hydrolysis of l-arginine â€“ chemical and enzymatic catalysis

Hydrolysis of L-Arginine –

Chemical and Enzymatic Catalysis

Pedro Miguel Cabral Campello Duarte Turras

Dissertação para obtenção do Grau de Mestre em

Engenharia Biológica

Júri

Presidente: Professor Luís Joaquim Pina da Fonseca (IST)

Orientadores: Professor Luís Joaquim Pina da Fonseca (IST)

Professor Maurice Franssen (WUR)

Vogais: Doutor Pedro Fernandes (IST)

Professor José António Leonardo dos Santos (IST)

Setembro 2008

i

Acknowledgments

I would like to thank my supervisor, Paul Könst, for teaching, supporting and helping

me through the course of this project.

I would also like to thank Professor Maurice Franssen for his guidance and the

opportunity to accomplish such gratifying work in a great research group.

Finally, I would like to thank everyone at the Valorisation of Plant Production Chains

group at Wageningen University. Not only was their advice and help essential during the

course of the project – a special word to Dr. Elinor Scott and Alniek van Zeeland – but I will

also keep everything I have learned during progress meetings, informal meetings and talks.

iii

Resumo

O esgotamento das reservas de combustíveis fosseis e a crescente preocupação

com as emissões de CO2 levaram à procura de uma fonte alternativa para obtenção

químicos funcionalizados em larga escala. O projecto N-ergy tem como principal objectivo a

utilização de resíduos agrícolas como principal matéria-prima na produção simultânea de

etanol e químicos azotados. Um dos passos intermediários no processo previsto é a

conversão de L-arginina em L-ornitina.

A hidrólise de L-arginina em L-ornitina e ureia pode ser catalisada quimicamente ou

enzimaticamente. A catálise quimica da reacção pode ser conseguida na presença de

condições fortemente ácidas ou alcalinas. A biocatálise da reacção pode ser conseguida

através da enzima arginase (EC 3.5.3.1).

A hidrotermólise da L-arginina em condições alkalinas levou à produção de L-ornitina

e de vários produtos secundários. O rendimento máximo de obtenção de L-ornitina foi de

15.2% a 150°C com um pH inicial ajustado a 12.0 com hidróxido de sódio. A adição de um

catalisador básico sólido, zeólito NaY, mostrou um efeito catalítico limitado mesmo a

elevadas concentrações. A adição de sais de diferentes metais não influenciou a reacção.

Arginase de Bacillus subtilis foi imobilizada com sucesso em três suportes activados

com grupos epóxi – Seapabeads EC-HFA, Sepabeads EC-EP e Eupergit C 250 L. Após uma

hora de incubação à temperatura ambiente na presença de cada suporte nenhuma arginase

foi detectada no sobrenadante. A ligação covalente aos três suportes testados não levou a

um aumento da estabilidade térmica da arginase e provocou uma redução na actividade

catalitica de 40% a 60%.

Palavras-chave: hidrólise da L-arginina; zeolito NaY; arginase; immobilização de enzimas.

v

Abstract

The depletion of fossil feedstocks and growing concern over CO2 emissions has led

to the search for an alternative source for bulk functionalized chemicals. The N-ergy project

has the ultimate aim of utilizing agricultural waste streams as the main raw material for the

simultaneous production of ethanol and bulk nitrogen-functionalized chemicals. One of the

intermediate steps in the projected process is the conversion of L-arginine to L-ornithine.

The hydrolysis of L-arginine to L-ornithine and urea can be chemically or

enzymatically catalysed. The chemical catalysis of the reaction can be achieved in the

presence of strong acidic or alkaline conditions. The biocatalysis of the reaction can be

achieved by the enzyme L-arginase (EC 3.5.3.1).

The hydrothermolysis of L-arginase in alkaline conditions led to the production of L-

ornithine and numerous secondary products. The maximum L-ornithine yield obtained was of

15.2% at 150°C with initial pH adjusted to 12.0 with sodium hydroxide. The addition of a solid

basic catalyst, NaY zeolite, showed limited catalytic effect even at high concentrations. The

addition of various metal salts did not influence the reaction.

Bacillus subtilis arginase was successfully immobilized in three different epoxy-

activated supports – Seapabeads EC-HFA, Sepabeads EC-EP and Eupergit C 250 L. After

one hour of incubation at room temperature in the presence of each support no arginase was

detected in the supernatant. The covalent-binding to the three tested supports did not lead to

a significant increase in arginase’s thermal stability and led to a decrease in catalytic activity

(recovered activity of 40%-60%).

Key words: L-arginine hydrolysis; NaY zeolite; arginase; enzyme immobilization.

vii

Table of Contents

1. Introduction 1

1.1. Motivation and Background 1

1.1.1. Biomass as a Source of Bulk Chemicals 1

1.1.2. Petrochemical Approach vs Bio-Refinery Approach 2

1.1.3. The N-ergy Project 3

1.1.4. Arginine Conversion 4

1.1.5. Product Applications 5

1.2. Chemical Catalysis 7

1.2.1. Chemically Catalyzed Hydrolysis of Arginine 7

1.2.2. Heterogeneous Catalysis – the use of Zeolites 9

1.3. Enzymatic Catalysis 10

1.3.1. Enzymatic Hydrolysis of L-Arginine – Arginase 10

1.3.2. Industrial Biocatalysis 14

1.3.3. Biocatalyst Immobilization 15

1.4. Aim of this Study 18

2. Chemical Hydrolysis of L-Arginine 21

2.1. Materials and Methods 21

2.1.1. Reagents 21

2.1.2. Solutions 22

2.1.3. Equipment 22

2.1.4. Analytical Techniques 23

2.1.5. Hydrothermolysis Experiments 24

2.1.6. Metal Salt Catalysis Experiments 25

2.1.7. Zeolite Catalysis Experiment 25

2.2. Results and Discussion 26

2.2.1. Hydrothermolysis Experiments 26

2.2.2. Metal Salt Catalysis Experiments 31

2.2.3. Zeolite Catalysis Experiment 32

2.2.4. Analytical Methods 32

3. Enzymatic Hydrolysis of L-Arginine 35

3.1. Materials and Methods 35

3.1.1. Bacillus Subtilis Arginase 35

3.1.2. Epoxy-activated Supports 35

3.1.3. Reagents 35

3.1.4. Solutions 36

3.1.5. Equipment 36

3.1.6. Analytical Techniques 37

3.1.7. Preparation of Arginase Stock Solution 38

viii

3.1.8. Immobilization of Arginase in Different Epoxy-activated Supports 38

3.1.9. Thermal Stability of Immobilized Arginase 40

3.2. Results and Discussion 40

3.2.1. Characterization of the Arginase Stock Solution 40

3.2.2. Immobilization of Arginase in Epoxy-activated Supports 42

3.2.3. Recovered Activity of Immobilized Arginase 45

3.2.4. Thermal Stability of Immobilized Arginase 46

3.2.4. Analytical Methods 48

4. Conclusions and Future Perspectives 51

5. References 53

Appendix A – Gradient Curves 57

Appendix B – NMR Spectra of L-Arginine 58

Appendix C – NMR Spectra of L-Ornithine 60

Appendix D – NMR Spectra of 3-Aminopiperid-2-one 62

Appendix E – NMR Spectra of Putrescine 64

Appendix F – NMR Spectra of 30 h Reaction Mixture 66

Appendix G – Certificates of Analysis 68

Appendix H – Recovered Support Masses 70

ix

Abbreviations and Symbols

CGP Cyanophycin Granule Peptide

DEPT Distortionless Enhancement by Polarization Transfer

DMF Dimethylformamide

HPLC High Performance Liquid Chromatography

NMR Nuclear Magnetic Resonance

TSP 2,2,3,3-d(4)-3-(trimethylsilyl)propionic Acid

Tables Index

Table 1.1. Cost breakdown of functionalized and non-functionalized bulk chemicals based on oil at $40 a barrel (adapted from Sanders et al.

4). 2

Table 1.2. Thermal properties of Nylon-6,6 and Stanyl® 11

. 6

Table 1.3. Techniques for arginase immobilization described in literature. The different techniques are listed in chronological order, from the oldest work to the most recent. 13

Table 1.4. Advantages and limitations associated to the use of immobilized enzymes (adapted from Cabral et al.

42) 15

Table 2.1. Reagents utilized during the course of the chemical hydrolysis of arginine experiments. 20

Table 2.2. Solutions prepared during the course of the chemical hydrolysis of L-arginine experiments. 21

Table 2.3. Gradient of eluents applied during HPLC analysis. 23

Table 3.1. Reagents utilized during the course of the enzymatic hydrolysis of L-arginine experiments. 33

Table 3.2. Solutions prepared during the course of the enzymatic hydrolysis of L-arginine experiments. 34

Table 3.3. Gradient of eluents applied during HPLC analysis. 35

Table H.1. Initial and recovered masses of the three tested epoxy-activated supports, including duplicates, during the immobilization and epoxy groups blockage steps. 67

x

Figures Index

Figure 1.1. Production of functionalized chemicals from naphta and from biomass (Sanders et al.

4). 2

Figure 1.2. Conversion of biomass to N-functionalized chemicals and ethanol – the N-ergy project. 4

Figure 1.3. Conversion of arginine to 1,4-butanediamine through hydrolysis to ornithine and decarboxylation. 5

Figure 1.4. Comparison of the petrochemical and bio-based approaches for 1,4-butane diamine bulk production (adapted from Sanders et al.

4). 5

Figure 1.5. Synthesis of nylon-4,6 from 1,4-butanediamine and adipic acid. 6

Figure 1.6. Acid catalyzed hydrolysis of arginine. 7

Figure 1.7. Alkali catalyzed hydrolysis of arginine 8

Figure 1.8. Thermohydrolysis of urea: (a), reversible conversion of urea to cyanate and ammonia; (b), hydrolysis of cynate to ammonia and carbon dioxide. 8

Figure 1.9. Topology diagram of rat liver arginase. Relative location of metal ligands is indicated by grey circles (adapted from Kanyo et al.)

28. 11

Figure 1.10. Proposed mechanism of rat liver arginase-catalysed arginine hydrolysis by metal-activated solvent

28. The α-amino and α –carboxylate groups are omitted

for clarity. 12

Figure 1.11. Methods of immobilization of biocatalysts. 16

Figure 1.12. Mechanism of immobilization of proteins on epoxy-activated Supports. The covalent reaction between soluble enzyme and epoxy support is extremely slow, but the previous adsorption to the support allows a faster covalent reaction (adapted from Mateo et al.

