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University of Groningen
Hydrolase-catalyzed synthesis of polyamides and polyester amidesStavila, Erythrina
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Hydrolase-catalyzed synthesis of
polyamides and polyester amides
Erythrina Stavila
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Hydrolase-catalyzed synthesis of polyamides and polyester amides
Erythrina Stavila
PhD thesis
University of Groningen
The Netherlands
Zernike Institute PhD thesis series 2014-07
ISSN: 1570-1530
ISBN: 978-90-367-6923-5 (printed)
978-90-367-6924-2 (electronic)
The research presented in this thesis was performed in the research group of
Macromolecular Chemistry and New Polymeric Materials of the Zernike
Institute for Advanced Materials at University of Groningen, The Netherlands.
This research was financially supported by a Ubbo Emmius PhD Scholarship of
University of Groningen.
Cover design by Erythrina Stavila and Muhammad Iqbal
Printed by Offpage, Amsterdam. www.offpage.nl
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Hydrolase-catalyzed synthesis of
polyamides and polyester amides
Proefschrift
ter verkrijging van de graad van doctor aan de
Rijksuniversiteit Groningen
op gezag van de
rector magnificus prof. dr. E. Sterken
en volgens besluit van het College voor Promoties.
De openbare verdediging zal plaatsvinden op
vrijdag 9 mei 2014 om 16.15 uur
door
Erythrina Stavila
geboren op 16 juni 1987
te Bandung, Indonesië
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Promotor
Prof. dr. K. Loos
Beoordelingscommissie
Prof. dr. N. Bruns
Prof. dr. G. Fels
Prof. dr. F. Picchioni
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Untuk Keluargaku Tersayang
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Contents
Chapter 1 General introduction
1
1.1. Enzymes in Organic Synthesis 2
1.2. Hydrolases 4
1.3. Enzyme Immobilization 8
1.4. Polyamide 10
1.5. Enzymatic Synthesis of Polyamide and its Copolymers 12
1.6. Aim and Outline of the Thesis 20
1.7. References 21
Chapter 2 Immobilization of Fusarium solani pisi cutinase
27
2.1. Introduction 28
2.2. Experimental Methods 30
2.3. Results and discussion 33
2.4. Conclusion 39
2.5. References 40
Chapter 3 Fusarium solani pisi cutinase-catalyzed synthesis of polyamides
43
3.1. Introduction 44
3.2. Experimental Methods 45
3.3. Results and discussion 47
3.4. Conclusion 56
3.5. References 57
Chapter 4 Synthesis of lactams using enzyme-catalyzed aminolysis
59
4.1. Introduction 60
4.2. Experimental Methods 61
4.3. Results and discussion 64
4.4. Conclusion 69
4.5. References 69
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Chapter 5 Enzyme-catalyzed synthesis of aliphatic-aromatic oligoamides
71
5.1. Introduction 72
5.2. Experimental Methods 73
5.3. Results and discussion 76
5.4. Conclusion 89
5.5. References 90
Chapter 6
Lipase-catalyzed ring-opening copolymerization of -caprolactone
and -lactam
93
6.1. Introduction 94
6.2. Experimental Methods 96
6.3. Results and Discussion 99
6.4. Conclusion 115
6.5. References 116
Chapter 7 Summary 119
Samenvatting 123
Acknowledgements 127
List of publications 131
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Chapter 1
General introduction
Abstract
The history of using enzymes in organic synthesis and the industrial
applications of enzymes are presented in this chapter. Two types of enzymes
used in this research are briefly reviewed in terms of their immobilization and
further use in enzymatic reactions; the enzymes are Candida antarctica lipase B
and Fusarium solani pisi cutinase. Furthermore, a literature overview describing
recent developments in the field of the enzymatic synthesis of polyamides and
its copolymers is presented in this chapter.
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Chapter 1
2
1.1. Enzymes in Organic Synthesis
Enzymes, also known as biocatalysts, are proteins with catalytic activity that
control the rates of metabolic reactions in living cells.1 Apart from their natural
function in living cells, since ancient times, enzymes have been used in the
preparation of food products such as cheese, beer, wine, vinegar, sourdough,
and in the manufacture of commodities such as linen, leather, and indigo.2
One unique feature of enzymes is the specificity towards a variety of
substrates. It was suggested by Fischer in 1894 and postulated as ‘the lock and
key’ model that both enzyme and substrates possess specific complementary
shapes so they can fit into another.3 This specificity is of tremendous benefit for
the use of enzymes in organic synthesis, as well as pharmaceutical and industry
applications. The chiral nature of the enzymes contributes to high
enantioselectivity towards substrates and results in the formation of stereo- and
regio-chemically defined reaction products.4 Enzymes reduce the amount of
undesirable byproduct formation leading to more efficient reactions, also
reducing the waste. Therefore, enzymes are considered environmentally friendly
catalysts.1,4
Over 3000 enzymes have been identified and classified into six enzyme
classes (EC) based on the type of reactions they catalyze.5,6
These enzymes are
considered to be highly efficient catalysts for a broad range of organic synthesis
transformations and some of them are also suitable for industrial scale
applications.6
EC.1. Oxidoreductases catalyze oxidation or reduction reactions.
EC.2. Transferases catalyze the transfer of functional groups, such as
aldehyde or ketonic, acyl, glycosyl, alkyl, aryl, nitrogenous, etc.
EC.3. Hydrolases catalyze the hydrolysis of esters, amides, ethers, carbon-
carbon, phosphorus-nitrogen, and so on.
EC.4. Lyases catalyze the addition of chemical groups to double bonds, as
well as the reverse reaction by removing chemical groups without
hydrolysis.
EC.5. Isomerases catalyze the rearrangement of isomers via racemization,
epimerization, and intramolecular reactions.
EC.6. Ligases catalyze the bond formation, but require nucleoside
triphosphates for activation of the enzymes.
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General introduction
3
In nature, enzymatic catalysis typically takes place in aqueous media,
whereas most enzymatic catalysis in organic synthesis takes place in organic
solvents as a reaction medium because many organic compounds are unstable or
insoluble in aqueous environments.7,8
Changing the reaction medium from an
aqueous to an organic solvent may lead to new types of enzymatic reactions.
For example, hydrolases catalyze the hydrolysis of esters to their corresponding
alcohols and acids in aqueous media. This reaction does not take place in
organic solvents due to the lack of water, but the addition of alternative
nucleophiles such as alcohols, amines or thiols leads to transesterification,
aminolysis, or thiotransesterification, respectively.7 Enzymatic reactions in
organic solvents offer advantages including easy recovery of products, the
possibility to use non-polar substrates, avoiding side reactions, increasing the
activity (in most cases for lipases), and shifting the thermodynamic equilibrium
to favor synthesis over hydrolysis that results in discovering new types of
enzymatic reactions.8,9
Due to remarkable progress in enzyme discovery, enzyme engineering and
biotechnology, more enzymes have been used for a plethora of in vitro
applications in organic synthesis, as well as in pharmacy and industry. Some of
the application of enzymes in various industries are summarized in Table 1.1.2
Table 1.1. Applications of enzymes in various industrial segments
Industry Enzyme Application
Detergent Protease, amylase, lipase,
mannanase
Stain removal
Starch Amylase, amyloglucosidase,
pullulanase
Saccharification
Food Protease, lipase, lactase Milk and cheese flavor
Baking Amylase, xylanase, lipase,
phospholipase, lipoxygenase
Dough stability conditioning
Animal feed Transglutaminase, phytase,
-glucanase
Digestibility
Beverage Pectinase, amylase, -
glucanase, laccase
Mashing, juice treatment, flavor
Textile cellulose, amylase, laccase,
peroxidase
Cotton softening bleaching
Pulp and paper Lipase, protease, amylase,
xylanase, cellulose
Contaminant control, biofilm
removal, starch-coating
Fats and oils Lipase and phospholipase Transesterification, de-gumming
Organic synthesis Lipase, acylase, nitrilase Resolution of chiral alcohol and
amides, synthesis enantiopure
Leather Protease, lipase Bating, de-pickling
Personal care Amyloglucosidase, glucose
oxidase, peroxidase
Antimicrobial, bleaching
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Chapter 1
4
1.2. Hydrolases
Enzymes from the hydrolase class (EC.3) are the most commonly used
enzymes in organic synthesis due to their acceptance towards a large variety of
substrates, considerable stability in organic solvents, direct use without addition
of cofactors, and the fact that many of them are commercially available.8
Candida antarctica lipase B (CAL-B) and Fusarium solani pisi cutinase both
members of the hydrolase class are described in-depth in this chapter.
