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Hydrolase-catalyzed synthesis of polyamides and polyester amidesStavila, Erythrina

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Hydrolase-catalyzed synthesis of

polyamides and polyester amides

Erythrina Stavila

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

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

Promotor

Prof. dr. K. Loos

Beoordelingscommissie

Prof. dr. N. Bruns

Prof. dr. G. Fels

Prof. dr. F. Picchioni

Untuk Keluargaku Tersayang

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




3.4. Conclusion 56

3.5. References 57

Chapter 4 Synthesis of lactams using enzyme-catalyzed aminolysis

59




4.4. Conclusion 69

4.5. References 69

Chapter 5 Enzyme-catalyzed synthesis of aliphatic-aromatic oligoamides

71




5.4. Conclusion 89

5.5. References 90

Chapter 6

Lipase-catalyzed ring-opening copolymerization of -caprolactone

and -lactam

93



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

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.

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.


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

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


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

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


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

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


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

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.


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

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-


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

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


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


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


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.


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.


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


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


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


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


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


6. Cheng, H. N. Enzyme-Catalyzed Synthesis of Polyamides and

Polypeptides. In Biocatalysis in Polymer Chemistry, Wiley-VCH


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.


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

university of groningen hydrolase-catalyzed synthesis of ......enzyme-catalyzed synthesis of...

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