43). 17

Figure 1.13. Functional groups of Sepabeads® EC-EP and Sepabeads

® EC-HFA

supports45

. 18

Figure 2.1. Parr® Series 500 Multiple Reactor System with 4871 Process Controller. 22

Figure 2.2. Time course of L-arginine consumption under different experimental conditions. The percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the reaction mixture at t=0. Error bars calculated using standard deviation of the two duplicate experiments. 26

Figure 2.3. Time course of L-ornithine formation under different experimental conditions. The percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the reaction mixture at t=0. Error bars calculated using standard deviation of the two duplicate experiments. 27

xi

Figure 2.4. Example of a typical HPLC chromatogram obtained after heating a L-arginine solution for 20h in the described experimental settings. The identified peaks correspond to: (A) L-aspartic acid (internal standard); (B) hydrolyzed excess derivatization reagent; (C) L-citrulline; (D) L-arginine; (E) L-ornithine; (F) ammonia; (G) 3-aminopiperid-2-one; (H) retention time of putrescine elution(none was detected in the samples). All unidentified peeks correspond to impurities originating either from the solutions utilized or from the HPLC system. 28

Figure 2.5. Evolution of the pH during the course of the reaction (125°C, initial pH of 12) and its comparison with ornithine concentration. Error bars were omitted for clarity. 29

Figure 2.6. Effect of the presence of different metal salts (equimolar concentrations – 25 mM) on the L-arginine thermohydrolysis reaction. The percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the reaction mixture at t=0. 30

Figure 2.7. Effect of different NaY zeolite concentrations on the L-arginine thermohydrolysis reaction. The percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the reaction mixture at t=0. Error bars calculated using standard deviation of the two duplicate experiments. 31

Figure 2.8. Detail from a HPLC chromatogram for a 25 mM 3-aminopiperid-2-one solution. The identified peaks correspond to: (O) L-ornithine; (A) ammonia; (L) 3-aminopiperid-2-one. All unidentified peeks correspond to impurities originating either from the solutions utilized or from the HPLC system. 32

Figure 3.1. Cole-Parmer Roto-Torque model 7637-10 Heavy Duty Rototator. 34

Figure 3.2. SDS-PAGE gel of the original arginase solution. Lane (A) corresponds to a 100X dilution while lane (B) corresponds to a 200X dilution. Lane (M) contains the molecular markers identified with the corresponding molecular weights in Daltons. 39

Figure 3.3. Protein concentration in the supernatant during the course of arginase immobilization in different epoxy-activated supports. The percentages are based on the initial (t=0) protein concentration. Error bars calculated using standard deviation of the two duplicate experiments. 40

Figure 3.4. Arginase activity in the supernatant during the course of arginase immobilization in different epoxy-activated supports. The percentages are based on the initial (t=0) protein concentration. Error bars calculated using standard deviation of the two duplicate experiments. 41

Figure 3.5. Protein content in the filtration supernatant after each step of the beads washing procedure. Error bars calculated using standard deviation of the two duplicate experiments. 42

Figure 3.6. Recovered activity of arginase immobilized in different epoxy supports. Percentages are calculated by comparison with the soluble form. Error bars calculated using standard deviation of the two duplicate experiments. 43

Figure 3.7. Evolution of residual activity of arginase immobilized in different epoxy supports compared to soluble arginase during the course of 24 hours incubation at 60°C. The percentages are based on the initial (t=0) enzyme activity. 45

xii

Figure 3.8. Example of a typical HPLC chromatogram obtained for an activity assay sample. 46

Figure A.1. Different gradient curves identified by the input number. 54

Figure B.1. 13

C-NMR spectrum of L-arginine. 55

Figure B.2. DEPT spectrum of L-arginine. 55

Figure B.3. 1H-NMR spectrum of L-arginine. 56

Figure C.1. 13

C-NMR spectrum of L-ornithine. 57

Figure C.2. DEPT spectrum of L-ornithine. 57

Figure C.3. 1H-NMR spectrum of L-ornithine. 58

Figure D.1. 13

C-NMR spectrum of 3-aminopiperid-2-one. 59

Figure D.2. DEPT spectrum of 3-aminopiperid-2-one. 59

Figure D.3. 1H-NMR spectrum of 3-aminopiperid-2-one. 60

Figure E.1. 13

C-NMR spectrum of putrescine. 61

Figure E.2. DEPT spectrum of putrescine. 61

Figure E.3. 1H-NMR spectrum of putrescine. 62

Figure F.1. 13

C-NMR spectrum of 30h reaction mixture. 63

Figure F.2. DEPT spectrum of 30h reaction mixture. 63

Figure F.3. 1H-NMR spectrum of 30h reaction mixture. 64

Introduction

1

1. Introduction

1.1. Motivation and Background

1.1.1. Biomass as a Source of Bulk Chemicals

Oil and other fossil fuels still are the main sources for energy, transport fuels and

(bulk) carbon-based chemicals, but the need for an alternative is undeniable. The depletion

of fossil feedstocks, the increasing oil and transport fuels prices, together with the growing

concern over climate changes and other consequences of CO2 emissions, has led to the

search for a cheap and environmentally friendly alternative.

Focusing on the production of chemicals, the replacement of fossil feedstocks with

CO2 neutral biomass offers a wide array of advantages: it’s a renewable resource which is

currently being produced in large amounts (170,000 million tones per annum accordingly to

1992 estimates by Eggersdorfer et al.1); it’s a relatively cheap resource – protein rich waste

streams are generated by plenty of industries, including bio-fuel production processes; the

use of biomass instead of fossil fuels would considerably reduce greenhouse gases

emissions. Moreover, it should be noted that in the long term, after the complete exhaustion

of fossil resources reservoirs, biomass will be the only available raw material for the bulk

production of organic carbon-based chemicals.

Bio-refinery can be defined as the fractionation of biomass into components that, after

further transformation and separation, can be used as final end products. When producing

chemicals through bio-refinery two options can be considered: the development of “new”

chemicals or the production of chemicals similar to the traditionally obtained from the

petrochemical industry. In the first case the focus is mainly on the use of carbohydrates as

raw materials in the production of new polymers, such as thermoplastic resins2 and multi-

application biodegradable additives3. While this option can provide chemicals and materials

with unique structures and properties, the introduction of bio-based alternative products

usually requires the re-engineering and optimization of whole production chains and implies a

considerable financial risk. On the other hand, the bio-based synthesis of existing chemicals

can profit from existing infrastructures. These structures – utilized in the production of bulk

chemicals and materials based on fossil fuels – are already highly optimized and do not

require large capital injections. Thus, in the short to mid-term it is expected that there will be a

strong investment in the development of bio-based bulk chemicals identical to those of

petrochemical origin that can be easily integrated in existing processes.

Introduction

2

1.1.2. Petrochemical Approach vs Bio-Refinery Approach

Despite its increasing prices, oil is still a reliable and inexpensive source of carbon-

based chemicals. However, the absence of functionalized substances in oil derivatives like

naphta, which is the main raw material used in the production of many chemicals, makes bio-

refinery a solid alternative to the petrochemical approach in the production of functionalized

chemicals.

Figure 1.1. Production of functionalized chemicals from naphta and from biomass (Sanders et al.4).

As shown in figure 1.1, only non-functionalized chemicals (olefins, parafins, etc.) can

be produced from naphtha without major enthalpy changes that require a significant heat

transfer. The preparation of functionalized chemicals from simple molecules, such as

ethylene, often implies the use of large amounts of energy, additional process steps,

dangerous working conditions (high temperatures, high pressures, corrosive/toxic

substances) and large amounts of various reagents (ammonia, chlorine). The differences in

production costs of non-functionalized and (O-,N-)functionalized chemicals are illustrated in

table 1.1.

Table 1.1. Cost breakdown of functionalized and non-functionalized bulk chemicals based on

oil at $40 a barrel (adapted from Sanders et al.4).

Cost type Non-functionalized (€/ton) Functionalized (€/ton)

Raw materials 200 650

Capital 300-500 400-650

Operational 50 50

Recovery 50-100 50-100

Total 725 1300

Comparatively, in the biomass mixture it is possible to find a variety of already

functionalized components, considerably reducing the heat necessary to introduce the

functionality. Additionally, these molecules have chemical compositions and structures similar

to the desired products, decreasing the number of conversion steps required. Hence, a well

Introduction

3

designed process using biomass as a raw material should be able to produce bulk

functionalized chemicals at low costs, without all the ecological complications associated with

the use of fossil resources.

A perfect example is the use of amino acids as raw materials in the production

nitrogen-containing bulk chemicals. This concept is one of the foundations of the N-ergy

project, in which this study is incorporated.

1.1.3. The N-ergy Project

The N-ergy project is a long term research project with the ultimate aim of developing

an economically feasible process for the combined production of ethanol and nitrogen

containing chemicals, utilizing biomass as the main raw material. Both end products can be

used as a resource (ethanol) or as a replacement (N-chemicals) for nitrogen containing

products that are traditionally of petrochemical origin. In addition, the produced ethanol can

be utilized as a transport fuel.

Initially, a fermentation step should convert agricultural waste products to ethanol and

insoluble cyanophycin granule peptide (CGP). This fermentation is performed by recombinant

species capable of simultaneous production of ethanol and CGP. Since cyanobacteria – the

natural producers of CGP – are not suitable for the bulk production of large quantities of

CGP5, cyanobacteria genes have been heterologously expressed in heterotrophic bacteria

and plants, with high percentages CGP per cell dry mass being obtained6,7

. Therefore, it is

expected that the genes responsible for CGP production can be successfully expressed in

species that are established ethanol producers, such as yeast and filamentous fungi. Within

the N-ergy project, research is being performed in order to obtain recombinant strains of

Saccharomyces cerevisiae and Rhizopus oryzae that are able to accumulate large volumes of

CGP.

The synthesis of CGP by cells is heavily dependent on the presence of certain amino

acids, particularly arginine, in the growing medium8. This can make the selection of a cheap

and reliable nutrient source a complicated task. However, recent studies have acknowledged

protoamylasse as a suitable medium for large-scale cyanophycin production8. Protamylasse,

or potato juice concentrate, is an abundant waste stream originating from the industrial

production of starch from potatoes. Its composition includes soluble peptides, amino acids,

with asparagine and aspartate as the main components, organic acids, carbohydrates, salts,

and minerals. The high arginine, aspartate, and asparagine contents of protamylasse are of

particular interest for the production and accumulation of CGP.

Cyanophycin granule peptide is a nonribosomally synthesized biopolymer, which

consists of equimolar amounts of arginine and aspartic acid arranged as a polyaspartate

backbone, with arginine residues linked to the β-carboxyl group of each aspartate by its α-

amino group9. In nature this polymer is synthesized by most cyanobacteria, as previously

Introduction

4

stated, and is accumulated as granules in the cells cytoplasm. Although originally thought to

be insoluble in water at neutral pH, later studies detected the formation of a soluble form of

CGP with identical chemical composition10

. Naturally, the insoluble form should allow a much

simpler separation of the polymer from the fermentation broth. The reasoning behind the

different behavior of the two CGP forms is not clear at this point.

After extraction, cyanophycin should be converted to nitrogen-functionalized

chemicals. In a first step, the polymer is completely hydrolyzed to its monomers: aspartic acid

and arginine. After separation, both amino acids should undergo further transformations until

the desired N-functionalized chemicals are obtained. Figure 1.2 illustrates the different steps

in the conversion of biomass to the target products.

NH

NH2

NH

OH

O

NH

O

NH

O

* *

n

OH

O

OH

NH2

O

OH

O

OH NH2

NH

NH

NH2

NH2

O

NH2

OH

NH2

NH2

O

NH2

NH2

BIOMASS

Acrylamide

Aminopropanol

Urea

1,4-Butanediamine

Aspartic Acid

Arginine

Cyanophycin

Ethanol

Figure 1.2. Conversion of biomass to N-functionalized chemicals and ethanol – the N-ergy project.

1.1.4. Arginine Conversion

The focus of this study is on the arginine route. In order to be converted to

1,4–butanediamine (a building block for nylon-4,6), arginine has to undergo two main

transformations: the hydrolysis of arginine to ornithine and urea; and the decarboxylation of

ornithine to 1,4–butanediamine and carbon dioxide (figure 1.3). Both steps can be catalyzed

chemically or enzymatically. The two catalysis methods applied to the first step are explored

and compared during this study.

Introduction

5

O

OH NH2

NH

NH

NH2

O

OH NH2

NH2

NH2

NH2

CO2

OH2

ArginineOrnithine

Urea

1,4-Butanediamine

Figure 1.3. Conversion of arginine to 1,4-butanediamine through hydrolysis to ornithine and

decarboxylation.

