course for the glycerol phosphorylation.
This concentration of glycerophosphate is calculated as described in section 2.1.5 – Glycerol
phosphorylation (equation 4).
The enzyme G3P-DH is specific for the L-glycerophosphate so only this enantiomer is
consumed in the coupled assay. Figure 3.11 shows that the concentration of L-glycerophosphate
obtained is around 1.1 mM and as we presumably obtain a racemic mixture of L and D
enantiomeres we assume that the total concentration of glycerophosphate formed is around 2.2
mM. These results are in accordance with the results obtained using the acid phosphatase,
PhoN-Sf, therefore it can be concluded that alkaline phosphatase has a good phosphorylating
activity towards glycerol.
Glycerol cascade reaction using immobilized AP
In the second cascade reaction, glycerol is phosphorylated using PPi as a phosphate
donor in the presence of AP to form 2 enantiomers L/D glycerophosphate. The L enantiomer is
then oxidized by glycerol-3-phosphate oxidase (GPO) in combination with catalase to form
60
DHAP. DHAP is then coupled to propanal by the aldolase RAMA and the resulting aldol-product
is then dephosphorylated by the alkaline phosphatase already present in the reaction. This
second cascade reaction was performed using only the immobilized enzyme since our results
show that immobilization improves alkaline phosphatase activity towards the cascade reactions.
The cascade mixture consists of 100 mM PPi, 500 mM glycerol, 100 mM propanal, 10
U/ml catalase, 6 U/ml RAMA and 50 U/ml GPO and the reaction was carried at 25°C and pH 6
and 8. For pH 8, 20 U/ml of immobilized AP were used (fresh beads) and for pH 6 we used the
immobilized acid phosphatase PhoN-Sf 1 U/ml to serve as a control.
Figure 3.12, 3.13 and 3.14 – Time course for the formation of product, phosphorylated product and Pi in
the glycerol cascade reaction using immobilized AP and PhoN-Sf. Reaction mixtures contained glycerol (500
mM), PPi (250 mM), propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized
AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml.
From the three figures above (figures 3.12, 3.13, 3.14) it is obvious that the results
obtained for the immobilized AP are very poor in comparison to the results obtained with PhoN-
Sf. The formation of product is extremely slow and leads to only 3 mM for AP whereas for PhoN-
Sf the product formation leads to around 50 mM (figure 3.12). The problem does not seem to be
61
the PPi hydrolysis since the formation of phosphate is increasing in time almost reaching the
desired final peak area of 10. The problem may rely in the fact that this cascade reaction is hard
to optimize since it involves the use of four enzymes and when a parameter concerning one of
the enzymes is varied the whole system is altered and this may influence the performance of all
the other enzymes.
A new cascade reaction was performed using fresh beads for both immobilized
enzymes, AP and PhoN-Sf and increasing the concentration of glycerol to 3 M. Figure 3.15
shows the comparison between the product concentrations for the two cascade reactions using
different concentrations of glycerol, for both immobilized enzymes.
Figure 3.15 – Product formation with immobilized AP and PhoN-Sf using different concentrations of glycerol
on the glycerol cascade reaction. Reaction mixtures contained glycerol (0.5 and 3 M), PPi (250 mM),
propanal (100 mM), GPO (50 U/ml), catalase (10 U/ml), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and
immobilized phoN-Sf (1 U/ml) pH 6 in 1 ml.
According to these results when the concentration of glycerol was increased to 3M
(utilizing fresh beads) the product formation was much more improved for AP: an increase of
almost 10 times against an increase of 2 times for PhoN-Sf, although the product peak areas are
still much lower for AP than PhoN-Sf. With the PhoN-Sf beads using 3 M of glycerol the
conversion, calculated from the initial concentration of propanal, reaches practically 100 %.
These results are in accordance with the Pradines et al47 who used immobilized AP for
the production of glycerophosphate and tried different concentrations of glycerol. They concluded
that the conversions were higher when the glycerol concentration was increased; the best results
were obtained using 80% glycerol (9.8 M) in the reaction mixture.