1.2.1. Candida Antarctica Lipase B (CAL-B)
Lipases are enzymes that predominantly catalyze the hydrolysis and
formation of ester bonds in lipids (see Scheme 1.1).10
Lipases have been used in
many industrial applications as shown in Table 1.1. CAL-B is one of its most
famous proponents and has been used in many organic syntheses.
Scheme 1.1. Hydrolysis or synthesis of ester bonds in lipid catalyzed by lipase
CAL-B was firstly produced from the basidomycetous yeast Candida
antarctica which was isolated from sediment at the bottom of the lake Vanda,
Antarctic.11-13
Nowadays, CAL-B is industrially produced by submerged
fermentation of a genetically modified Aspergillus microorganism.14
The crystal structure of CAL-B was elucidated by Uppenberg et al.15
The
results revealed that CAL-B has an α/β-hydrolase-like fold and an active site
catalytic triad (Ser105, His224, and Asp187) that is accessible from the
solvent.15,16
The results also revealed that the polypeptide chain of CAL-B is
built out of 317 amino acids with a molecular weight of 33 kDa and
approximate dimension of 30 x 40 x 50 Å3.15
Moreover, it was also suggested
that a 5-residue long hydrophobic helix (5 helix) in CAL-B act as a lid
because of its observed disorder in some crystal structures, which indicates a
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General introduction
5
region of high mobility.15
The structure of CAL-B is presented in Figure 1.1 and
the catalytic triad is emphasized in purple.
Figure 1.1. Structure of Candida antarctica lipase B; image from the RSCB PDB
(www.pdb.org) of PDB ID 1TCA15
The study of interfacial activation of CAL-B has been performed by
Martinelle et al.17
They revealed that the hydrophobic lid (5 helix) is not
involved in any conformational change in regulating the access to the catalytic
site, which is why CAL-B does not display interfacial activation. Therefore,
CAL-B has an easily accessible catalytic site towards substrates and can display
maximum activity at the water/lipid interface as well as towards monomeric
substrates.17
CAL-B has a broad substrate specificity and exhibits a very high degree of
substrate selectivity (regio- and enantioselectivity).12
CAL-B is a well-known
catalyst in organic synthesis due to its high effectiveness in the resolution of
alcohols and especially amines, which led to the synthesis of an important
variety of optically active hydroxyl and amino compounds.18
CAL-B has high
stability in an immobilized form and can be used at elevated temperatures for
hours without any significant loss in activity.19
Because of these intrinsic
benefits, immobilized CAL-B is used in many enzyme catalyzed syntheses of
polymers.
The most well-known immobilized CAL-B is the commercially available
Novozym® 435 (N435). N435 is a heterogeneous biocatalyst that consists of
physically immobilized CAL-B within macroporous poly(methyl methacrylate)
beads, which are known as Lewatit beads with a particle size of 315–1000 m
and an average pore size of 140–170 Å.14
N435 has been used as catalyst in
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Chapter 1
6
chemical synthesis and production of polyesters,20-25
polycarbonates,20-22
polyamides and its copolymers.20,23,25-27
Some of these enzymatic
polymerizations have been carried out on large scale, for instance Baxenden
Chemical Ltd. has performed a large scale process for the enzymatic synthesis
of poly(hexane-1,6-diyl adipate).28,29
Furthermore, BASF AG has patented
different methods in enzymatic synthesis of polyamides for the production of
aqueous polyamide dispersions.29-31
1.2.2. Fusarium Solani Pisi Cutinase (F. solani pisi)
Cutinase (cutin hydrolase) is a serin esterase that catalyzes the hydrolysis of
cutin.32,33
Cutin is the structural component of plant cuticle and is an insoluble
polymer composed of hydroxy fatty acids (C16 and C18) and contains up to three
hydroxyl groups,33-35
as shown in Figure 1.2.
Figure 1.2. Cutin and some cutin monomers that originate upon hydrolysis (adapted
from Carvalho et al.35
)
Cutinase from pathogenic fungus Fusarium solani pisi is the smallest
lypolytic enzyme composed of 197 amino acids and has a relative molecular
mass of 22–25 kDa.33,35-37
The crystal structure of Fusarium solani pisi was
elucidated by Martinez et al.37
They reported that cutinase is an α/β protein with
the catalytic triad composed of Ser120, Asp175, and His188. Cutinase does not
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General introduction
7
display interfacial activation because the catalytic serin is not buried under the
surface loops (lid), and therefore accessible to solvents. Furthermore, cutinase is
a compact one-domain molecule, with dimensions of approximately 45 x 30 x
30 Å3. In Figure 1.3, the structure of Fusarium solani pisi cutinase is presented
and the active site of the enzyme is highlighted in purple.
Figure 1.3. Structure of Fusarium solani pisi cutinase; image from the RSCB PDB
(www.pdb.org) of PDB ID 1CUS37
Cutinase is able to hydrolyze lipids and may thus be considered as a lipase,
although it does not display interfacial activation like most lipases due to the
absence of a hydrophobic lid in its structure that covers the catalytic site.36,37
Studies on the dynamics of F. solani pisi cutinase via structural comparison
among different crystal forms has been performed by Longhi et al.38
They
revealed that the absence of any significant structural rearrangements upon
binding highlights the important feature of cutinase,32
which is probably shared
only by Candida antarctica lipase B (CAL-B).15,38
Egmond et al. reported
kinetic studies of cutinases and showed they resemble similar kinetic behavior
to lipases and allow cutinase to be used as a model in studying more complex
lipases.39
Therefore, cutinase establishes a bridge between lipases and esterases
because it has the capabilities of both families.32,37,39
Studies on the applications of cutinases have been performed extensively
and were reported in several review articles.35,40,41
Cutinases have been used in
various chemical reactions (e.g., hydrolysis,42-45
esterification,46-48
and
transesterification49
) and as a lipolytic enzyme in industry or dishwashing
detergent composition to remove fats.35,40
Furthermore, several patents have
been registered concerning the application of cutinase in industry, such as in
industrial cleaning processes, degradation of biodegradable polymers, surface
modification of polyacrylonitrile and polyamide fibers, production of esters in
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Chapter 1
8
the absence of organic solvents, enhancing the effect of agricultural pesticides,
increasing the surface permeability of fruits and vegetables, and removing the
excess dye in industrial textile dyeing processes.41
Cutinases are not only successfully used in polymer degradation
(hydrolysis)42-45
but have also been applied as catalysts in the synthesis of
polymers, for example in the syntheses of polyesters.50-52
Gross et al. 50-52
have
used cutinase from Humicola insolens (HIC) in its immobilized form in the
syntheses of polyesters. They discovered the similar kinetic and mechanistic
characteristic between CAL-B and HIC in the polymerization of -
caprolactone.50
In our studies, syntheses of polyamides were performed using
cutinase from Fusarium solani pisi cutinase and will be discussed in detail in
chapter 2, 3, and 5.
Fusarium solani pisi cutinase is the most studied and well characterized
cutinase.35,37,40,53
It has been used in different forms in various reaction media,
such as dissolved in aqueous solution, suspended as powder, microcapsulated in
reversed micelles, and as immobilized enzyme.35
Cutinase has been used in
immobilized form in polymer synthesis, and because of this no denaturation
occurred when used in organic solvents and at elevated temperature. Unlike
CAL-B, immobilized Fusarium solani pisi cutinase is not yet commercially
available, although several patents have been registered for application of this
enzyme.41,54
Therefore, immobilization of the enzyme will have to be performed
before using this enzyme in enzymatic polymerization.
1.3. Enzyme Immobilization
The limiting factor in the use of enzymes in enzymatic polymerizations is
generally their low stability towards both organic solvents and the high
temperatures required to perform the polymerization reactions. Because many
chemical compounds can only be dissolved in organic solvents,8 one way to
improve the stability of the enzyme in organic solvents is immobilization.