Although this work is contained in the N-ergy project, the methods utilized to convert

L-arginine to L-ornithine and 1,4-butanediamine are not limited to the treatment of

cyanophycin and can be applied to any L-arginine source. In fact, many agriculture based

industries, including the production of biofuels, generate waste streams with high protein

content. A good example is the protein rich (50%) soybean meal obtained by grinding the

flakes that remain after extraction of most of the oil by solvent or mechanical process. The

protein can be hydrolyzed to its amino acids which, after separation, can be individually

treated.

1.1.5. Product Applications

Putrescine (commercial name of 1,4-butanediamine) is the product obtained from

arginine with the highest market value. However, secondary products like urea and ammonia

(mainly originating from the spontaneous hydrolysis of urea at high temperatures) can also

provide a significant financial return.

Currently 1,4-butanediamine is produced using chemicals of petrochemical origin:

propylene, ammonia and hydrogen cyanide. This process and its comparison with the

proposed bio-based approach are illustrated in figure 1.4. As can be observed, the suggested

process requires less conversion steps, lower working temperatures and is environmentally

friendly. These advantages were previously discussed in section 1.1.2.

O

OH NH2

NH

NH

NH2

NH2

NH2

CO2NH3

O2

CH4

N

CH N

NH3

Arginine(Biomass)

1,4-Butanediamine

- Urea

-

+

+

+

1.5+

Petrochemical Products

Figure 1.4. Comparison of the petrochemical and bio-based approaches for 1,4-butanediamine bulk

production (adapted from Sanders et al.4).

Introduction

6

Putrescine has the potential to be used as an intermediate in a large array of

industries, including the pharmaceutical industry, the agrochemical industry and the textile

industry, among others. However, at the moment its only relevant application is its use as a

co-monomer, along with adipic acid, in the production of nylon-4,6 (figure 1.5),

commercialized by DSM under the trade name of Stanyl®.

NH2

NH2 HOOCCOOH

NH

NH

CO CO **n

-H2O

1,4-Butanediamine

+

Adipic acid Nylon-4,6

Figure 1.5. Synthesis of nylon-4,6 from 1,4-butanediamine and adipic acid.

Stanyl® is a high performance, high temperature polyamide characterized for its

strong mechanical properties at high temperatures, excellent resistance to wear, low friction,

easy processing and exceptional design freedom. This properties make this polymer an

excellent alternative to the popular nylon-6,6 in processes where high temperatures are

utilized (table 1.2). Key applications for this material include the substitution of metal

components in the automotive industry and Electric & Electronics11.

Table 1.2. Thermal properties of Nylon-6,6 and Stanyl® 11

.

Properties Nylon-6,6 Stanyl®

Melting point (°C) 265 295

Density (kg/m3) 1140 1180

Crystallinity rate (sec-1

):

-at 200°C

-at 230°C

6

0.7

>15

10

Glass transition temperature 65 78

As previously stated, urea and ammonia produced in considerable amounts can

provide a significant source of income. Although some minor applications (like the bulk

chemicals industry) are also relevant, both chemicals are mainly utilized as fertilizers. Urea

and ammonia are an excellent nitrogen source for plant growth that offer several advantages:

are relatively cheap products with inexpensive handling, storage and transportation costs; can

be applied to soil as a solid, solution or even spray to certain crops; involve no fire hazard and

little explosion hazard. The prices of these products are rapidly increasing at the moment,

with urea and ammonia prices close to reaching $800/tonne and $600/tonne respectively12

.

Introduction

7

1.2. Chemical Catalysis

1.2.1. Chemically Catalyzed Hydrolysis of Arginine

The hydrolysis of arginine to ornithine is catalyzed in the presence of strongly acid or

alkaline conditions13,14

.

The acid hydrolysis is a considerably slow process even at high temperatures –

Murray et al.13

obtained a conversion of 98% of L-arginine to L-ornithine after 120 hours of

heating at 176°C in the presence of 6 N hydrochloridic acid. The acid catalyzed reaction

(figure 1.6) leads solely to the formation of L-ornithine, no undesirable by-products are

produced.

O

OH NH2

NH

NH

NH2

O

OH NH2

NH2

H+

NH2

NH2

O

OH2

ArginineOrnithine

+

Urea

+

Figure 1.6. Acid catalyzed hydrolysis of arginine.

Alkaline catalyzed hydrolysis (figure 1.7) can be performed faster and at lower

temperatures, with the disadvantage of the formation of by-products. For each 5.5

equivalents of L-arginine that is converted to L-ornithine and urea, 1 equivalent is converted

to citrulline and ammonia. The citrulline is further hydrolyzed to ornithine but at a slow rate.

Other by-product comes from the reversible conversion of ornithine to its lactam (3-amino-

piperid-2-one) at high temperatures under strong alkali concentration. Despite the existence

of these side-reactions high yields of L-ornithine have been obtained. The heating at 110°C

for 24 hours with the pH adjusted to 12 with ammonia13

resulted in a yield of 90% L-ornithine

with 8% 3-amino-piperid-2-one and 1% citrulline. It should be noted that these results were

obtained through an experimental procedure that is far from being reproducible in an

industrial setting: the reactions were carried in evacuated, sealed 1 ml tubes and heated with

refluxing toluene (as a way of controlling the heating temperature – 110°C – which is

toluene’s boiling point) .

Introduction

8

OH-

O

OH NH2

NH2

O

OH NH2

NH

NH

NH2

OH2

O

OH NH2

NH

O

NH2

-NH3

NH

O

NH2

Ornithine

-Urea

Citrulline

Arginine

+

3-Amino-piperid-2-one

Figure 1.7. Alkali catalyzed hydrolysis of arginine

Other possible by-products suggested in literature are proline15

and

diketopiperazine16

(the product of the reaction between two ornithine molecules).

In both cases, acid and alkaline catalysis, the urea formed is immediately hydrolyzed

to ammonia and cyanate due to the high reaction temperature14

. The cyanate is further

hydrolyzed to ammonia and carbon dioxide (figure 1.8)

NH2

CNH

2

O

OC

NNH4

++

(a)

OH3

+NH3 CO2O

CN + +

(b)

Figure 1.8. Thermohydrolysis of urea: (a), reversible conversion of urea to cyanate and ammonia; (b),

hydrolysis of cynate to ammonia and carbon dioxide.

Other than traditional acid and basic catalysis, other techniques have potential to

produce interesting results and are explored in this study. These include the use of metal

salts and heterogeneous (solid) catalysis, with the second one being specially important if this

reaction is integrated in an industrial process. The background to this last method will be

presented in the next section.

Introduction

9

1.2.2. Heterogeneous Catalysis – the use of Zeolites

Zeolites comprise a group of hydrated aluminosilicate minerals and have a micro-

porous structure. The zeolites are framework silicates consisting of interlocking tetrahedrons

of SiO4 and AlO4. In order to be called a zeolite, a mineral should have a (Si +Al)/O ratio of

1/2. The alumino-silicate structure is negatively charged and attracts the positive cations that

reside within. More than 150 zeolite types have been synthesized and 48 naturally occurring

zeolites are known, natural zeolites are formed when volcanic rocks and ash layers react with

alkaline groundwater.

Since synthetic zeolites were first synthesized in 1949 at Union Carbide

Corporation17

, they have proven to be a versatile material with applications in several

industrial processes. These days, major uses are as detergent builders, as adsorbents, and

as catalysts. Although catalytic application of zeolites represents only 12.5% of the total

tonnage utilization, it is 55% of the total market value for synthetic zeolites. Clearly, catalysis

is the major economic driver in the search for new zeolitic materials.

The main use of zeolites as catalysts has always been in the petroleum industry.

Previously, acid catalysts in the fuel industry included silica-alumina gel, supported

phosphoric acid and chlorine treated platinum or alumina17

, but since the late 50’s

strong-acid zeolites have been successfully utilized as catalysts in the cracking of petroleum

for gasoline production. In the last 30–40 years, between 30% and 50% of all motor fuels

(gasoline, jet, and diesel) have been produced world wide with Y zeolite catalysts.

Due to their important role in petroleum refinery, heterogeneous acidic catalysts have

attracted much more attention than heterogeneous basic catalysis. The catalytic capabilities

of basic catalysts were first reported in the early 70’s. Yashima et al.18

reported that side

chain alkylation of toluene was catalyzed by alkali ion-exchanged X and Y type zeolites. In

basic zeolites and other basic heterogeneous catalysts the basic sites are believed to be the

surface oxygen atoms19

. Oxygen atoms existing on any materials may act as basic sites

because the O atoms are able to interact attractively with a proton.

The acid-base properties of these materials can be controlled by selecting the types

of ion-exchanged cations and by the Si/Al ratio of the zeolite framework. Wide variation of

acid-base properties can be achieved by ion-exchange and ion-addition, while relatively small

changes in acid-base properties are yielded by changing the Si/Al ratio. When preparing basic

zeolites, two approaches are possible. One approach is to ion-exchange with alkali metal

ions, and the other is to impregnate the zeolite pores with fine particles that can act as bases

themselves. The former produces relatively weak basic sites, while the latter results in the

strong basic sites. In alkali ion-exchanged zeolites, the type of ions used affects the basic

strength of the resulting zeolites. Effects of the alkali ions on basic strength are in the

following order: Cs+ > Rb

+ > K

+ > Na

+ > Li

+.

In the present study, a Y zeolite ion-exchanged with sodium ions (CBV 100 –

SiO2/Al2O3 ratio of 5:1) is tested as a possible catalyst for the arginine hydrolysis reaction.

Introduction

10

The research on the use of NaY zeolite as a catalyst has been focused on the treatment of

toxic compounds. This includes the catalysis of the conversion of nitriles to primary amides20

and the reduction of nitric oxide21

. Other studies use this material together with other catalysts

such as metal ions (platinum22

, cobalt22

and gold23

) and metallocenes24

.

Concerning the catalysis of the arginine hydrolysis reaction, no work with zeolites can

be found in literature. Nonetheless, Ikeda et al.25

reported mixed results with another

heterogeneous catalyst. In this study montmorillonite was utilized, this material can adsorb

molecules into its interlamellar layers, in which the regular spacing of clay sheets plays an

important role as a shape-selective catalyst. L-arginine was successfully adsorbed into the

interlamellar layer and the hydrolysis reaction was detected using the pressure-jump

relaxation method with electric conductivity detection. However, it was established that the

release of ornithine from the interlamellar layer was very slow step, making this catalyst not

suitable for industrial applications.

1.3. Enzymatic Catalysis

1.3.1. Enzymatic Hydrolysis of L-Arginine – Arginase

The hydrolysis of L-arginine to L-ornithine and urea is catalyzed by L-arginase (L-

arginine amidinohydrolase, EC 3.5.3.1). This is accomplished by the cleaving of the

guanidinium group from arginine which yields urea, a small nitrogen rich molecule. Thus,

arginase plays a fundamental role in the nitrogen metabolism. It is widely spread through the

evolutionary spectrum and can be find in significantly distinct organisms such as bacteria,

yeast, plants and animals.

In this study is utilized bacterial arginase from Bacillus subtilis. The information found

in literature concerning this organism’s arginase is scarce. Nevertheless, the structure of

arginase is conserved across bacterial species and even across eukaryotic organisms. All

arginases are multimeric metallo-enzymes comprised of identical or near identical sub-units

(although it has been shown for human arginase that the monomer is active26

). Each

monomer has a binuclear manganese spin-coupled cluster located in the active site that is

profoundly involved in the reaction mechanism. Both Bacilus caldovelox arginase and rat liver

arginase have been subjected to extensive structural studies27,28

.