62
3.2. Recombinant acid phosphatase from Salmonella enterica: V78L, V78Y and V78H
Previous optimizations of the one-pot method for DHA phosphorylation and subsequent
aldol condensation to an aldehyde by the aldolase RAMA have been made by this group, in
which it was concluded that the wild-type bacterial non-specific acid phosphatase from
Salmonella enterica ser. typhimurium (PhoN-Se) had a higher DHA phosphorylating activity than
the wild-type from Shigela flexneri (PhoN-Sf) in the more alkaline region43.
In the same publication it was shown that a directed evolution approach applied to the
wild-type PhoN-Se gene provides a mutant library with enhanced phosphorylating activity towards
DHA. EpPCR mutagenesis was successfully used to create a PhoN-Se mutant library, and DNA
sequencing revealed 1.8 base changes on average per mutant. After several screenings
monitored by continuous DHAP assays, DNA sequencings revealed that the most promising
mutants all contained the mutation at the V78 residue43.
This residue appears to be very important in tuning the phosphorylation and
dephosphorylation reaction, since it has been previously reported an identical mutation in directed
evolution experiments in which these results were confirmed43, 78.
The group in Amsterdam extensively studied the role of V78 and currently the best
mutants obtained from saturation mutagenesis of the V78 residue of PhoN-Se are: V78Q, V78G,
V78H, V78Y and V78L. These mutants have been screened for DHA and glycerol
phosphorylation at different pH values and the results have been very satisfactory as shown in
figure 3.16.
Figure 3.16 – pH profile for the DHA and glycerol phosphorylation using the best mutants from PhoN-Se
and their comparison with WT PhoN-Se and WT PhoN-Sf. Reactions contain DHA (100 mM) or glycerol
(100 mM), PPi (50 mM) and acetate or Tris/acetate buffer (100 mM), together with PhoN (1 M).
63
In order to improve the acid phosphatase/aldolase cascade reaction for the production of
non natural hydrocarbons, expression, purification and isolation of three of these mutants was
carried out. The mutants were V78 L, V78 Y and V78 H, which introduce respectively a Leucine
(Leu, L), a Tyrosine (Tyr, Y) and a Histidine (His, H) amino acid at the V78 position (figure 3.17).
Figure 3.17 – Amino acid introduced in the V78 position of the enzyme PhoN-Se.
3.2.1. Expression and purification of V78L, V78Y and V78H
The expression of the three mutants was not completely successful. The V78 H mutant
didn‟t show any active fractions after FPLC purification, nor did the microtiter activity assay with
pNPP show any activity for this mutant at this stage. Therefore this mutant wasn‟t purified: as for
the other two mutants the results of expression and purification can be seen in table 3.3. It‟s
clear that the yield of the enzyme mutants in both cases is very low.
Table 3.3 - Results of expression and purification of the V78 L and V78 Y mutants.
Volume culture (L) Final volume (ml) mg/ml mg U/ml U/mg Total U
V78 L 2 4 0.3 1.2 5.2 17.3 20.8
V78 Y 2 6 0.08 0.5 2.8 34.6 17.0
64
3.2.2. Cascade reaction
The two mutants grown and purified, V78 L and V78 Y were then tested for their use in
the DHA cascade reaction.
The cascade mixture consisted of 500 mM DHA, 250 mM PPi, 100 mM propanal, 20 mM
Na-acetate pH 6, 6 U/ml RAMA and different concentrations of mutants were used: 0.5, 1 and 2
µM. The reaction was carried in the thermomixer at pH 6 and 32°C.
The effect of the mutations on the levels of conversion in the one-pot cascade reaction
were evaluated and compared to the results obtained using WT PhoN-Se 1 and 2 µM (figure 3.18
and 3.19).
Figure 3.18 – Time course of the DHA one-pot cascade reaction using WT PhoN-Se and the mutants V78 L
and V78 Y. Reaction mixtures contained DHA (500 mM), PPi (250 mM), propanal (100 mM), Na-acetate pH
6 (20 mM), RAMA (6 U/ml), PhoN (0.5 and 1 µM) in 0.5 ml at pH 6.