Enzyme immobilization is the localization or physical confinement of an
enzyme in a certain region of space with retention of its catalytic activities and
which can be used repeatedly and continuously.55
Since the beginning of the
19th century, long before the enzyme immobilization process was well
established, immobilized microorganisms were already employed industrially in
vinegar production and waste-water treatment.56
For instance in the production
of vinegar, an alcohol-containing solution was passed over wood chips adsorbed
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General introduction
9
with ‘acetic acid bacteria’. In a water purification process, a trickling filter
(mixed with microorganism) was used in the breakdown of waste substances.
Nowadays, immobilized enzyme has been applied in various organic
syntheses,8,18
polymerizations,20-23,57-59
and industrial applications.2,29
There are several advantages for using enzyme in immobilized form, which
are: (a) immobilization facilitates separation from the product (e.g., by filtration
or centrifugation), and thus minimize or eliminate protein contamination of the
product; (b) increased enzyme activity (up to a factor of 100) in organic
solvents; (c) increased temperature stability; (d) increased enantioselectivity; (e)
improved long term stability; (f) improved reusability of the enzymes even for
other types of reactions.60-63
1.3.1. Enzymes Immobilization Methods
Enzyme immobilization methods can be divided into three types: support
binding, entrapment (encapsulation), and cross-linking, 60-63
as shown in Figure
1.4.
Figure 1.4. Different types of enzyme immobilization methods via (a) support binding,
(b) entrapment (encapsulation), (c) cross-linking
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Chapter 1
10
The support binding method can be performed via adsorption (physical
interaction between enzyme and support such as by hydrophobic and Van der
Waals interactions), ionic, or covalent binding. The binding strength decreases
via the following order: covalent > ionic > adsorption. Entrapment
(encapsulation) is carried out through inclusion of the enzyme in a polymer
network or gel lattice (such as organic polymer or silica sol-gel) or in a
membrane-device (hollow fiber or microcapsule). For the cross-linking method,
individual enzymes are linked together using a bifunctional reagent, such as
glutaraldehyde, to prepare carrierless macroparticles.61-63
Our studies focus on the immobilization of cutinase. There are many
examples of immobilized cutinase in literature and they have been used in
hydrolysis, esterification, and transesterification reactions.35
Immobilized
cutinase has been prepared via adsorption (onto zeolite, Teflon, silica, PA6,
polystyrene EP 100, or chromosorb P), covalent binding (on porous silica or
derivatized silica supports), entrapment (in calcium alginate), and
microencapsulation (using anionic, cationic, or nonionic surfactants).35
Specifically for the use as catalysts in the synthesis of polyesters, the
immobilizations have been carried out via physical adsorption onto Lewatit
beads50,52
and covalent attachment on Amberzyme oxirane beads.51
. The results
demonstrated that immobilized cutinase (from Humicola insolens) catalyzed the
synthesis of polyesters with high monomer conversion and molecular weight,
which was similar to polyesters synthesized by CAL-B (N435). In respect to
recent developments, same approach was chosen in our studies for the
preparation of immobilized F. solani pisi cutinase for the enzymatic synthesis
of polyamides.
1.4. Polyamide
Polyamide is a polymer that contains a repeating unit linked together with an
amide bond (–CONH–). Polyamides can be found as a natural (protein) or
synthetic polymer; however, in this chapter, the discussion will be devoted to
synthetic polyamides. According to the composition of their repeating units,
polyamides are classified into three types, which are (a) aliphatic, (b) aromatic,
and (c) aliphatic-aromatic polyamides, as presented in Figure 1.5.
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General introduction
11
Figure 1.5. Classification of polyamides based on the composition of repeating units (a)
aliphatic, (b) aromatic, (c) combination of aliphatic and aromatic
Aliphatic polyamides, which are commercially known as nylon, are versatile
synthetic fibers and engineering plastic materials due to their high mechanical
and heat resistance properties.64
The first nylon was poly(hexamethylene
adipamide) and it was commercialized as nylon-6,6 and used for toothbrush
filament by DuPont in 1938.65
Another nylon designated nylon-6 was
synthesized from -caprolactam and first described in 1938.66
Even today,
nylon-6,6 and nylon-6 are the most used polyamides. In addition, about two-
thirds of the production of nylon-6,6 and nylon-6 is converted to fibers, with the
remainder used as engineering plastic materials.64
Aromatic polyamide or aramid is known as a high performance material. It
has high chemical resistance, superior mechanical and thermal resistance
properties compared to aliphatic polyamide.67-71
Aramid is different from nylon
because over 85% of the amide bonds are bound to two aromatic rings.68,72
The
fully aromatic structure and amide linkages in aramid contribute to the stiff rod-
like macromolecular chains that interact with each other via highly directional
hydrogen bonding, which results in a highly compact intermolecular
packing.67,68
Aramid has been used for several applications, such as high
strength and modulus fibers, high temperature resistant coatings, and highly
efficient semipermeable membranes.69
Moreover, commercial aramids, such as
poly(p-phenylene terephthalamide) (PPPT) under trade name ‘Kevlar®’ and
poly(m-phenylene isophthalamide) (PMPI) are used as electrical insulations,
bullet-proof body armor, industrial fillers, etc.67,68,70
Certain polyamides are synthesized from a combination of aliphatic and
aromatic monomers. They are known as aliphatic-aromatic polyamides and
mostly have better solubility than aramid and better mechanical and thermal
properties than nylon. The commercially available aliphatic-aromatic polyamide
polyphthalamides (PPAs) is produced by DuPont. The aromatic structure in
PPA give advantages like higher glass transition temperature (Tg), higher
melting temperature, and lower adsorption of moisture and solvent compared to
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Chapter 1
12
nylon.73
PPAs have been used mainly as engineering thermoplastics in
automotive components.73
Another example of application in the intumescent
flame retardant (IFR) of polypropylene (PP) is using poly(hexamethylene
terephthalamide) (PA6,T) as carbonization agent. Studies showed that PA6,T
has promising flame retardancy properties74
compared to using nylon-6 as
carbonization agent.75
The synthesis of polyamide can be carried out via three different methods (a)
polycondensation of -aminocarboxylic acid, (b) polycondensation of
diester/diacid and diamine, and (c) ring-opening polymerization of lactam. In
this chapter, the discussion is focused on enzymatic synthesis of polyamide and
polyamide copolymers.
1.5. Enzymatic Synthesis of Polyamide and its Copolymers
The global sales of polyamides (nylon-6 and nylon-6,6) was up to 2.6
million ton in 2006 (around 30% in total market of engineering plastics) and
they are therefore considered to be one of the largest engineering polymer
families.76
The industrial synthesis of most polyamides is carried out by a
melting process. The use of elevated temperature reactions leads to thermal
degradation and undesired polymer products.70,77
Polymerization under the
polymer’s melting temperature (~ 215 °C) can be performed by using anionic
polymerization methods in the synthesis of nylon-6. However, the
polymerization involves the use of alkali metal catalyst such as Na, NaH,
C2H5MgBr, LiAlH4, etc.64
Because of increasing environmental concerns, many
efforts in the development of green chemistry have taken place in recent
decades and one of them is the use of enzymes as green catalysts. The
enzymatic synthesis of polymers not only allows the application of milder
reaction conditions, but also the use of enzymes as non-toxic catalyst which are
derived from renewable resources.20
Over the past decades, many enzymatic polymerizations have been
developed. The first attempt in enzymatic synthesis of an oligoester was
performed in 1984 by Okumara et al.78
More studies of enzymatic syntheses of
polymer have been carried out, such as synthesis of polyesters, polysaccharides,
polycarbonates, vinyl polymers, polyamides, and polyaromatics.20-24,26-29,57-59,62,79
Enzymatic polymerization is defined as an in vitro synthesis of polymer
catalyzed by enzyme and does not follow biosynthetic pathways.57,58
Therefore,
polymers synthesized via bacterial fermentation methods such as poly(3-
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General introduction
13
hydroxyalkanoates) and poly(γ-glutamic acid)20
are not classified as enzymatic
polymerizations.
In the following section of this chapter, the discussion focuses on the recent
developments in the enzymatic synthesis of polyamide and its copolymers using
lipase or cutinase as catalyst.
1.5.1. Polycondensation
Synthesis of polyamide via polycondensation is divided into two: A-B type
and AA-BB type polycondensations.
Polycondensation of A-B Monomer
A-B type monomer denotes a monomer with two different reactive end
groups. Polycondensation of A-B type monomer is also known as self
polycondensation where groups A and B react with each other,23
as presented in
Scheme 1.2.