Bewley et al. report a trimeric or hexameric quaternary structure, depending on the

pH of the medium, for Bacilus caldovelox arginase27

. The trimer is comprised of three

identical subunits that associate to form a flat disc-like structure. The monomers interact

through a total of 12 hydrogen bounds between each interface. At pH values above 7.0 a

hexamer is formed when one trimer is rotated 20° with respect to the other about a common

threefold axis.

Introduction

11

The fold of Rat liver arginase is similar to that of B. caldovelox, with 248 structurally

equivalent Cα atoms (83% of the sequence27

). However, the trimer monomer-monomer

interaction is mediated by an additional “S”-shaped oligomerization motif (14 extra residues)

at the carboxy terminus, the conformation of this segment is stabilized by numerous inter-

monomer van der Walls interactions, hydrogen bonds and salt links28

(figure 1.9). The

hexameric form is not found in rat liver arginase.

Figure 1.9. Topology diagram of rat liver arginase. Relative location of metal ligands is indicated by

grey circles (adapted from Kanyo et al.)28

.

The geometry of the active site also appears to be similar in rat liver and bacterial

arginases27

. The catalytic mechanism proposed by Kanyo et al.28

for rat liver arginase

involves the formation of hydrogen bonds between the guanidinium group of arginine and

carboxylate side chain from the Glu277 residue located in the active site (figure 1.10). Later

studies not only confirmed the existence of these bonds but also indicate that other residues –

His141 and Thr246 – accept hydrogen bonds from the guanidinium group and help stabilize

the substrate29

. This array of hydrogen bonds orients the guanidinium group for nucleophilic

attack by a hydroxide ion that bridges both manganese ions. Several bridging metal ligands

coordinate the two manganese ions and stabilize the solvent bridge between them, including

Asp128 which also hydrogen bonds with the hydroxide ion.

The nucleophilic attack leads to the formation of metastable tetrahedral intermediate.

The metal ions are essential in transition state stabilization by keeping the metal-bridging

hydroxide in position, with Christiansson et al.29

suggesting that one of the manganese ions

also directly interacts with one of the NH2 groups from the guanidinium. Following a proton

transfer to the leaving amino group mediated by Asp128, the collapse of the tetrahedral

intermediate yields the products L-ornithine and urea. The subsequent addition of a water

molecule to the binuclear manganese cluster facilitates urea departure, which may trigger the

ionization of the metal-bridging water molecule to regenerate the nucleophilic metal-bridging

hydroxide ion.

Introduction

12

Figure 1.10. Proposed mechanism of rat liver arginase-catalysed arginine hydrolysis by metal-activated

solvent28

. The α-amino and α –carboxylate groups are omitted for clarity.

Arginase´s catalytic action is highly specific for the hydrolysis of its natural substrate,

L-arginine. Its enantiomer D-arginase is not a substrate and the use of similarly structured

molecules and derivatives as alternative substrates severely attenuates the catalytic activity

of rat liver arginase30

. Tested molecules include: L-canavanine, L-homoarginine, L-argininic

acid, agmatine and L-argininamide (best alternative substrate tested with a 12-fold increase in

KM and a 14-fold decrease in Vmax).

The best arginase inhibitors are those bearing N-hydroxyguanidinium or boronic acid

“warheads” that bridge the binuclear manganese cluster, including Nω-hydroxy-L-argininine

and N-hydroxy-L-lysine. The simplest arginase inhibitor is the fluoride ion, an uncompetitive

inhibitor with a Ki of 1.3 mM. The crystal structure of the binding of this inhibitor to the

enzyme-substrate complex has been determined31

and shows an unusual mode of inhibition:

the metal-binding hydroxide ion is displaced by a fluoride ion and another fluoride ion is

added to the vacant coordination site of one of the manganese ions (Mn2+

A). The metal

bound fluoride ions are stabilized by short hydrogen bonds with the guanidinium group of the

substrate, in a typical uncompetitive inhibition substrate binding is required to stabilize the

inhibitor.

The properties of Bacilus subtillis arginase were the subject of only one published

study by Nakamura et al.32

. The enzyme purified during this work has a specific activity of 858

U/mg-protein and a native molecular weight of 115.000 Da. It is also reported an optimum pH

at pH=10, a relatively high KM for L-arginine of 13.5 mM and an increased in thermal stability

through the addition of Mn2+

ions.

In literature various methods for arginine (usually originating from mammal liver)

immobilization have been studied with mixed results (table 1.3). Several of these studies were

conducted with the intent of using arginase has an arginine detection tool.

Introduction

13

Table 1.3. Techniques for arginase immobilization described in literature. The different techniques are

listed in chronological order, from the oldest work to the most recent.

Immobilization Method Particular Aspects

Covalent binding to controlled pore glass bead

derivative33

Arginase and urease were simultaneously

immobilized in the beads and assembled on an

ammonia sensing electrode for arginine detection.

Entrapment in a highly porous polymer matrix34

The enzyme is immobilized in the matrix trough

radiation induced polymerization of acrylic

monomers.

After immobilization on bead shaped matrix only

30-40% of the initial enzymatic activity was

retained.

Covalent binding on various types of carbodiimide

activated carboxyl-functionalized polyacrylamide

beads35

Optimum pH for the catalytic activity shifted in the

acid direction.

Optimum temperature for the catalytic activity of

the immobilized arginase much higher than that

for the soluble enzyme.

Kmapp

of the immobilized arginase for L-arginine

was an order of magnitude higher than that of the

soluble enzyme.

Entrapment in membrane reactor system36

Enzyme recovered with a UF membrane after a

batch for re-use in the next batch.

Conversion after 24h decreased from 53% to zero

after 5 runs.

Covalent binding on epoxy-functionalized resin37

The resin is contained inside a continuous-flow

reactor for arginine detection.

The enzyme reactor is stable for more than 6

months and retains about 80% of its initial activity

after 800 assays.

Entrapment in gelatin gel38

Arginase immobilized together with urease on the

surface of a pH electrode for arginine detection.

The gelatin matrix has no effect on the system.

Covalent binding on N,N’-disuccinimidyl suberate

activated aminopropyl silica39

Enzyme immobilized on a chromatography

support to study the binding of nor-NOAH to

arginase.

The arginase column is stable during a long

period of time.

The next sections encompass a brief overview of the use of biocatalysts in industrial

processes and a section on the importance of biocatalysts immobilization, with focus on the

immobilization methods utilized in this study.

Introduction

14

1.3.2. Industrial Biocatalysis

For millennia enzymes have been utilized in the production of several food products.

However, it was only decades ago that biocatalysis started to be regarded as a valuable tool

by the chemical industry. Examples of initial applications include the use of acylases,

hydantoinases, and aminopeptidases in the production of optically pure amino acids, and the

use of nitrile hydratase in the enzymatic production of the bulk chemical acrylamide from

acrylonitrile40

.

Since then the industrial use of biocatalysis has expanded, with enzymes being

utilized as catalysts in the industrial synthesis of bulk chemicals, pharmaceutical and

agrochemical intermediates, active pharmaceuticals, and food ingredients40

. Most these

commercial enzymatic processes share several attributes: high product concentrations and

productivities, no undisirable by-products and the use of enzymes that do not require

expensive co-factores41

. Today, the industrial community sees biocatalysis as a highly

promising area of research, especially for the development of sustainable technologies for the

production of chemicals40

. This sustainable, environmentlaly friendly, production of bulk

chemicals (green chemistry) is the aim of the N-ergy project, and of this study in particular.

However, the number and diversity of the applications of biocatalysis are still modest

when compared with traditional catalysis methods. This happens, in part, because of the

limitations inherent to a process catalysed by enzymes: substrate scope, limited enzyme

availability and operational stability40

. Recent breakthroughs in certain areas should help

overcome these limitations. Advances in genomics, directed evolution and bioinformatics

allow not only the discovery of new enzymes but also the optimization of existing ones. In

addition, the development of a feasible biocatalytic process usually requires a major financial

investment. Depending on the type of biocatalyst to be used, specific reactor and hardware

configurations are needed. Biocatalytic processes are typically highly heterogeneous and

need specific designs of the catalyst–hardware interface to allow efficient immobilization and

re-utilization41

.

The lack of operational stability of certain enzymes when utilized in industrial scale

processes is one of the constraints that has received more attention in recent years. Various

techniques have been developed to improve stability so that enzymes can be used with

organic solvents, high temperatures and extreme pHs40

. Among these techniques are protein

engineering, process modification and, as discussed in the next section, immobilization of the

biocatalysts.

Introduction

15

1.3.3. Biocatalyst Immobilization

Immobilization of biocatalysts consists in its confinement to a defined area

(bioreactor), ensuring the maintenance of the catalytic activity and allowing its repeated or

continuous use. Immobilization methods have been applied to a wide array of biocatalysts,

ranging from pure enzymatic extracts to whole microbial cells or even animal and vegetal

tissues. When applied to enzymatic extracts on an industrial setting, immobilization offers

several advantages and some restrictions, the most relevant of which are listed in table 1.4.

Table 1.4. Advantages and limitations associated to the use of immobilized enzymes (adapted from

Cabral et al.42

)

Advantages Particular Aspects

Retention of the catalyst inside the reactor Allows reutilization and continuous processes

Possibility of operating on high dilution rates

without the risk of wash-out

High catalyst concentrations Allows higher volumetric production rates

Faster bioconvertion, important when secondary

reactions are a issue

Controlled microenvironment of the catalyst Allows manipulation of enzymatic activity and

specificity

Improves enzyme stability

Protects the enzyme against shear stress

Easy separation between catalyst and product Minimizes product contamination

Precise control of bioconversion time

Limitations Particular Aspects

Loss of catalytic activity May occur during the immobilization process,

during the bioconversion or due to the physical

properties of the immobilization matrix.

Empiric process Specific optimization is needed for each particular

application

Complex control and modeling

The different types of immobilization of biocatalysts have been the subject of several

classification systems. One of them, adapted from Cabral et al.42

, is presented in figure 1.11.

Introduction

16

Figure 1.11. Methods of immobilization of biocatalysts.

Due to its multimeric structure, arginase is an excellent candidate for immobilization

by covalent bonds. The binding of this enzyme to a support can play an important an

important role in avoiding dissociation of the enzyme by keeping the sub-units together. This

kind of immobilization not only should enhance enzyme stability but also is essential when

utilizing arginase in an industrial setting.

Most protocols for protein immobilization described in literature are difficult to

reproduce on an industrial scale where long support handling may be necessary and some

dangerous substances cannot be utilized, problems that are not considered on a laboratory

scale. Comparatively to other immobilization methods, covalent binding supports, and the

epoxy-activated supports utilized in this study in particular, are almost ideal for performing

easy industrial immobilization of enzymes. Epoxy-activated supports are very stable during

storage and also when suspended in neutral aqueous media. Hence, they can be easily

handled before and during immobilization procedures. In addition, these supports are able to

directly form very stable covalent linkages with different protein groups (amino, thiol, and

phenolic ones) under very mild experimental conditions43

.

The immobilization of enzymes in epoxy-activated supports usually follows a two step

mechanism: first a rapid mild physical adsorption between the protein and the support is

produced, and secondly the covalent reaction between adsorbed protein and epoxy groups

occurs (figure 1.12). In order to adsorb proteins during incubation at high ionic strenghts,

commercial epoxy supports are fairly hydrophobic, in hydrophilic supports (e. g., agarose) this

preliminary physical hydrophobic adsorption is not possible. The remaining epoxy groups may

be easily blocked after the protein immobilization, to stop any kind of undesired covalent

support-protein reaction.