Figure 3.18 shows that the WT PhoN-Se is less active leading to lower product
concentrations than any of the other mutants. The best results were obtained with the mutant V78
Y: using 1 µM of this mutant the maximum amount of product (43 mM) was obtained after only 3
hours and using 0.5 µM the maximum amount of product was higher (48 mM) but was obtained
only after 6 hours. This variant is 5 times more active than the WT PhoN-Se in the acid
phosphatase/aldolase cascade reaction at pH 6 as measured after 2 h. The variant V78 L is not
as active being just 2 times more active than the WT PhoN-Se. These results are not in
agreement with previous work done on this subject43 where the mutant V78 L was much more
active in the DHA cascade reaction showing an activity 17 times higher than the WT PhoN-Se for
65
the same concentrations of enzyme used 1 µM. This can be explained by the fact that the mutant
V78 L used in the previous work43 had a specific activity around 10 times higher than the specific
activity of the mutant V78 L expressed and purified here, 17 U/mg. The fact that the V78 Y mutant
presents a higher activity than the V78 L is in accordance with the recent work performed by this
group towards DHA phosphorylation which is illustrated in figure 3.16.
In any case, as can be seen in figure 3.18, the use of even 0.5 µM of the V78 Y and V78
L mutants result in higher conversions than are achieved with 1 µM of the WT enzyme.
The reaction was repeated but using 2 µM of WT PhoN-Se and of each mutant V78 Y
and V78 L. Figure 3.19 shows the amount of product formed using 1 µM and 2 µM of these
variants.
Figure 3.19 – Time course of the DHA one-pot cascade reaction using WT PhoN-Se and the mutants V78L
and V78Y. Reaction mixtures contained DHA (500 mM), PPi (250 mM), propanal (100 mM), Na-acetate pH
6 (20 mM), RAMA (6 U/ml), PhoN (1 and 2 µM) in 0.5 ml at pH 6.
As expected the increase in enzyme concentration leads to higher amounts of product for
every variant V78 L, V78 Y and the WT enzyme. Again the best results were obtained for the
mutant V78 Y where 55 mM of product was obtained in just 2 hours against 50 mM of product for
the V78 L after 4 hours and 40 mM for the WT enzyme after 24 hours.
Also figure 3.19 shows that the use of 1 µM of the V78 Y mutant results in higher
conversions than achieved with 2 µM of the WT enzyme.
66
67
4. Conclusion
4.1. Alkaline Phosphatase
The phosphorylation reactions of both DHA and glycerol by the soluble alkaline
phosphatase appeared to be reasonably effective leading to concentrations of product similar to
those obtained using acid phosphatases (PhoN-Se/Sf) 5, 6.
In the DHA cascade reaction performed using soluble AP the maximum conversion to the
aldol product, 5,6-dideoxy-D-threo-hex-2-ulose, after 24 hours was only 7% at pH 8 and 30°C.
These results were very disappointing when compared to the values obtained with the acid
phosphatases, 60 % conversion to dephosphorylated aldol product43. Two hypotheses were
considered to explain these poor results: AP was suffering from inhibition by the substrate PPi or
by inorganic phosphate, Pi. The second hypothesis was proved to be right since the activity of the
enzyme is extremely affected by addition of phosphate. When a pNPP activity assay is
performed, as described in section 2.1.5 Analytical Techniques - Enzymatic activity assay, the
addition of only 10 mM of Pi decreases the enzyme activity almost 15% and the enzyme
becomes completely inhibited by addition of 100 mM Pi.
The immobilization of the enzyme in three different supports was attempted: epoxy-
functionalized supports Immobeads-150 and Sepabeads EC-EP and amino-functionalized
Sepabeads EC-HA support. The only successful immobilization occurred when using the support
Sepabeads EC-HA aldehyde activated in which after 4 hours the enzyme was completely bound
to the support. This may be explained by the fact that Sepabeads EC-HA is the only carrier with a
relatively long and branched spacer. The length and other properties of spacers such as
hydrophobicity/hidrophilicity and charged/neutral character might influence the binding capability
due to the higher molecular mobility in the presence of spacers between the enzyme and the
carrier.