Scheme 1.2. Enzymatic polymerization of -aminocarboxylic acid
Kong et al.30
have filed a patent application on the method of preparing
aqueous polyamide dispersions by lipase-catalyzed condensation of -
aminocarboxylic acids (C2–C30). The conventional processes for preparing
aqueous polyamide dispersions are generally multistage, technically very
complex, and energetically demanding. However, they succeeded in preparing
aqueous polyamide dispersion via enzymatic polymerization. They performed
the reaction in miniemulsions using dispersants (non-ionic or anionic
emulsifiers) and addition of water-immiscible solvent up to 60% (w/w) (such as
toluene). The dispersed phase has an average diameter of 1 m in the aqueous
medium. Miniemulsions containing lipase (0.5–8% (w/w)), dispersants, and
water were also prepared and further mixed with the miniemulsions containing
aminocarboxylic acids. The reactions were carried out at 60 °C and stirred for
20 hours under a N2 atmosphere.
Poulhès et al have performed the synthesis of chiral polyamide using amino
ester as a monomer.80
The chiral monomer of (R)-amino ester was synthesized
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Chapter 1
14
prior to use in the polymerization. The polymerization was carried out in
diphenyl ether at 80 °C using N435 as catalyst for 240 h under reduced pressure
(3 mbar), as shown in Scheme 1.3. They reported that the chiral polyamide had
a molecular weight of 1316 Dalton and PDI 1.9. No polyamide was formed in
the control reaction (without the addition of N435).
Scheme 1.3. Enzymatic synthesis of chiral polyamide using CAL-B as catalyst,
reproduced from Poulhès et al.80
Copyright (2012) with permission from Elsevier
Poulhès et al.81,82
have also reported another enzymatic synthesis of
polyamide using an amino-ester containing an ethylene glycol moiety as
monomer (Figure 1.6). All the polymerizations were performed using
immobilized CAL-B (N435) as catalyst. The presence of the ethylene glycol
moiety increases the solubility of the resulting polymer in organic solvent.
Moreover, Poulhès et al.82
also observed that by using -amino-α-alkoxy-
acetate as monomer, 93% conversion can be reached within 30 minutes.
Figure 1.6. Amino-esters contain ethylene glycol moiety. Adapted from Poulhès et
al.81,82
Polycondensation of AA and BB Monomers
The AA and BB type monomers are difunctional monomers with the same
end group, such as diesters, diacids, diamines, etc. An example of the
polycondensation of an AA and BB monomer is presented in Scheme 1.4.
Scheme 1.4. Enzymatic polycondensation of diacid/diester and diamine
-
General introduction
15
The preparation of high molecular weight polyamides via enzymatic
polymerization was first reported by Cheng et al.83
The polyamides had
molecular weights of 3000–10000 Daltons. The reactions were carried out using
different dialkyl esters (adipate, malonate, phenylmalonate, or fumarate) and
amines (NH2-terminated triethylene glycol, triethylene tetraamine, or diethylene
triamine) using commercial lipases.
The synthesis of water soluble poly(aminoamide) has been reported by Gu et
al.84
The poly(aminoamide)s were synthesized using lipase as catalyst at 70–90
°C. They found that CAL-B (N435) and Mucor miehei (e.g., Amano lipase M)
showed the highest activities. Various types of poly(aminoamide)s with
different molecular weights were produced from these reactions, DETA
(diethylene triamine)–adipate polyamide (Mw 8400 and PDI 2.73), TETA
(triethylene tetraamine)–adipate polyamide (Mw 8000 and PDI 2.10), TEGDA
(triethylene glycol diamine) polyamide (Mw 4540 and PDI 2.71).
BASF AG has patented a method for the preparation of aqueous polyamide
dispersions via lipase-catalyzed polycondensation.31
The AA-BB type
polycondensation is presented in Scheme 1.5. The reactions were carried out at
60 °C at a pH between 3 and 9. The resulting polyamide had a molecular weight
(Mw) of 5200 g/mol.
Scheme 1.5. Enzymatic synthesis of polyamide in aqueous dispersion
Azim et al.85
have reported the synthesis of oligoamides catalyzed by N435.
The reaction was carried out between dialkyl ester (e.g., diethyl allylmalonate)
and diamine (e.g., 1,12-dodecanediamine) and the resulting oligoamides had a
degree of polymerization (DP) up to 9. Another patent from Panova et al.86
described that the polycondensation of diester and diamine not only resulted in
linear polyamides or oligoamides, but also resulted in the formation of
macrocyclic amide oligomers that could be useful for the subsequent production
of higher molecular weight polyamides.86
-
Chapter 1
16
The synthesis of silicone aromatic polyamides (SAPAs) was reported by
Poojari et al.87
, as shown in Scheme 1.6 where diaminopropyl-terminate
poly(dimethylsiloxane) was reacted with dimethyl terephthalate. Two different
molecular-masses of APT-PDMS (Mn 1000 and 4700 g/mol) were used and 96
hours reaction time resulted in SAPAs of Mn 5700 and 40000 g/mol,
respectively.
Scheme 1.6. Lipase-catalyzed polycondensation of α,-(diaminopropyl)-terminated
poly(dimethylsiloxane) (APT-PDMS) with dimethyl terephthalate (DMT) carried out in
toluene at 80 °C for 48–96 h under reduced pressure (copyright from Poojari et al.87
copyright (2010) American Chemical Society)
The synthesis of chiral polyamides and polyamides containing ethylene
glycol moieties were also reported by Poulhès et al.80-82
By using AA-BB
polycondensation, chiral polyamide was synthesized using immobilized CAL-B
as catalyst, as shown in Scheme 1.7. The reaction was performed in diphenyl
ether at 80 °C for 240 h under reduced pressure (3 mbar) and resulted in the
formation of a chiral polyamide with a yield up to 87% and Mw 19280 D. The
crystallinity and thermal properties of the resulted polyamides were determined
as well.
Scheme 1.7. Enzymatic synthesis of chiral polyamide from AA-BB monomer using
CAL-B as catalyst, reproduced from Poulhès et al.80
-
General introduction
17
Ragupathy et al.88
have reported syntheses of nylon-8,10, nylon-6,13, nylon-
8,13, and nylon-12,13 using immobilized CAL-B as catalyst. The reactions
were carried out in one- (in toluene at normal pressure), two- (in dried diphenyl
ether at reduced pressure), or three-steps (in diphenyl ether at two-step reduced
pressure) enzymatic reactions. By performing three-step enzymatic reactions,
yields up to 97% can be reached and a molecular-mass Mn (determined from 1H
NMR) up to 5380 g/mol. Detailed studies found that nylon-8,10 had 4 identified
microstructures, amine-ester, amine-amine, amine-amine, and ester-ester.
1.5.2. Ring-Opening Polymerization
Kong et al.30
have reported the preparation of aqueous polyamide
dispersions via lipase-catalyzed ring-opening polymerization of lactam, as
shown in Scheme 1.8. They used various sizes of lactams, among them -
caprolactam. They successfully produced nylon-6 (poly(-caprolactam)) with
Mw of 212000 g/mol and Mn of 47000 g/mol.
Scheme 1.8. Enzymatic ring-opening polymerization of lactams
Mechanistic study on enzymatic ring-opening polymerization of β-
propiolactam has been established by Schwab et al.89
and collaborators.90,91
Its
proposed mechanism scheme is summarized in Scheme 1.9. Schwab et al.89
also
reported the synthesis of linear nylon-3 (poly(β-alanine) by lipase-catalyzed
ring-opening polymerization β-lactam. The resulting nylon-3 had a low average
DP of 8, which is likely caused by low solubility of nylon-3 in the reaction
medium. They also performed control experiments with β-alanine as a substrate
and confirmed that the ring structure of β-lactam was necessary to obtain the
polymer.
-
Chapter 1
18
Scheme 1.9. Mechanism of enzyme-catalyzed ring-opening polymerization of β-lactam
developed by Baum et al.90,91
(reproduced from Schwab et al.92
)
1.5.3. Enzymatic Copolymerization
The enzymatic synthesis of copolymers containing amide bonds or
polyamide blocks can be carried out via ring-opening copolymerization,
polycondensation, or via a combination of ring-opening and polycondensation
in a one-pot reaction. Several selected reports are briefly discussed in this
section.