Biocatalyst Immobilization

Insoluble Soluble

Reticulation Bonding to a Support With Modification of the Microenvironment

Without Modification of the Microenvironment

Adsorption Ionic Bond Covalent Bond

Gel Micro-encapsulation

Ultrafiltration Membranes

Microcapsules Inverted Micelles

Introduction

17

Figure 1.12. Mechanism of immobilization of proteins on epoxy-activated Supports. The covalent

reaction between soluble enzyme and epoxy support is extremely slow, but the previous adsorption to

the support allows a faster covalent reaction (adapted from Mateo et al.43

).

Even though other immobilization techniques inside porous supports can increase the

enzyme operational stability by preventing any intermolecular process (proteolysis,

aggregation) and by preserving the enzyme from interactions with external interfaces (air,

oxygen, immiscible organic solvents, etc.), these techniques do not necessarily increase the

conformational stability of the enzyme43

. This kind of stability should be achieved if the

immobilization of each enzyme occurs through several residues. This way, all the residues

involved in immobilization preserve their relative positions and the enzyme is unaffected by

conformational changes promoted by heat, organic solvents, or any other distorting agents43

.

Thus, multipoint covalently immobilized enzymes should become more stable than their

soluble counterparts or than randomly immobilized derivatives. It is important that the reactive

groups that react with the enzyme are bound to the surface of the support by short spacer

arms (two to three carbon atoms), allowing the reaction to occur only with the external

residues of the enzyme but not residues located in internal pockets. The bond to a rigid

support through short spacer arms is vital when dealing with multimeric enzymes. Fernadez-

Lafuente et al.44

suggests the use of short spacer arms supports together with cross-linking

agents to achieve the stabilization of the quaternary structure of multimeric enzymes with no

side modifications.

In this study three epoxy supports are tested for arginase immobilization:

Sepabeads® EC-EP, Sepabeads

® EC-HFA and Eupergit C 250 L. All supports are

microporous, epoxy-activated, acrylic polymer matrix spherical beads.

Sepabeads® EC-EP is a highly activated support functionalized with short chain

epoxy groups while the Sepabeads® EC-HFA supports are functionalized with epoxy groups

on a longer, more complex, spacer (figure 1.13). Both are very rigid supports that may be

used in stirred tanks or bed reactors. These supports have low swelling tendency in high

molar solutions and in common solvents. Also, their internal geometry offers large internal

Introduction

18

plain surfaces where the enzyme may undergo intense interactions with the support43

. The

standard grade beads have a diameter of 150-300 µm with an average pore diameter of 30-

40 nm and a specific gravity of 1.13 g/ml45

. The epoxy group density on the beads is around

100 µmol/(g of wet support)43

.

Figure 1.13. Functional groups of Sepabeads® EC-EP and Sepabeads

® EC-HFA supports

45.

Eupergit C 250 L is activated similarly to Sepabeads® EC-EP, having on its surface a

dense monolayer of reactive and stable epoxy groups (200 µmol/(g of dry support) according

to the supplier). The bead diameter is 100-250 nm with an average pore diameter of 100

nm46

. This kind of supports is amongst of the most extensively studied due to their capability

of immobilizing enzymes quickly and easily, both at laboratory and industrial scale.

Examples of the industrial application of epoxy-activated supports are the use of

Sepabeads supports on the production of 6-amino penicillanic acid and on the conversion of

cephalosporin C into alpha-keto-adipoyl-7-amino-cephalosporanic acid47

. The only case of

arginase immobilization on an epoxy-activated support found in literature is the already

referred work of Alonso et al.37

, where arginase was successfully immobilized on a

commercial resin for arginine detection.

1.4. Aim of this Study

The main goal of this project is to establish an industrial viable method of converting

L-arginase to L-ornithine, contributing to the aim of the overall N-ergy project of coverting

biomass to ethanol and bulk N-fuctionalized chemicals. With this objective in mind two

approaches are studied: the chemically catalyzed hydrolysis of L-arginine and the hydrolysis

of L-arginine catalyzed by the enzyme arginase.

Concerning the chemical catalysis, the experimental conditions studied are closer to

an industrial setting than previous works found in literature13,14,15,24

. The effects of pH and

temperature in the yield of ornithine are studied, as well as the influence of metal ions on the

reaction mixture. NaY zeolite, a heterogeneous catalyst potentially suitable for industrial

application is also tested. It should be noted that all research on the chemically catalysis of

the conversion of arginine is focused on the basic hydrolysis. The acid catalyzed reaction,

regardless of being cleaner, is considerably slower and requires very high temperatures,

being inadequate for a large scale process.

Introduction

19

Regarding the biocatalysis, the hydrolysis of L-arginine to L-ornithine and urea

catalyzed by Bacilus subtillis arginase is studied. In this case, the main objective is to

investigate the effect of immobilization in covalent-binding supports on the performance of the

enzyme. Different commercially available epoxy-activated supports suitable for industrial

use48,46

are tested for optimum stability/activity. After selecting the best performing enzyme

preparation, the chemical and enzymatic hydrolysis of arginine are compared.

Finally, even though the main goal is the production of ornithine, a secondary

objective of this project is to further contribute to the developing N-ergy project. This includes,

if possible, the collection of data on the conversion of ornithine to 1,4-butanediamine and the

monitoring of the formation of economically important secondary products like ammonia and

urea.

Chemical Hydrolysis of L-Arginine

21

2. Chemical Hydrolysis of L-Arginine

2.1. Materials and Methods

2.1.1. Reagents

Table 2.1. Reagents utilized during the course of the chemical hydrolysis of arginine experiments.

Chemical Supplier Purity (min)

1,4-Butanediamine Sigma-Aldrich 99%

2,2,3,3-d(4)-3-(trimethylsilyl)propionic

acid sodium salt (TSP) Alfa Aesar 98%

3-Aminopiperid-2-one Activate Scientifics 95%

Acetic acid Riedel-de-Haen 99.8%

Acetone Merck 99.8%

Acetonitrile Merck 99.9%

Al(NO3)3 Merck 98.5%

Borax Riedel-de-Haen 99.5%

Boric acid Merck 99.5%

CuSO4 Sigma-Aldrich 99%

Dabsyl chloride Sigma-Aldrich 99%

Dimethylformamide (DMF) Lab-Scan 99.8%

Deuterium oxide Sigma-Aldrich 99

Ethanol Merck 99.9%

L-Aspartic acid Merck 99%

L-Arginine Sigma-Aldrich 99%

L-Citrulline Sigma-Aldrich 99%

L-Ornithine Sigma-Aldrich 99%

MnSO4 Sigma-Aldrich 99%

NaY Zeolite Zeolyst International -

Sodium Acetate Merck 99%

Sulfuric acid-D2 Sigma-Aldrich 99%

ZnCl2 Boom 98%


22

2.1.2. Solutions

Table 2.2. Solutions prepared during the course of the chemical hydrolysis of L-arginine experiments.

Solution Composition

Dabsyl chloride solution 24 mg of Dabsyl chloride in 10 ml of acetone

Derivatization Buffer 0.1 M borate buffer, pH=9.0

(addition of a 6.2 g/l solution of boric acid in Milli-Q

water to a 38.15 g/l solution of borax in Milli-Q

water until pH = 9.0 is obtained)

Dilution Buffer 50% Acetonitrile

25% Ethanol

25% Eluent A

Eluent A 96% 20mM sodium acetate in Milli-Q water

4% DMF

pH adjusted to 6.4 with concentrated acetic acid

Eluent B 80% Acetonitrile

20% Milli-Q water

2.1.3. Equipment

High Performance Liquid Chromatography

Reverse-phase HPLC analyses were performed on a Waters™ System consisting of:

Waters™ 600s Controller; Waters™ In-Line Degasser; Waters™ 616 Pump; Waters™

717plus Autosampler; Waters™ 484 Tunable Absorbance Detector; Nova-Pak® C18 column

60 Ǻ 3.9 x 150 mm with a Nova-Pak® C18 4 µm Guard-Pak™ pre-column insert.

Nuclear Magnetic Resonance

1H-NMR and

13C-NMR analyses were performed on a Bruker AVANCE III 400 MHz

NMR spectrometer.

Multiple Reactor System

All reactions were performed on a Parr® Series 500 Multiple Reactor System

equipped with a 4871 Process Controller. This system is fully programmable and allows the

simultaneous heating of up to six 75 ml reactors with internal stirring. The individual reactors

are made of alloy C-276 and tolerate operational temperatures up to 300°C and operational

pressures up to 3000 psi. Each reactor includes: a thermocouple mounted inside the reactor

and protected by stainless steel sheaths; a dip tube with sample valve; an optional glass liner

to protect the metal walls from corrosive substances (using the glass liners reduces the

reactor volume to about 300 ml).


23

Figure 2.1. Parr® Series 500 Multiple Reactor System with 4871 Process Controller.

Thermomixer

An Eppendorf® Thermomixer Comfort was utilized. This thermomixer is equipped with

a rack that allows the simultaneous heating of up to 24 1.5 ml eppendorf Safe-Lock tubes. It is

fully programmable, capable of heating or cooling samples from 1°C to 99°C and of agitating

from 300 rpm to 1500 rpm (also has a no mixing mode).

2.1.4. Analytical Techniques

Dabsyl Chloride Derivatization

Previously to HPLC analysis, a Dabsyl chloride derivatization procedure adapted from

Kause et al.49

was applied to each sample. This pre-column derivatization method allows the

efficient separation and detection in the visible region of amino acids as well as primary and

secondary amines, including putrescine.

Samples were diluted five times with a 1.25 mM L-aspartic acid (internal standard) in

Milli-Q water solution. The total amino acid concentration of the diluted samples should be

around 5 mM, with a concentration of internal standard of 1 mM.

Aliquots of 20 µl of the diluted samples were further diluted with 180 µl of the

derivatization buffer and, after mixing on a vortex mixer, 100 µl of dabsyl chloride solution was

added to the samples, immediately followed by thorough mixing. Samples were incubated at

70°C for 15 minutes in a thermomixer. The reaction was stopped by placing the samples on

ice for 5 minutes, followed by the addition of 200 µl of dilution buffer for a final volume of 500

µl per sample. Finally, the samples were centrifuged for 5 minutes at 14 000 rpm and 200 µl

of the supernatant were used for HPLC analysis.

HPLC Analysis

After derivatization, the samples were analyzed on the previously described reverse-

phase HPLC Waters™ system. 10 µl per sample were injected and the column was eluted at

50°C with a flow rate of 1 ml/min. The gradient of the eluents utilized is described in table 2.3,


24

the total time of analysis for each sample was of 80 minutes. UV detection was carried at the

wavelength of 436 nm.

Table 2.3. Gradient of eluents applied during HPLC analysis.

Time % Eluent A % Eluent B Curvea

0 92 8 6

2 92 8 5

7 80 20 7

35 65 35 6

45 50 50 6

66 0 100 6

71 0 100 6

75 92 8 6

80 92 8 6

a for the slope of the different curves see Appendix A.

NMR Analysis

The samples for NMR analysis were collected from experimental settings where the

reactions were carried out using D2O as solvent. NMR tubes were filled with 1 ml of each

sample. To each tube was added 15 µl of 10% D2SO4 in D2O and small amounts of TSP.

Samples were analyzed for 1H-NMR,

13C-NMR and DEPT spectra.

2.1.5. Hydrothermolysis Experiments

These experiments were conducted with the objective of studying the influence of pH

and temperature in the alkaline hydrolysis of L-arginine and identifying the setting that leads

to maximum L-ornithine yield.

A solution of 25 mM L-arginine in Milli-Q water was prepared. Portions of this solution

were adjusted to pH 11 and 12 with a concentrated (0.1 M) sodium hydroxide solution.