Performing the DHA cascade reaction with the immobilized AP improves the enzyme
activity in the reaction, leading to a conversion of 35% at 25°C pH 8 in a batch system. This result
was promising and even better than the one obtained using immobilized PhoN-Sf at 25°C pH 6,
30% conversion. However when a 2nd
cycle of the reaction was performed (maintaining the beads
and refreshing the cascade mix) a large decrease in product formation is observed using the AP
beads when compared to the PhoN-Sf beads. This demonstrates that the beads are deactivated
after one cycle.
Regarding the glycerol cascade reaction using the immobilized alkaline phosphatase the
results were not at all promising; 3% conversion using 0.5 M of glycerol and 30% conversion
68
using 3 M of glycerol, at 25°C pH 8. Using the immobilized PhoN-Sf the results were much better;
50 mM for 0.5 M of glycerol and 100 mM for 3 M of glycerol (100% conversion) at 25°C pH 6.
Despite the fact that the use of alkaline phosphatase in this cascade led to very poor product
formation (particularly when using 0.5 M glycerol) one must regard the great improvement on
product formation with the increase in glycerol concentration (10 times increase). This implies that
the enzyme activity is favored by high concentrations of glycerol.
Further research should focus on identifying the operational conditions that maximize the
production of dephosphorylated aldol product and the stability of the immobilized alkaline
phosphatase. Naturally, the next step should be the lab-scale simulation of a continuous process
to perform the cascade reactions, as a flow system with recovery and re-utilization of the
immobilized AP, avoiding the early deactivation of the beads.
4.2. Recombinant acid phosphatase from Salmonella enterica: V78L, V78Y and V78H
In order to improve the acid phosphatase/aldolase cascade reaction for the production of
carbohydrates, expression, purification and isolation of three mutants from PhoN-Se mutagenesis
on V78 residue was carried out. The mutants were V78 L, V78 Y and V78 H.
The expression of the three mutants was not completely successful. The V78 H mutant
didn‟t show any active fractions after FPLC purification, nor did the microtiter activity assay with
pNPP show any activity for this mutant at this stage. Therefore this mutant wasn‟t purified; as for
the other two mutants the expression resulted on a final volume of 4 ml of the V78 L mutant and 6
ml of the V78 Y mutant. After purification the V78 L mutant displayed an activity of 5.2 U/ml and
the V78 Y mutant 2.8 U/ml and a specific activity of 17.3 U/mg and 34.6 U/mg, respectively.
These results were not very satisfactory since the amounts of mutants expressed were very low,
resulting on low specific activities. This may have occurred due to the newly utilized pET 26b
plasmid used for expression, which still has to be optimized.
The two mutants grown and purified, V78 L and V78 Y were tested for their use in the
DHA cascade reaction. The best results were obtained using the V78 Y mutant where 55 mM of
aldol product, 5,6-dideoxy-D-threo-hex-2-ulose, (55% conversion) was obtained in just 2 hours
(reaction led to completion) against 50 mM of product for the V78 L after 4 hours and 40 mM for
the WT PhoN-Se enzyme after 24 hours. These results are not in agreement with previous work
done on this subject43 were the mutant V78 L was much more active showing an activity 17 times
higher than the WT PhoN-Se for the same concentrations of enzyme used, 1 µM. This can be
explained by the fact that the mutant V78 L used in the previous work43 had a specific activity
around 10 times higher than the specific activity of the mutant V78 L expressed and purified here.
69
4.3. Future prospects
The directed evolution approach seems to be the most promising, since the continuing
advances on recombinant DNA technology has made it feasible to express almost every enzyme,
making it possible to modify their selectivity. As it has been done for the acid phosphatases43, 78, it
seems to be of great interest to generate a mutant library for the alkaline phosphatase in order to
enhance its phosphorylating activity towards DHA and glycerol, optimizing its use in the cascade
reactions.