The first report on the block copolymer synthesis using enzymatic
polycondensation was by Gross and Scandola et al.93
They described the
synthesis and solid state properties of polyesteramides with a
poly(dimethylsiloxane) (PDMS) block. The polycondensation was carried out
with various ratios of dimethyl adipate, octanediol, and diamine-functionalized
PDMS (Mw = 875). The physical properties of the resulted copolymer varied
from hard solids to sticky materials.
In the paper discussing the preparation of aqueous polyamide dispersions,
Kong et al.30
also produced copolyesteramides. The copolymer was prepared
via enzymatic ring-opening copolymerization of pentadecanolide and -
caprolactam with ratio of 1:2.3. The produced copolymer had amide and ester
bonds in its backbone chain, as shown in Scheme 1.10. The copolymer had a
-
General introduction
19
weight-average molecular weight (Mw) of 16600 g/mol and two melting points
at 97 °C and ~ 210 °C.
Scheme 1.10. Enzymatic synthesis of aqueous polyamide dispersion by Kong et al.
30
Palsule et al.94
have reported the synthesis of silicone fluorinated aliphatic
polyesteramides (SAFPEAs) using immobilized CAL-B as catalyst, as shown in
Scheme 1.11. They proposed that SAFPEA’s have the potential for a variety of
low surface energy applications. The SAFPEA’s are viscous materials due to
the presence of highly flexible silicone segments in the backbone chain.
SAFPEA’s with a silicone content above 15% no longer exhibit crystallinity.
Scheme 1.11. Lipase-catalyzed synthesis of silicone fluorinated polyesteramides
(SFAPEPs) by transesterificatioan and amidation of α,-aminopropyl terminated
poly(dimethylsiloxane) (APDMS) and 3,3,4,4,5,5,6,6-octafluoro 1,8-diol (OFOD) with
dimethyl adipate (DEA), respectively. Reprinted from Palsule et al.,94
copyright (2010)
with permission from Elsevier.
The synthesis of poly(amide-co-ester)s (PEAs) via a combination of ring-
opening polymerization and polycondensation in a one-pot reaction has been
performed by Ragupathy et al.88
By using a three-step polymerization process,
as described in Scheme 1.12, molecular weights up to Mn of 17550 g/mol can be
obtained, with melting points of approximately 164 °C.
-
Chapter 1
20
Scheme 1.12. Synthesis of poly(amide-co-ester)s by N435-catalyzed polymerization.
Reproduced from Ragupathy et al., copyright (2012) with permission from Elsevier
1.6. Aim and Outline of the Thesis
The goal of this thesis is to gain a better understanding on enzymatic
polymerization in polyamide syntheses. Therefore, the synthetic activity of two
enzymes (CAL-B and F. solani pisi cutinase) and their effectiveness in the
synthesis of polyamides, using a variety of monomers and techniques, was
investigated.
In chapter 2, the preparation of immobilized F. solani pisi cutinase is
described. Three different methods of immobilization via physical adsorption on
Lewatit beads, cross-linked enzyme aggregates using gluteraldehyde as cross-
linking agent and covalent linkages on modified Eupergit CM beads are
explored. The quality of these immobilized cutinases was assessed through
extensive hydrolytic and synthetic activity studies.
Chapter 3 focuses on the enzymatic synthesis of aliphatic polyamides
(nylon-4,10, nylon-6,10, and nylon-8,10). The enzymatic polymerization was
carried out using different length of diamines (1,4-butanediamine, 1,6-
hexanediamine, or 1,8-diaminooctane) and diethyl sebacate by using
immobilized cutinase or CAL-B as catalyst. The selectivity of enzymes toward
different diamines is studied.
The enzymatic synthesis of lactams is described in chapter 4. By using
different chain length -aminocarboxylic acids (4-aminobutanoic acid, 5-
aminovaleric acid, 6-aminocaproic acid, 8-aminooctanoic acid, and 12-
aminododecanoic acid), lactams with different ring sizes were produced.
-
General introduction
21
In chapter 5, the enzymatic synthesis of aliphatic-aromatic oligoamides is
examined. By combining aliphatic and aromatic monomers, for instance diethyl
sebacate and p-xylylene diamine or dimethyl terephthalate and 1,8-
diaminooctane, in a one-pot enzymatic polymerization, oligoamide containing
both aromatic and aliphatic units were obtained. The conversions, DPmax,
structure, and thermal properties of the oligoamides are examined.
The ring-opening copolymerization of -caprolactone and -lactam is
reported in chapter 6. The structure and thermal analysis of the synthesized
copolymer are studied.
Finally, the main results of the enzymatic synthesis of polyamides are
summarized in chapter 7.
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-
Chapter 2
Immobilization of Fusarium solani
pisi cutinase
Abstract
In this study, the immobilization of Fusarium solani pisi cutinase using three
different methods of immobilization (i.e., hydrophobic interactions on Lewatit
beads, cross-linked enzyme aggregates (CLEA) using glutaraldehyde, or
covalent linkage on modified Eupergit CM beads) are presented. From the
hydrolysis and synthetic activity studies, CLEA cutinase showed higher activity
compared to the other immobilized cutinases.
Part of this chapter is published as:
Stavila, E.; Arsyi, R. Z.; Petrovic, D. M.; Loos, K., Eur. Polym. J. 2013, 49,
834-842.
-
Chapter 2
28
2.1. Introduction
Enzymes are known as biocatalysts and are protein with catalytic activity
that control the rates of metabolic reactions.1 They have also been used as in
vitro catalysts in organic synthesis, chemical industry, as well as pharmaceutical
processes. In chemical reactions, the chiral nature of the enzymes contributes to
high enantioselectivity towards substrates and results in the formation of stereo-
and regio-chemically defined reaction products.2 Thus minimizes the formation
of undesirable byproducts and contributes to enzymes being labelled as
environmentally benign catalysts.1,2
Therefore, enzymes are excellent
alternative catalysts in chemical reactions compared to the traditionally
employed organometallic catalysts.
The hydrolases type enzymes are the most used enzymes in chemical
synthesis due to their broad substrate acceptance and considerable stability in
organic solvents.3 Among the hydrolases, lipases are the most frequently used
and several immobilized lipases are commercially available. Particularly the
enzymes lipase, cellulase, hyaluronidase, and papain have been used in polymer
synthesis.4-6
In addition, other hydrolase enzymes such as cutinases have
recently also been investigated for use as catalysts in polymer synthesis.
Cutinases are hydrolytic enzymes that degrade cutin, which contains
polyester and epoxy fatty acids (C16 and n-C18). This enzyme is known as a
small carboxylic ester hydrolase (Mw 22 kD) that bridges functional properties
between lipases and esterases.7,8
Different from lipases, cutinases do not exhibit
interfacial activation due to the absence of a hydrophobic lid restricting access
to the catalytic site.7 Because of this, cutinases are expected to accept the
substrates easier than lipases. Therefore, in this study the catalysis of cutinase in
the enzymatic synthesis of polyamides was investigated. The enzyme Humicola
insolens cutinase (HIC) has already been used in the synthesis of polyesters.9-12
In our studies, Fusarium solani pisi cutinase was chosen as catalyst for
enzymatic polymerizations as the best of our knowledge the use of this enzyme
in enzymatic polymerization has never been reported.
The limiting factors for using enzymes in polymerization reactions are
generally the relatively low stability of enzymes in organic solvents and the
high reaction temperatures compared to metabolic reactions. One method to
improve stability of enzymes in organic solvents is via enzyme immobilization.
The immobilization of the enzyme not only improves the stability in organic
solvents, but also bestows additional benefits such as: increased enzyme activity
and temperature stability, remarkable long-term stability, increased
-
Immobilization of Fusarium solani pisi cutinase
29
enantioselectivity, enables efficient recovery (by filtration or centrifugation) and
reuse of the immobilized enzymes.13-15
In the enzymatic synthesis of polyester, HIC was used in its immobilized
form.10,12
Immobilizations were achieved via physical interaction on Lewatit
beads10
or covalent linkage on Amberzyme oxirane beads.12
By using a similar
approach Fusarium solani pisi cutinase was immobilized via three methods.