Volumes of 60 ml of the three solutions were heated in Parr pressurized stirred reactors at

110°C, 125°C and 150°C for a total of 9 distinct experimental conditions. The reactors were

heated accordingly to a pre-programmed temperature gradient. The clock was started after 30

minutes of heating, corresponding to the time needed for the reactors’ internal temperature to

rise from room temperature to the desired temperature.

3 ml samples were taken from each reactor through a dip tube at 0, 1, 2, 18 and 20

hours. The reaction was stopped by immediately placing the samples on ice. The experiments

were conducted in duplicate.


25

Before each sample was taken, 4 ml aliquots of the reaction mixture were collected

through the dip tube. This portion is not utilized for analysis and corresponds to the volume of

liquid stored inside the tube which is not at the temperature measured inside the reactor.

The samples were stored in the fridge (5˚C) until dabsyl chloride derivatization and

HPLC analysis.

For NMR analysis 60 ml of 25 mM L-arginine in D2O were heated in the same

reactors at 125°C for 15 and 30 hours. The dip tube was not used, instead the reaction

mixture was collected after the reactor cooled down to room temperature.

2.1.6. Metal Salt Catalysis Experiments

This experiment was conducted with the objective of verifying if the presence of metal

ions has any catalytic effect in the arginine hydrolysis reaction.

Different metal salts were added to solutions of 25 mM L-arginine in Milli-Q in

equimolar concentrations. The following salts were added: Aluminium Nitrate (Al(NO3)3·H2O)

9.38 g/l; Copper(II) Sulfate (CuSO4) 3.99 g/l; Manganese(II) Sulfate (MnSO4·H2O) 4.25 g/l:

Zinc Chloride (ZnCl2) 3.41 g/l. Volumes of 30 ml of the four solutions and a blank solution with

no salts added were heated in Parr pressurized stirred reactors (utilizing a glass liner) at

125°C accordingly to a pre-programmed temperature gradient. The clock was started after 30

minutes of heating, corresponding to the time needed for the reactors’ internal temperature to

rise from room temperature to the desired temperature.

3 ml samples were taken from each reactor through a dip tube at 0, 2 and 4 hours.

The reaction was stopped by immediately placing the samples on ice.





HPLC analysis.

2.1.7. Zeolite Catalysis Experiment

This experiment was conducted with the objective of replicating, or improving, the L-

ornithine yield values obtained through traditional alkaline catalysis utilizing a solid catalyst

suitable for an industrial process. Different concentrations of NaY zeolite were tested for this

effect.

Different quantities of NaY zeolite were added to solutions of 25 mM L-Arginine in

Milli-Q for final concentrations of 0.5 gz/lsol, 1 gz/lsol, 2 gz/lsol, 5 gz/lsol. Volumes of 30 ml of the

three suspensions and a blank solution with no zeolite added were heated in Parr pressurized


26

stirred reactors (utilizing a glass liner) at 125°C accordingly to a pre-programmed temperature

gradient. The clock was started after 30 minutes of heating, corresponding to the time needed

for the reactors’ internal temperature to rise from room temperature to the desired

temperature.

3 ml samples were taken from each reactor through a dip tube at 0, 18 and 20 hours.

The reaction was stopped by immediately placing the samples on ice. The experiments were

conducted in duplicate.





HPLC analysis.

2.2. Results and Discussion

2.2.1. Hydrothermolysis Experiments

The alkaline hydrothermolysis of L-arginine was followed during 20 hours at different

temperatures. Previous studies on this reaction under similar pH and temperature conditions

reported high L-ornithine yields. Murray et al.13

, for example, obtained a yield of 90% L-

ornithine at pH=12 and T=110ºC. Yet, the various works found in literature utilize

experimental setups considerably different from the one that was employed in the present

study. Murray et al.13

, Wong et al.16

, and Vallentyne., et al.15

performed similar thermal

degradation experiments in sealed evacuated tubes, with the reaction being stopped at

certain point by cooling the tubes. The experimental setting now employed is closer to an

industrial scale process, the reactions are carried in pressurized reactors and samples are

regularly taken through a dip tube.

Figure 2.2 shows the rate of L-arginine consumption under the diverse experimental

conditions tested.


27

110 C

0

20

40

60

80

100

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

pH=10.6

pH=11

pH=12

125 C

0

20

40

60

80

100

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

pH=10.6

pH=11

pH=12

150 C

0

20

40

60

80

100

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

pH=10.6

pH=11

pH=12

Figure 2.2. Time course of L-arginine consumption under different experimental conditions. The

percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the

reaction mixture at t=0. Error bars calculated using standard deviation of the two duplicate experiments.

The small L-arginine consumption registered at t=0 is due to the 30 minutes of

heating necessary for the reactors to reach the desired work temperature. Accordingly to what

is described by Warner et al.14

, temperature positively influences arginine hydrolysis. The

higher conversion values were detected at 150°C where, after 20 hours of reaction, the high

temperature nullifies the pH effect, with concentrations of remaining arginine reaching 6%.

The influence of pH is also clear, especially at lower temperatures. Although the adjustment

of the initial pH to 11 with sodium hydroxide has almost no effect on the rate of consumption,

the adjustment to pH=12 leads to a decrease of arginine concentration of 25% at 110°C and

125°C after 20h of reaction. This effect was also previously described by Warner et al.

Figure 2.3 shows the rate of L-ornithine formation under the diverse experimental

conditions tested.


28

110 C

0

5

10

15

20

25

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

pH=10.6

pH=11

pH=12

125 C

0

5

10

15

20

25

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

pH=10.6

pH=11

pH=12

150 C

0

5

10

15

20

25

0 5 10 15 20 25

Time (hours)

Rela

tive

Concentr

atio

n (

%)

pH=10.6

pH=11

pH=12

Figure 2.3. Time course of L-ornithine formation under different experimental conditions. The

percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the


Again, the small L-ornithine formation registered at t=0 are due to the initial 30

minutes of heating before the clock was started. The influence of pH in ornithine formation is

concordant with its influence in arginine degradation: in every temperature higher pH leads to

a higher ornithine yield. However, in this case, the effects of temperature are not as linear.

The maximum yield, 15,2%, was obtained at 125°C pH=12.

Comparing arginine consumption with ornithine yield, it becomes clear that secondary

products are being formed. For example, at 125°C pH=12, after 20 hours a consumption of

85,4% of the arginine leads to the formation of only 15,2% of ornithine. The complex HPLC

chromatograms and NMR spectra (Appendix F) reinforce the idea that multiple products are

present in the reaction mixture. Probable side-products are: citrulline, formed directly from

arginine; 3-aminopiperid-2-one, formed from the lactamization of ornithine; putrescine (1,4-

butanediamine), formed from the decarboxylation of ornithine. The decrease of ornithine

concentration between 18h and 20h at 150°C suggests that ornithine is being converted to

another product.


29

Figure 2.4. Example of a typical HPLC chromatogram obtained after heating a L-arginine solution for

20h in the described experimental settings. The identified peaks correspond to: (A) L-aspartic acid

(internal standard); (B) hydrolyzed excess derivatization reagent; (C) L-citrulline; (D) L-arginine; (E) L-

ornithine; (F) ammonia; (G) 3-aminopiperid-2-one; (H) retention time of putrescine elution(none was

detected in the samples). All unidentified peeks correspond to impurities originating either from the

solutions utilized or from the HPLC system.

After identification of the different peaks on the HPLC chromatogram (figure 2.4), the

presence of citrulline and 3-aminopiperid-2-one in the reaction mixture was established. Still,

citrulline appears only in small amounts and the presence of the lactam reasonable amounts

is not enough to explain the substantial disparity between arginine conversion and ornithine

yield. It should be noted that it was not possible to calculate the exact amount of 3-

aminopiperid-2-one in solution for reasons explained in section 2.2.4.

Despite not being detected during HPLC analysis, the hypothesis of putrescine

formation can’t be neglected. Putrescine is a relatively volatile product, and there is a

possibility that it escapes as a gas at the moment a sample is being collected. This

supposition is supported by the presence of foul odor felt during sample collection (putrescine

is known for its strong odor).

However, the comparison of 1H-NMR spectra appears to demonstrate that there is no

putrescine formed during the experiments. The 1H-NMR spectum of putrescine (Appendix E)

shows a distinct peak at 2.6 ppm that clearly is not present on the 1H-NMR spectum for the

reaction mixture (Appendix F). Samples for NMR analysis are collected only after the reactors

cool to room temperature, which guarantees that this product is in liquid state.

Other possible secondary products referred in literature are proline15

and

diketopiperazine16

. The proline detected by Vallentyne et al. is probably citrulline incorrectly

identified, which is an easily understandable mistake considering the less accurate detection

methods available at the time of the study (1968). The formation of diketopiperazine from the


30

reaction between two molecules of ornithine seems unlikely in diluted solutions, which is the

present case. Thus, new and more precise analytical methods should be employed to follow

the thermohydrolysis of arginine reaction in order to clarify which compounds are being

formed and in what amounts.

Also noticeable in figure 2.3 is the big discrepancy (error bars) between the values of

both measurements (duplicates) for each point at high pH values. The experiments were

repeated more than once, also in duplicate, and the big standard deviation persisted. This can

be explained by the complexity of the reactions occurring in the system. At each moment

multiple reactions are happening in the reaction mixture – lactamization, hydrolysis, formation

and degradation of citrulline – and the reactions are dependent from each other and from the

pH of the medium which, in turn, depends from the concentration of the different products (as

seen in figure 2.5). At high initial pH values this equilibrium is even more fragile so a minor

perturbation, like a valve from the dip tube that is opened too much or too little, can greatly

affect the system. On a lab scale the use of buffers would minimize this problem, but the use

of salts is not desirable in an industrial setting.

0

4

8

12

16

0 5 10 15 20 25

Time (hours)

Re

lative

Co

ncen

tration

%

10

10.5

11

11.5

12

pH

[ORN]

pH

Figure 2.5. Evolution of the pH during the course of the reaction (125°C, initial pH of 12) and its

comparison with ornithine concentration. Error bars were omitted for clarity.

During analysis no urea was detected. This is easily explained as at temperatures

above 100°C in alkaline conditions urea is almost immediately hydrolyzed to ammonia and

carbon dioxide14

. Considerable amounts of ammonia were detected through HPLC analysis.

Despite being less valuable than urea, ammonia market prices at the moment (August 200)

are extremely high, reaching $600/tonne12

. Bearing in mind that each tonne of arginine as the

potential to produce 195 kg of ammonia, it could be of financial interest to recuperate this

secondary product.


31

2.2.2. Metal Salt Catalysis Experiments

This experiment was performed based on the idea that the presence of metal ions

could help stabilize the guanidinium group of L-arginine making it a better leaving group. This

would lead to faster arginine conversion and lower working temperatures. The lower

temperatures would also minimize the lactamization of ornithine to 3-aminopiperid-2-one. The

influence of manganese ions was specially anticipated, as this metal has an important role on

the catalytic mechanism of arginase28

.

This premise is based on studies on the successful catalysis of the hydrolysis of

amides using different metal ions including: copper(II)50,51

, nickel(II)51

, cobalt(II)51

and cobalt

(III)52

. Curiously, the work developed Meriwether et al.51

is motivated by the catalytic

mechanism of another metallo-enzyme. In this case, different metals are tested to try to mimic

the catalytic activity of exopeptidases.

Different metal salts were tested at 125°C in equimolar concentrations with arginine:

manganese sulfate, zinc chloride, aluminium nitrate and copper sulfate.

0

0,5

1

1,5

2

2,5

3

3,5

4

0 1 2 3 4 5

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

MnSO4

ZnCl2

Al(NO3)3

no salts added

Figure 2.6. Effect of the presence of different metal salts (equimolar concentrations – 25 mM) on the L-

arginine thermohydrolysis reaction. The percentages are based on the total concentration of amino

acids (L-arginine and L-ornithine) in the reaction mixture at t=0.