70
71
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78
Appendix A – PPi inhibition
To determine if AP was suffering from PPi inhibition the phosphorylation of DHA and the
PPi hydrolysis reactions were performed with different concentrations of MgCl2 since it has been
reported that the addition of this metal ion improved the pyrophosphatase activity.
Figure A1 – DHA Phosphorylation adding different concentrations of MgCl2. Reaction mixtures contain DHA
(100 mM), PPi (50 mM), AP (6 U/ml) and MgCl2 (1 and 25 mM) at pH 8 and 32°C.
The results are clear; the presence of a small concentration of Mg2+ (Mg2+/PPi = 1:50)
does not lead to great improvements. When the concentration ratio used was Mg2+
/PPi = 1:2 the
reaction nearly does not occur which implies that the enzyme is severely inhibited by excess of
Mg2+
, which is concordant with the mentioned publications74, 75
.
79
Figure A2 – PPi hydrolysis adding different concentrations of MgCl2. Reaction mixtures contain PPi (100
mM), AP (6 U/ml) and MgCl2 [100 (1:1), 66 (2:3) and 50 mM (1:2)] at pH 8 and 32°C.
Figure A2 shows the formation of Pi in 24h in the presence of different concentrations of
Mg2+ as well as for the reaction without addiction of this ion. As concluded for the DHA
phosphorylation, the addition of Mg2+
didn‟t improve the reaction in any of the concentration ratios
employed, the reaction without magnesium leads to higher peak areas of the product and in a
faster rate. In the presence of magnesium the best concentration ratio was Mg2+
/PPi = 1:2, as
advised in literature74, 75. When the concentration ratio used was Mg2+
/PPi = 1:1 the enzyme is
clearly affected by the excess of magnesium, the hydrolysis is really slow and stops at 40%
conversion.
80
Appendix B – HPLC chromatogram of DHA cascade using immobilized AP (1
st cycle)
Figure B1 – HPLC chromatogram obtained for the DHA cascade reaction using immobilized AP
after 0 and 24 hours (black and blue line respectively). The identified peaks are in the UV spectra and
correspond to: PPi – pyrophosphate, Phosphorylated product, DHA and Product. All unidentified peaks
correspond to impurities originated either from the solutions utilized or the HPLC system.
As depicted in figure B1 the pyrophosphate is almost entirely consumed after 24 hours of
reaction, since its peak decreases largely from time zero. In addition, the phosphorylated product
after 24 hours is also practically inexistent, being its peak much higher in the middle of the
reaction (not shown). These facts show that the reaction is led to completion after 24 hours, with
a relatively good amount of product formed.
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Figure B2 – HPLC chromatogram obtained for the DHA cascade reaction using immobilized AP
after 0 and 24 hours (black and blue line respectively). The identified peaks are in the IR spectra and
correspond to: PPi – pyrophosphate, Pi – inorganic phosphate, DHA and Propanal. All unidentified peaks
correspond to impurities originated either from the solutions utilized or the HPLC system.
Figure B2 indicates that the pyrophosphate is almost entirely hydrolysed to Pi after 24
hours of reaction. In addition, the propanal after 24 hours is practically inexistent. These facts
also show that the reaction is led to completion after 24 hours.
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Appendix C – DHA cascade using immobilized AP (2nd
cycle)
Figure C1 – Time course for the formation of phosphorylated product in the 2nd
cycle of the DHA cascade
reaction using immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM),
propanal (100 mM), RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6
in 1 ml.
Figure C2 – Time course for the formation of Pi in the 2nd
cycle of the DHA cascade reaction using
immobilized AP and PhoN-Sf. Reaction mixtures contain DHA (500 mM), PPi (100 mM), propanal (100 mM),
RAMA (6 U/ml), immobilized AP (20 U/ml) pH 8 and immobilized PhoN-Sf (1 U/ml) pH 6 in 1 ml.
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As depicted in the figures above when a 2nd cycle of the DHA cascade reaction is
performed using the same AP beads as for the 1st cycle, these become inactive towards the
cascade reaction. That is explained due to the fact that the phosphate is being produced very
slowly and the phosphorylated product is still present after 24 hours.