The enzyme was immobilized on Lewatit beads, cross-linked enzyme
aggregates (CLEA) and modified Eupergit CM beads. Lewatit are macroporous
poly(methyl methacrylate) beads that have also been used as solid support for
immobilization of Candida antarctica lipase B. Eupergit CM beads have
oxirane groups that enable covalent linkages in enzyme immobilization. The
immobilization scheme of Fusarium solani pisi cutinase is shown in Scheme
2.1. In addition to the increased solvent and temperature stability, using an
immobilized enzyme for enzymatic polymerizations also facilitates the
separation from the product compared to free enzyme and minimizes any
potential protein contamination.13,14
Scheme 2.1. Simplified schematic immobilization of cutinase via (a) immobilization on
Lewatit beads, (b) cross-linked enzyme aggregates (CLEA), and (c) immobilization on
modified Eupergit CM beads
-
Chapter 2
30
2.2. Experimental Methods
2.2.1. Materials
Fusarium solani pisi cutinase (Novozym 51032) was generously supplied by
Novozymes, Denmark. Lewatit OC VOC 1600 was donated by Lanxess,
Belgium. Ethanol, ammonium hydroxide and calcium hydride were purchased
from Merck. 1,2-Dimethoxyethane, sodium phosphate dibasic, sodium
phosphate monobasic, 4-nitrophenol butyrate, sodium cholate hydrate, -
caprolactone, chloroform-d (CDCl3), Eupergit CM and Lipase acrylic resin
from Candida antarctica (immobilized CAL-B and commercially available as
N435) were purchased from Sigma-Aldrich. A bicinchoninic acid (BCA)
Protein assay kit was purchased from Thermo Scientific. 4-Nitrophenol was
purchased from Fluka. Tetrahydrofuran and glutaraldehyde were obtained from
Acros. Except for toluene and -caprolactone, all chemicals were used without
further purification. Toluene was dried by a solvent purification system (SPS).
-Caprolactone was dried using CaH2 and distilled under reduced pressure.
2.2.2. Immobilization Procedures
Immobilization of Fusarium solani pisi on Lewatit
The Lewatit beads were first activated with ethanol and dried at 50 C under
vacuum for 60 min to remove traces of ethanol.10
Then, beads (0.2 g) were
added to 3 mL cutinase solution (used as received with concentration of 20
mg/mL). The samples were incubated in a shaker at 100 rpm at 4 C for 48 h.
Subsequently, the supernatant was removed and the remaining beads were
washed with sodium buffer phosphate (0.1 M, pH 7.8). The concentration of the
remaining cutinase in the supernatant and the washing solutions was determined
using the bicinchoninic acid (BCA) protein assay, and then compared to the
concentration cutinase before immobilization in order to estimate the amount of
immobilized cutinase on Lewatit beads (enzyme loading). Prior to use, the
icutinase on Lewatit was freeze-dried for 48 h.
-
Immobilization of Fusarium solani pisi cutinase
31
Preparation of CLEA Fusarium solani pisi cutinase
Three milliliters of cutinase stock solution (20 mg/mL) was dissolved in 6
mL sodium phosphate buffer (100 mM, pH 7) according to literature
procedures.16
Subsequently, 18 mL of 1,2-dimethoxylethane and 480 L
glutaraldehyde (25% w/v in water) were added. The mixture was stirred at 4 C
for 17 h. After 17 h, 6 mL of 1,2-dimethoxyethane was added and followed by
centrifugation at 7000 rpm for 20 min. Afterwards, the supernatant was
separated from the residue. The residue was washed with 30 mL of 1,2-
dimethoxyethane, centrifuged, and decanted. The washing step was repeated
three times. The CLEA cutinase was collected and dried in a vacuum oven at 25
C for 1 h.
Immobilization of Cutinase on Modified Eupergit CM
Preparation of Modified Eupergit CM
Eupergit CM was first modified by amination as described in literature.17,18
The amination reaction was performed by adding 0.3 g of Eupergit CM to 20
mL of ammonium hydroxide (25% w/w). The reaction mixture was kept at 50
°C for 24 h and stirred at 100 rpm. After 24 h, the aminated Eupergit CM was
washed with an excess of demineralized water and then dried at 40 °C for 2 h in
the vacuum oven. Subsequently, the aminated Eupergit CM was activated with
glutaraldehyde by addition to 15 mL of sodium phosphate buffer (0.05 M, pH 8)
containing glutaraldehyde (10% w/w) and stirred at 100 rpm at room
temperature for 3 h. After 3 h, the modified Eupergit CM was washed with an
excess of sodium phosphate buffer and dried at 40 °C for 2 h in the vacuum
oven.
Immobilization of Cutinase
The modified Eupergit CM (0.2 g) was added to 3 mL stock solution of
cutinase (concentration of protein = 20 mg/mL). The immobilization was
conducted at 30 C for 24 h and stirred at 100 rpm. After 24 h, the solution was
separated from the beads and the remaining beads were washed with sodium
phosphate buffer (0.1 M pH 7.8). The supernatant and washing solution were
treated with BCA reagent and analyzed on a UV-Vis spectrophotometer at 562
nm to determine the enzyme loading. The resulting immobilized cutinase
(icutinase) on modified Eupergit CM was freeze-dried for 48 h before use.
-
Chapter 2
32
2.2.3. Hydrolysis and Synthetic Assay
Cutinase Hydrolysis Assay
The hydrolytic activity of cutinase was determined by hydrolysis of 4-
nitrophenyl butyrate (p-NPB) to 4-nitrophenol (p-NP). In a standard procedure,
immobilized cutinase (10 mg) was added to 1 mL of sodium phosphate (11.3
mM), tetrahydrofuran (0.43 M), and p-NPB (0.55 mM) in sodium phosphate
buffer (50 mM, pH 7.0). The hydrolytic activity of the cutinase was measured
by UV-Vis spectrophotometer at 399 nm over a time range of 1 minute against
the blank solution.19
The molar absorptivity of p-NP (p-NP) was determined by
the calculation of the slope from the calibration curve and was found to be
7919.2 M-1
cm-1
. The specific activity of cutinase was defined as nmol of p-
nitrophenol for 1 min per gram solid support.
The activity was calculated as shown in equation 2-1 and 2-2:
Which:
Cutinase Synthetic Assay
The synthetic activity of cutinase was determined by ring-opening
polymerization (ROP) of -caprolactone (-CL). One mL of -CL was added to
a 25 mL two-neck flask containing immobilized cutinase (10 mg or 10 L for
free cutinase). The reactions were carried out at 70 C for 24 h at 100 rpm,
under N2 atmosphere. The monomer conversion and the degree of
polymerization were determined by 1H NMR measurements using CDCl3 as
solvent.
The conversion of -CL was calculated by comparing the area signal of CH2
next to the carbonyl group of the polymer backbone (I1) at 4.09 ppm and of the
monomer (Ia) at 4.19 ppm. The ratio of the area from the signal of CH2 next to
the carbonyl group of the polymer backbone to the total area from the signal of
the CH2 next to the carbonyl group of the polymer backbone and in the
monomer was determined as monomer conversion, as depicted in Figure 2.1,
see formula.20
-
Immobilization of Fusarium solani pisi cutinase
33
Figure 2.1. Monomer conversion calculated from the
1H NMR spectrum of the reaction
mixture after 1 h of ROP of -CL at 70 °C using N435 as catalyst
2.2.4. Instrumental Methods
Attenuated Total Reflectance-Fourier Transform Infrared (ATR FT-IR)
measurements were carried out on a Bruker IFS88 FT-IR spectrometer. UV-Vis
measurements were performed on a Spextra Max M2 spectrophotometer. 1H
NMR measurements were performed on a 400 MHz Varian VXR apparatus,
with CDCl3 as solvent.
2.3. Results and Discussion
2.3.1. Immobilization of Cutinase on Lewatit Beads
In the immobilization experiments on Lewatit, cutinase was attached to the
beads by physical adsorption/non-covalent linkages. From enzyme loading
determination, the amount of attached cutinase varied between 127–143 mg
cutinase per gram Lewatit beads.
Further characterization of the icutinase on Lewatit was carried out by ATR
FT-IR measurement. Figure 2.2 shows the ATR FT-IR spectra of the Lewatit
beads before and after immobilization. After immobilization there are three
additional peaks at 1653, 1541, and 3286 cm-1
, that can be attributed to amide I
-
Chapter 2
34
(C=O), amide II (N–H) and intermolecular hydrogen bonding, respectively,
which confirms successful immobilization of the cutinase.