Again, the ornithine concentrations registered at t=0 are due to the preliminary

heating (30 minutes) needed for reaching he desired work temperature. As observed in figure

2.6 the presence of metal salts appears to have no significant influence on the arginine

hydrolysis reaction. Copper sulfate interfered in the derivatization reaction (certain amino

acids, including the internal standard aspartic acid weren’t derivatized) and its effect couldn’t

be determined.


32

2.2.3. Zeolite Catalysis Experiment

This experiment was performed with the objective of replicating or improving the

results previously obtained for ornithine yield, utilizing a catalyst suitable for an industrial

process. In literature, no comparable work is found on the arginine hydrolysis reaction with

zeolites or other similar materials. The reaction was followed for 20 hours (figure 2.7) with

addition of NaY zeolite in different concentrations: 0.5 g/l, 1 g/l, 2 g/l, 5 g/l and no zeolite

added.

L-Arginine

0

20

40

60

80

100

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

1g/l

2g/l

0.5 g/l

5 g/l

no zeolite added

L-Ornithine

0

5

10

15

20

25

0 5 10 15 20 25

Time (hours)

Re

lative

Co

nce

ntr

atio

n (

%)

1g/l

2g/l

0.5 g/l

5 g/l

no zeolite added

Figure 2.7. Effect of different NaY zeolite concentrations on the L-arginine thermohydrolysis reaction.

The percentages are based on the total concentration of amino acids (L-arginine and L-ornithine) in the


As can be observed in figure 2.7, the effects of the zeolite in the reaction are limited

even at high concentrations of 5 g/l. Nonetheless, there is a mild catalytic effect, the best

results, 11.2%, are obtained with 5 g/l at 18 hours. It should be noted that concentrations this

high (5 g/l) of zeolite are probably unsuitable for application in an industrial process.

The zeolite utilized also showed to be the impractical, even at laboratory scale, due to

its accumulating on the side walls of the glass liners, being extremely difficult to remove.

2.2.4. Analytical Methods

Dabsyl Chloride derivatization and the HPLC analysis method utilized were

appropriate for the experiments performed. With the copper sulfate exception already

mentioned, derivatization and separation of all relevant components of the reaction mixture

was achieved. Nonetheless, ideally, specific methods should be applied for some

components, such as ammonia. It was not possible to determine the exact concentrations of

3-aminopiperid-2-one. During derivatization, the relatively high temperatures (70°C) led to the


33

partial conversion of this lactam to ornithine13

(figure 2.8), not allowing a viable standard

calibration curve.

Concerning in the NMR analysis, a similar problem is observed in the 3-aminopiperid-

2-one spectra (appendix D). The lactam’s spectra show peaks that are also present in the

ornithine NMR spectra (appendix C). In this case the conversion is catalyzed by the addition

of sulfuric acid13

.

Figure 2.8. Detail from a HPLC chromatogram for a 25 mM 3-aminopiperid-2-one solution. The

identified peaks correspond to: (O) L-ornithine; (A) ammonia; (L) 3-aminopiperid-2-one. All unidentified

peeks correspond to impurities originating either from the solutions utilized or from the HPLC system.

Enzymatic Hydrolysis of L-Arginine

35

3. Enzymatic Hydrolysis of L-Arginine

3.1. Materials and Methods

3.1.1. Bacillus Subtilis Arginase

The Bacillus Subtillis arginase utilized during the enzymatic hydrolysis of L-arginine

experiments was supplied with the K-LARGE commercial kit for L-arginine/urea/ammonia

detection from Megazyme International Ireland Ltd. According to the supplier, the solution has

an activity of 8300 U/ml and contains pure Bacillus Subtillis protein with the addition of

manganese chloride (25 mM) for enzyme activation and of lithium sulphate (2.5 M), used for

precipitation of the protein during purification.

3.1.2. Epoxy-activated Supports

Sepabeads® EC-EP and Sepabeads

® EC-HFA were supplied by Resindion Srl

(Mitsubishi Chemical Corporation). Eupergit® C 250 L was supplied by Sigma-Aldrich. For the

certificates of analysis of the supplied Sepabeads supports please consult Appendix G.

3.1.3. Reagents

Table 3.1. Reagents utilized during the course of the enzymatic hydrolysis of L-arginine experiments.

Chemical Supplier Purity (min)

Acetic acid Riedel-de-Haen 99.8%

Acetonitrile Merck 99.9%

Borax Riedel-de-Haen 99.5%

Boric acid Merck 99.5%

Bradford Dye Reagent Bio-Rad -

di-Sodium hydrogenphosphate Merck 99%

Fluorescamine (Fluram®) Sigma-Aldrich 99%

Glycine Merck 99.7%

L-Arginine Sigma-Aldrich 99%

L-Ornithine Sigma-Aldrich 99%

Manganese (II) Chloride Sigma-Aldrich 98%

Sodium Acetate Merck 99%

Sodium Azide Merck 99%

Sodium dihydrogenphosphate Merck 98%

Triethanolamide Sigma-Aldrich 98%


36

3.1.4. Solutions

Table 3.2. Solutions prepared during the course of the enzymatic hydrolysis of L-arginine experiments.

Solution Composition

Activation buffer 50 mM triethanolamide in Milli-Q water

1 Mm manganese (II) chloride added

pH adjusted to 8.0 with glacial acetic acid

Derivatization buffer 0.1 M borate buffer, pH=9.0

(addition of a 6.2 g/l solution of boric acid in Milli-Q

water to a 38.15 g/l solution of borax in Milli-Q

water until pH = 9 .0 is obtained)

Eluent A 50 mM sodium acetate in Milli-Q water

pH adjusted to 4.5 with concentrated acetic acid

Fluorescamine solution 10 mg of fluorescamine in 20 ml of acetonitrile

Substrate solution 250 mM L-Arginine in Milli-Q water

pH adjusted to 9.5 with glacial acetic acid

3.1.5. Equipment

High Performance Liquid Chromatography

Reverse-phase HPLC analyses were performed on a Waters™ System consisting of:

Waters™ 600s Controller; Waters™ In-Line Degasser; Waters™ 616 Pump; Waters™

717plus Autosampler; Jasco® 820-FP Intelligent Spectrofluorometer; Nova-Pak

® C18 column

60 Ǻ 3.9 x 150 mm with a Nova-Pak® C18 4 µm Guard-Pak™ pre-column insert.

Rotator

During immobilization experiments a Cole-Parmer Roto-Torque model 7637-10

Heavy Duty Rotator was utilized. This rotator allows the gentle but efficient mixing, at variable

speed, of flasks, bottles, and test tubes with different shapes and sizes. The angle of rotation

is adjustable.

Figure 3.1. Cole-Parmer Roto-Torque model 7637-10 Heavy Duty Rototator.


37

Thermomixer

An Eppendorf® Thermomixer Comfort was utilized. This thermomixer is equipped with

a rack that allows the simultaneous heating of up to 24 1.5 ml eppendorf Safe-Lock tubes. It is

fully programmable, capable of heating or cooling samples from 1°C to 99°C and of agitating

from 300 rpm to 1500 rpm (also has a no mixing mode).

3.1.6. Analytical Techniques

Fluorescamine Derivatization

Previously to HPLC analysis, a fluorescamine derivatization procedure was applied to

each sample. This pre-column derivatization method allows the efficient separation and

fluorescence detection of amino acids.

To 5 µl aliquots of each sample were added 75 µl of the derivatization buffer and 20

µl of fluorescamine solution, immediately followed by thorough mixing. 100 µl of the

derivatized samples were used for HPLC analysis.

HPLC Analysis

After derivatization, the samples were analyzed on the previously described reverse-

phase HPLC Waters™ system. 10 µl per sample were injected and the column was eluted at

30°C with a flow rate of 1 ml/min. The gradient of the eluents utilized is described in table 3.3,

eluent B consists of pure acetonitrile of liquid chromatography gradient grade. The total time

of analysis for each sample was of 20 minutes.

Table 3.3. Gradient of eluents applied during HPLC analysis.

Time % Eluent A % Eluent B Curvea

0 80 20 6

6 80 20 6

6.5 60 40 6

9.5 60 40 6

15 80 20 6

a for the slope of the different curves see Appendix A.

Bradford Assay

5 µl of each sample were pipeted to separate wells of a 96 wells microplate. To each

well was added 250 µl of Quick Start Bradford Dye Reagent (Bio-Rad). Absorbance was

immediately measured at 595 nm and compared with a simultaneously obtained standard

curve from Bovine Serum Albumin. Each assay was performed in triplicate.


38

3.1.7. Preparation of Arginase Stock Solution

An arginase stock solution was prepared diluting 0.8 ml of enzyme solution to a total

volume of 5.5 ml with Milli-Q water. Sodium azide was added (0.05% mass) to inhibit

microbial growth. The solution was filter sterilized (0.2 µm filter) and samples were collected

for Bradford and activity assays.

For determination of the solution’s activity, the L-arginine and L-ornithine

concentrations were monitored during an activity assay. A portion of the enzyme stock

solution was diluted 25 times with Milli-Q Water and 300 µl of the dilution were incubated with

300 µl of Milli-Q water for 5 minutes at 37°C in a thermomixer (agitation of 1200 rpm). After

the incubation time, 400 µl of the substrate solution were added and the clock was started.

The reaction was carried in the thermomixer at 37°C. 10 µl samples were taken at 2, 4, 6 and

8 minutes. The samples were immediately quenched with 150 µl of 1 M acetic acid. 100 µl of

the quenched samples were diluted with 400 µl of Milli-Q water and stored in the fridge (5˚C)

until fluorescamine derivatization and HPLC analysis.

3.1.8. Immobilization of Arginase in Different Epoxy-activated Supports

Immobilization in Sepabeads EC-HFA

For the immobilization in Sepabeads EC-HFA, 0.5 ml of arginase stock solution were

added to 7.5 of filter sterilized (0.2 µm filter) 10 mM sodium phosphate buffer, pH=8.0. After

gentle mixing, the solution was added to a sterile 10 ml test tube containing 2.2 g (wet weight;

1 g dry weight) of support, the tube was immediately placed on the rotator at slow rotation

speed and the clock was started. The immobilization was carried for 24 hours with 60 µl

samples of the supernatant being collected at 0, 1, 2, 4, 6, 23 and 24 hours. 30 µl of each

sample were immediately utilized for an activity assay, while the remaining of the sample was

stored in the fridge (5˚C) for posterior Bradford assay. The immobilization was conducted in

duplicate and in the presence of a blank (also in duplicate) containing the same enzyme

solution/buffer ratio but no support added.

Immobilization in Sepabeads EC-EP

For the immobilization in Sepabeads EC-EP, 0.5 ml of arginase stock solution were

added to 7.5 of filter sterilized (0.2 µm filter) 0.8 M sodium phosphate buffer, pH=8.0. After

gentle mixing, the solution was added to a sterile 10 ml test tube containing 2.4 g (wet weight,

1 g dry weight) of support, the tube was immediately placed on the rotator at slow rotation

speed and the clock was started. The immobilization was carried for 24 hours with 60 µl

samples of the supernatant being collected at 0, 1, 2, 4, 6, 23 and 24 hours. 30 µl of each

sample were immediately utilized for an activity assay, while the remaining of the sample was

stored in the fridge (5˚C) for posterior Bradford assay. The immobilization was conducted in


39

duplicate and in the presence of a blank (also in duplicate) containing the same enzyme

solution/buffer ratio but no support added.