Figure 2.2. ATR FT-IR spectra of (a) Lewatit beads and (b) Lewatit beads with
immobilized cutinase
In order to determine icutinase on Lewatit’s activity in toluene, hydrolysis
assay tests were performed after stirring in toluene at 70 °C for 4 days and
compared to fresh icutinase on Lewatit. The results from the hydrolysis assay
are presented in table 2.1. The hydrolytic activity decreased from 13 to 7 nmol
of p-NP min-1
mg-1
. This decrease in hydrolytic activity of icutinase on Lewatit
means that less cutinase leach out from the Lewatit beads and immobilization
able to inhibit denaturation of cutinase. Therefore, icutinase on Lewatit is able
to maintain its activity after stirring in toluene.
Table 2.1. Hydrolytic activity of immobilized cutinase on Lewatit
Enzyme Unit activity
(nmol of p-NP min-1
)
Specific activity
(nmol of p-NP min-1
mg-1
)
iCutinase on Lewatita)
132 13
iCutinase on Lewatita)
after
stirring 4 d in toluene 78 ± 17 7 ± 2
a) Enzyme loading was 127 mg/g.
2.3.2. Immobilization of Cutinase by CLEA
Immobilization of Fusarium solani pisi via the CLEA method was
performed via a simple cross-linking process using glutaraldehyde as cross-
linker agent. Around 0.1 g CLEA was isolated from 3 mL free cutinase solution
(20 mg/mL). The cross-linking was achieved via the reaction of a primary
-
Immobilization of Fusarium solani pisi cutinase
35
amine (from lysine for example) with the carbonyl from glutaraldehyde, which
react rapidly under neutral pH.21
In reactions Scheme 2.2, the simplified
reaction of glutaraldehyde with primary amine is shown. However, the reaction
is not as straightforward because glutaraldehyde may form oligomeric (dimer,
trimer, tertramer, etc.) or cyclic species apart from its monomeric form and all
of them are in the equilibrium state in an aqueous solution.21
Scheme 2.2. Schematic presentation of reaction scheme of cross-linking of cutinase and
glutaraldehyde
ATR FT-IR spectra of free cutinase and cutinase after cross-linking are
shown in Figure 2.3a and b. By comparing the spectra before and after cross-
linking, we observed three new peaks at 861, 978, and 1049 cm-1
after the
enzyme was cross-linked, as indicated in Figure 2.3b. The peaks at 861 and 978
cm-1
can be attributed to the quarternary pyridium compound from the aromatic
vibrational stretch (C–H). It is also likely that the amino acid of the free
cutinase can be observed in this region, because the CLEA after cross-linking
precipitates, whereas the free cutinase remains in solution. Furthermore, the
peak at 1049 cm-1
originates from the stretching vibration of an aliphatic amine
(C–N). This peak indicates that the cross-link between glutaraldehyde and the
amine group in the cutinase was successfully formed.
-
Chapter 2
36
Figure 2.3. ATR FT-IR spectra of (a) free cutinase and (b) CLEA cutinase
2.3.3. Immobilization on Modified Eupergit CM Beads
An important feature of the Eupergit CM beads is that they can be used as
solid support in immobilization of enzyme without further modification steps.
The oxirane group present in these beads is crucially important in the process of
immobilizing an enzyme and allows for covalent binding after physical
adsorption of the enzyme on the beads because of the reaction of the enzyme
with the epoxy groups (oxirane groups).14
. However, we observed no cutinase
immobilized on the unmodified Eupergit CM, because no enzyme loading could
be determined after the immobilization. Therefore, the Eupergit CM beads were
modified to include a spacer to allow immobilization of cutinase. The
modification of Eupergit CM beads and the subsequent immobilization of
cutinase are shown in Scheme 2.3.
Scheme 2.3. Schematic reaction of the modification of Eupergit CM and the
immobilization of cutinase on modified Eupergit CM where (a) is the amination step,
(b) the activation using glutaraldehyde, and (c) the immobilization of cutinase
-
Immobilization of Fusarium solani pisi cutinase
37
The modification was carried out by first aminating the oxirane group and
subsequent activation using glutaraldehyde, as shown in Scheme 2.3a and b. By
comparing the ATR FT-IR spectra, we observed the difference before and after
modification steps, as shown in Figure 2.4. In Figure 2.4a the oxirane group
from Eupergit CM beads can be observed clearly at 871 (stretching C–O–C) and
909 (stretching C–O) cm-1
. After the amination step, the peaks belonging to
oxirane were no longer detected, as can be seen in Figure 2.4b. The final step in
the modification was performed by activation using gluteraldehyde. The ATR
FT-IR spectrum in Figure 2.4c shows no significant different with the spectrum
after amination step. Because of the formation of an amine, a new peak was
expected to appear around 1025–1200 cm-1
, which belongs to the stretching
band of C–N (amine). It is likely the amine peak was hidden under the main
peak at 1106 cm-1
, which is probably from stretching C–O of ester group of
Eupergit CM beads. The modified Eupergit CM was further used for solid
support in immobilization of cutinase.
Figure 2.4. ATR FT-IR spectra of Eupergit CM beads (a) before modification, (b) after
amination, and (c) after activation using glutaraldehyde
The immobilization of cutinase on modified Eupergit CM beads was
achieved via hydrophobic interaction and the formation of a covalent bond after
reaction between the glutaraldehyde and the amine from cutinase. The enzyme
loading after immobilization procedure and purification was 77 mg/g. Further
characterization of cutinase attached on modified Eupergit CM was carried out
by ATR FT-IR measurements. From the ATR FT-IR spectrum of modified
Eupergit CM beads after immobilization (Figure 2.5b), a new peak was
-
Chapter 2
38
observed at 1026 cm-1
and comes from the stretching band of C–N (amine).
Thus, the presence of new peak proves that cutinase attached to the modified
Eupergit CM bead.
Figure 2.5. ATR FT-IR spectra of modified Eupergit CM beads (a) before
immobilization of cutinase and (b) after immobilization of cutinase (enzyme loading =
77 mg/g)
Because the immobilization on Eupergit CM occurs via the formation of
covalent bonds as well as physical interactions, we would expect to observe
higher enzyme loading than immobilized cutinase on Lewatit. However, the
immobilization of cutinase on modified Eupergit CM resulted in lower enzyme
loading values. This is because Eupergit CM beads (particle size of 50–300 m)
have smaller particle size compared to Lewatit beads (315–1000 m).
Therefore, less cutinase can be immobilized on Eupergit CM beads.
2.3.4. Activity Comparison of Immobilized and Free cutinase
The hydrolytic and synthetic activities were examined for icutinase on
Lewatit, the CLEA-cutinase, icutinase on modified Eupergit CM, and free
cutinase. The hydrolytic activity was investigated via a hydrolysis assay, where
the hydrolysis of 4-nitrophenyl butyrate (p-NPB) to 4-nitrophenol (p-NP) was
analyzed. In the synthetic assay, the ring-opening polymerization (ROP) of -
caprolactone was performed. The results of both tests are summarized in Table
2.2. The hydrolysis assay revealed higher activity of free cutinase compared to
cutinase after immobilization. The icutinase on modified Eupergit CM showed
the lowest activity, because the enzyme loading of icutinase on modified
-
Immobilization of Fusarium solani pisi cutinase
39
Eupergit CM was lower compared to icutinase on Lewatit beads, therefore
icutinase on modified Eupergit CM showed lower hydrolytic activity compared
to the other immobilized cutinase.
The synthetic assay illustrated that the free cutinase has better catalytic
activity than any of the immobilized cutinases. It is likely that some of the
active sites are no longer accessible to solvent or monomer because they are
blocked by the solid support or cross-linked with glutaraldehyde compromising
their activity. Although the synthetic and hydrolytic activities are affected by
the immobilization, the immobilized cutinases were used in the synthesis
reactions described in the following chapters, because they facilitate separation
from the reaction mixture, as well as minimizing protein contamination of the
product.13,14
From Table 2.2, it becomes obvious that CLEA seems to be the better
immobilization method for cutinase compared to icutinase on Lewatit or
Eupergit CM. This behavior is in line with to literature,14
because enzymes in
CLEA preparations are usually more concentrated. Furthermore, immobilization
on macroporous beads such as Lewatit beads or modified Eupergit CM leads to
dilution of activity, due to the large amount of non-catalytic solid support.
Table 2.2. Enzyme loading and activity of free and immobilized cutinase
Sample Hydrolytic activity
(nmol of p-NP min-1
)
Amount of
sample in
synthetic activity
Poly(-CL)
̅̅ ̅̅ Monomer
conversion
(%)
Free cutinasea)
124 ± 25b)
10 L 3 34 ± 8
iCutinase on Lewatitc)
108 ± 19 10 mg 1 4 ± 1
CLEA cutinase 124 ± 12 10 mg 3 19 ± 4
iCutinase on modified
Eupergit CMd)
11 10 mg 1 5
a) Enzyme concentration was 20 mg/mL.
b) Hydrolysis assay was carried out using 20 µL of free cutinase.
c) Enzyme loading was 135 ± 8 mg/g.
d) Enzyme loading was 77 mg/g.
2.4. Conclusion
Immobilization of Fusarium solani pisi cutinase by physical interaction on
Lewatit beads, cross-linked enzyme aggregates (CLEA) or covalent linkage on
modified Eupergit CM beads has been performed. Immobilization of cutinase
using the CLEA method resulted in better catalytic activity than immobilization
-
Chapter 2
40
on Lewatit or modified Eupergit CM beads. Therefore, in this case, the
immobilization by CLEA method was the preferred method for Fusarium solani
pisi cutinase immobilization.
For immobilization on modified Eupergit CM, low enzyme loading and
lower reproducibility of the immobilization methods resulted in relatively
complicated immobilization. Despite the fact that free cutinase has higher
hydrolytic and synthetic activity than the immobilized cutinases, only the
immobilized cutinase on Lewatit and the CLEA-cutinase will be used in the
enzymatic polymerization reactions described in the following chapters.
2.5. References
1. Akiyama, A.; Bednarski, M.; Kim, M.-J.; Simon, E. S.; Waldman, H.;
Whitesides, G. M. CHEMTECH 1988, october, 627-634.
2. Koeller, K. M.; Wong, C.-H. Nature 2001, 409, 232-240.
3. Gotor-Fernández, V.; Vicente, G. Use of Lipases in Organic Synthesis.
In Industrial Enzymes, Polaina, J.; MacCabe, A., Eds. Springer
Netherlands: 2007; pp 301-315.
4. Kobayashi, S.; Uyama, H. Enzymatic Polymerization to Polyesters. In
Biopolymers Online, Wiley-VCH Verlag GmbH & Co. KGaA: 2005.
5. van der Vlist, J.; Loos, K. Enzymatic Polymerizations of
Polysaccharides. In Biocatalysis in Polymer Chemistry, Wiley-VCH
Verlag GmbH & Co. KGaA: 2010; pp 211-246.
6. Cheng, H. N. Enzyme-Catalyzed Synthesis of Polyamides and
Polypeptides. In Biocatalysis in Polymer Chemistry, Wiley-VCH
Verlag GmbH & Co. KGaA: 2010; pp 131-141.
7. Carvalho, C. M.; Aires-Barros, M. R.; Cabral, J. M. S. Electron. J.
Biotechnol. 1998, 1, 160-173.
8. Egmond, M. R.; de Vlieg, J. Biochimie 2000, 82, 1015-1021.
9. Gross, R. A.; Ganesh, M.; Lu, W. Trends Biotechnol. 2010, 28, 435-
443.
10. Hunsen, M.; Abul, A.; Xie, W.; Gross, R. Biomacromolecules 2008, 9,
518-522.
11. Hunsen, M.; Azim, A.; Mang, H.; Wallner, S. R.; Ronkvist, A.; Xie,
W.; Gross, R. A. Macromolecules 2007, 40, 148-150.
12. Feder, D.; Gross, R. A. Biomacromolecules 2010, 11, 690-697.
-
Immobilization of Fusarium solani pisi cutinase
41
13. Miletic, N.; Nastasovic, A.; Loos, K. Bioresour. Technol. 2012, 115,
126.
14. Sheldon, R. A. Adv. Synth. Catal. 2007, 349, 1289-1307.
15. Mateo, C.; Palomo, J. M.; Fernandez-Lorente, G.; Guisan, J. M.;
Fernandez-Lafuente, R. Enzyme Microb. Technol. 2007, 40, 1451-1463.
16. López-Serrano, P.; Cao, L.; van Rantwijk, F.; Sheldon, R. A.
Biotechnol. Lett. 2002, 24, 1379-1383.
17. Bilici, Z.; Camli, S. T.; Unsal, E.; Tuncel, A. Anal. Chim. Acta 2004,
516, 125-133.
18. Miletić, N.; Rohandi, R.; Vuković, Z.; Nastasović, A.; Loos, K. React.
Funct. Polym. 2009, 69, 68-75.
19. Almeida, C. F.; Cabral, J. M. S.; Fonseca, L. s. P. Anal. Chim. Acta
2004, 502, 115-124.
20. Schwab, L. W.; Kroon, R.; Schouten, A. J.; Loos, K. Macromol. Rapid
Commun. 2008, 29, 794-797.
21. Migneault, I.; Dartiguenave, C.; Bertrand, M. J.; Waldron, K. C.
BioTechniques 2004, 37, 790-802.
-
Chapter 2
42
-
Chapter 3
Fusarium solani pisi cutinase-
catalyzed synthesis of polyamides
Abstract
Polyamides, or nylon, are widely used in fiber and engineering plastic materials,
due to their good mechanical and thermal properties. Synthesis of oligomers
from nylon-4,10, nylon-6,10, and nylon-8,10 were performed via
polycondensation of diamine (1,4-butanediamine, 1,6 hexanediamine, or 1,8-
diaminooctane) and diester (diethyl sebacate). These reactions were catalyzed
by immobilized cutinase from Fusarium solani pisi on Lewatit beads, cutinase
in the form of cross-linked enzyme aggregates (CLEA), or by immobilized
Lipase B from Candida antarctica (CAL-B). The highest maximal degree of
polymerization (DPmax), up to 16, can be achieved in the synthesis of nylon-8,10
catalyzed by CLEA in diphenyl ether at 70 C. By performing a reaction at
cutinase optimal temperature (70 C), CLEA cutinase in the synthesis of nylons
shows good catalytic activity, like CAL-B.
Part of this chapter is published as:
Stavila, E.; Arsyi, R. Z.; Petrovic, D. M.; Loos, K., Eur. Polym. J. 2013, 49,
834-842.
-
Chapter 3
44
3.1. Introduction
Polyamides, also known as nylon, are widely used in fiber and engineering
plastic materials, due to their good mechanical and thermal properties.1
Synthesis of nylons can be accomplished via ring-opening polymerization2 and
polycondensation.3 Industrial synthesis of nylon is conducted at high
temperatures, which leads to thermal degradation and results in undesired
polymer products.4 On the other hand, alternative options for synthesizing
polymers at relative low temperatures can be achieved via enzymatic
polymerization. Enzymatic polymerizations are known as environmentally
friendly reactions, as they can be carried out in relatively low temperatures, and
they use enzymes as catalysts.5-10
Candida antarctica lipase B (CAL-B) is a highly used enzyme in
biocatalysis. CAL-B is commercially available as N435, which is physically
immobilized CAL-B within macroporous poly(methyl methacrylate) resin
(known as Lewatit OC VOC 1600).11
CAL-B is known to have high catalytic
activity over a broad range of esters, amides, and thiols.12
CAL-B has been used
in esterification and transesterification,13
aminolysis,14,15
and synthesis of
polyesters,5,7,16
polyamides,17-19
and the poly(amide-co-ester).20
Here, we present, for the first time, the synthesis of polyamides catalyzed by
Fusarium solani pisi cutinase. Cutinases are hydrolytic enzymes that degrade
cutin, which contains polyester and epoxy fatty acids (C16 and n-C18). This
enzyme is known as a small carboxylic ester hydrolase (Mw 22 kD) that bridges
functional properties between lipases and esterases.21
Like CAL-B, Fusarium
solani pisi cutinase belongs to the / hydrolase family, which has a catalytic
triad in Ser120, His188, and Asp175. It is known, that the act