Immobilization in Eupergit

For the immobilization in Eupergit C 250, 0.5 ml of arginase stock solution were

added to 7.5 of filter sterilized (0.2 µm filter) 0.8 M sodium phosphate buffer, pH=8.0. After

gentle mixing, the solution was added to a sterile 10 ml test tube containing 1 g (dry weight) of

support, the tube was immediately placed on the rotator at slow rotation speed and the clock

was started. The immobilization was carried for 24 hours with 70 µl samples of the

supernatant being collected at 0, 1, 2, 4, 6, 23 and 24 hours. The samples were centrifuged (1

minute at 1200 rpm) and 60 µl of the supernatant were collected. 30 µl of each sample were

immediately utilized for an activity assay, while the remaining of the sample was stored in the

fridge (5˚C) for posterior Bradford assay. The immobilization was conducted in duplicate and

in the presence of a blank (also in duplicate) containing the same enzyme solution/buffer ratio

but no support added.

Soluble Arginase Activity Assay

For the activity assay of the samples collected during the immobilizations, 30 µl of the

sample was incubated with 30 µl of Milli-Q water for 5 minutes at 37°C in a thermomixer

(agitation of 1200 rpm). After the incubation time, 40 µl of the substrate solution were added

and the clock was started. The reaction was carried in the thermomixer at 37°C. 10 µl

samples were taken at 2, 4, 6 and 8 minutes. The samples were immediately quenched with

150 µl of 1 M acetic acid. 100 µl of the quenched samples were diluted with 400 µl of Milli-Q

water and stored in the fridge (5˚C) until fluorescamine derivatization and HPLC analysis.

Washing

After the immobilization, all supports containing the immobilized enzyme were

washed and stored following the same protocol. The suspensions were filtered using a

sintered glass filter. The filtrate was rinsed with 8 ml of 50 mM sodium phosphate buffer

(pH=8.0) and filtrated once more. The filtered support was collected on a 10 ml sterile test

tube, to which was added 8 ml of sterile 50 mM sodium phosphate buffer (pH=8.0), and

placed on the rotator at slow rotation speed for 45 minutes. After this washing step, the

filtering/rinsing procedure was repeated and the dry support with the immobilized arginase

was stored in the fridge (5˚C). After each filtration step a 1 ml sample of the supernatant was

collected for posterior analysis.

Blockage of the Remaining Epoxy Groups

After immobilization, immobilized arginase can be submitted to the blockage of the

epoxy groups on the support surface that did not react with the enzyme. With this objective,

portions of the three supports containing the immobilized arginase were submitted to a similar


40

blockage procedure. 0.5 g of each support were placed in a 10 ml sterile test tube to which

were added 4 ml of 3 M glycine in 50 mM sodium phosphate buffer (pH=8, filter sterilized).

The suspensions were place in the rotator at slow rotation speed. After 18h the blocked

supports were washed and stored accordingly to the previously described procedure.

Immobilized Arginase Activity Assay

The activity of the enzyme immobilized in the different supports was assayed through

an activity assay comparable to the one utilized for soluble enzyme. 25 mg of the supports

containing immobilized arginase were incubated with 600 µl of activation buffer for 5 minutes

at 37°C in a thermomixer (agitation of 1200 rpm). After the incubation time, 400 µl of the

substrate solution were added and the clock was started. The reaction was carried in the

thermomixer at 37°C. 10 µl samples were taken at 2, 4, 6 and 8 minutes. The samples were

immediately quenched with 150 µl of 1 M acetic acid. 100 µl of the quenched samples were

diluted with 400 µl of Milli-Q water and stored in the fridge (5˚C) until fluorescamine

derivatization and HPLC analysis.

3.1.9. Thermal Stability of Immobilized Arginase

This experiment was conducted with the objective of studying the effect of

immobilization in the stability of arginase. Arginase immobilized in the three tested supports

and in soluble form was incubated at 60°C for different periods.

Multiple 25 mg portions of each of the three supports containing the immobilized

enzyme (with and without blockage of the remaining epoxy groups) were suspended in 600 µl

of activation buffer water in 1.5 ml test tubes. A set of tubes containing the soluble enzyme

was also prepared by diluting 20 µl of arginase stock solution with 580 µl of activation buffer.

The 7 sets of tubes were placed on a water-bath and the clock was stared. Periodically, a

tube from each set was withdrawn and the remaining activity of the immobilized arginase was

immediately assayed at 60°C as previously described. Tubes were assayed after 1, 2, 4, 6,

16 and 24 hours of incubation at 60°C.

3.2. Results and Discussion

3.2.1. Characterization of the Arginase Stock Solution

The exact protein concentration and activity in the Bacilus subtillis arginase stock

solution where determined by Bradford and activity assays respectively. The Bradford assay

gave a protein concentration of 4.1 mg/ml. This value together with the soluble enzyme


41

activity assay, which reported an activity of 325 U/ml, gives a specific activity of 80 U/mg-

protein in the arginase stock solution. One unit is defined as one mol of L-ornithine produced

per minute at pH=9.5 and 37°C.

The value of specific activity obtained is only 9% of the value obtained by Nakamura

et al.32

for purified Bacilus subtillis arginase (858 U/mg). This low value can not be explained

by the relatively low concentration of manganese ions. The Mn2+

concentration in the stock

solution is close to 7 mM, well above the concentration suggested by Nakamura et al.32

for full

enzyme activation (3 mM). A possible explanation can be the use of more aggressive

purification methods that damages the enzyme’s structure. The purification methods

employed were not specified by the supplier.

A SDS-PAGE gel of the original arginase solution was prepared in order to verify its

purity. The run was done in the presence of a reducing agent, guarantying that the enzyme is

dissociated to its sub-units.

Figure 3.2. SDS-PAGE gel of the original arginase solution. Lane (A) corresponds to a 100X dilution

while lane (B) corresponds to a 200X dilution. Lane (M) contains the molecular markers identified with

the corresponding molecular weights in Daltons.

A strong band is observed slightly below 38.000 Da. Due to the presence of reducing

agents, this band should correspond to the monomeric units of arginase and is in accordance

with the range of values reported for the molecular weight of monomers of other bacterial

arginases, 31.000-34.000 Da53,54

. This value also suggests that the enzyme purified by

Nakamura et al.32

was probably in the trimeric form, as a molecular weight of 115.000±5.000

Da was obtained by this author for the purified native arginase.

The only other bands found on the arginase lanes are located around 70.000 Da and

110.000 Da (only observed in the 100X dilution lane). These bands should correspond to the

dimeric and trimeric forms of the enzyme, showing that arginase is not fully dissociated in the


42

presence of the reducing agent. This leads to the conclusion that the arginase solution

supplied is highly pure.

3.2.2. Immobilization of Arginase in Epoxy-activated Supports

The arginase covalent immobilization in Sepabeads EC-HFA, Sepabeads EC-EP and

Eupergit C epoxy-activated supports was followed for 24 hours. Figure 3.3 shows the

evolution of protein concentration in the supernatant during the course of the immobilization

step.

Sepabeads EC-HFA

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Pro

tein

Concentr

atio

n (

µg/m

L)

Sepabeads EC-HFA

Blank

Sepabeads EC-EP

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Pro

tein

Co

nce

ntr

atio

n (

µg

/mL

)

Sepabeads EC-EP

Blank

Eupergit C

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Pro

tein

Co

nce

ntr

atio

n (

µg

/mL

)

Eupergit C

Blank

Figure 3.3. Protein concentration in the supernatant during the course of arginase immobilization in

different epoxy-activated supports. The percentages are based on the initial (t=0) protein concentration.

Error bars calculated using standard deviation of the two duplicate experiments.

The immobilization course of the enzyme was also followed by measuring the

remaining supernatant activity, as shown on figure 3.4.


43

Sepabeads EC-HFA

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Su

pe

rna

tan

t A

ctiv

ity

(%)

Sepabeads EC-HFA

Blank

Sepabeads EC-EP

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Su

pe

rna

tan

t A

ctivi

ty (

%)

Sepabeads EC-EP

Blank

Eupergit C

0%

20%

40%

60%

80%

100%

0 5 10 15 20 25

Time (h)

Su

pe

rna

tan

t Activ

ity (

%)

Eupergit C

Blank

Figure 3.4. Arginase activity in the supernatant during the course of arginase immobilization in different

epoxy-activated supports. The percentages are based on the initial (t=0) protein concentration. Error

bars calculated using standard deviation of the two duplicate experiments.

The immobilization of arginase in the three different supports was clearly successful.

After one hour no protein was found in the supernatant of all suspensions (figure 3.3),

indicating that all enzyme is bound to the supports. The analysis of the supernatant activity

(figure 3.4) shows comparable results. After 24 hours of incubation only the suspension with

Sepabeads EC-EP shows slight remaining activity in the supernatant.

Previous studies48

showed similarly successful results with Sepabeads EC-HFA, the

complete immobilization of different enzymes is observed after 1 to 6 hours. In the case of

Sepabeads EC-EP and Eupergit C, the results obtained for arginase immobilization were

better than the results reported for other enzymes. Using these supports and similar

incubation conditions for covalent immobilization of various enzymes, Mateo et al.48

observed

an immobilization inferior to 65% of the enzyme after 25 hours of incubation.

After 24 hours, low concentrations of protein are detected on the supernatants of

Sepabeads EC-EP and Eupergit C immobilizations, suggesting that arginase might be only

temporarily bond to this supports. This apparent release of arginase from both short-chained

epoxy group supports is contained in the experimental error margin for this experiment, but


44

the immobilization should be followed for a longer period in order to clarify the results. If the

release of arginine from the supports after longer periods of incubation is confirmed, a

possible explanation can be damage inflicted to the structure of the beads by mixing or the

high concentration of sodium phosphate buffer (0.8 M). This seems unlikely, as the mixing

should also affect the Sepabeads EC-HFA beads and previous work with higher

concentration of sodium phosphate buffer during incubation did not show similar problems43

.

The blanks have the exact same composition of the immobilization suspensions

without the addition of support: arginase in 10 mM sodium phosphate buffer for Sepabeads

EC-HFA immobilization; arginase in 0.8 M sodium phosphate buffer for Sepabeads EC-EP

and Eupergit C 250 L immobilization. The evolution of protein concentration and enzyme

activity in the blanks shows a slight decreasing pattern. Again, this apparently abnormal

observation can be due to experimental error. Still, it is possible that 24 hours of incubation at

room temperature in the presence of a buffer can lead to minor degradation of the enzyme,

affecting the Bradford assay and the enzyme’s catalytic activity.

After incubation for 24 hours in the presence of arginase, a washing protocol was

applied to the support beads. During this procedure, the supernatants of the filtration steps

were analyzed in order to detect any protein that might be released from the beads (figure

3.5). In a first observation, the analysis confirms the presence of small amounts of enzyme in

the supernatants of Sepabeads EC-EP and Eupergit C suspensions at the end of incubation

time, similarly to what was observed in figure 3.3. Moreover, during the first rinse step it is

clear that some enzyme is released from the Eupergit C beads, showing that not all arginase

is covalently bound to the support. The protein that is released when the beads are rinsed

with 50 mM sodium phosphate buffer has yet to complete the second step of the binding

mechanism, and is only physically adsorbed to the support’s surface by hydrophobic

interactions. When in contact with a buffer with lower ionic strength (50 mM as opposed to

800 mM), this interactions are weaker and the enzyme is released. The presence of residual,

non-covalently bound enzyme after incubation has previously been reported for Eupergit C

supports46

.

Arginase Immobilization - Rinse/Wash Steps

0,0

10,0

20,0

30,0

40,0

50,0

60,0

Supernatant Rinse 1 Wash Rinse 2

Pro

tein

Con

centra

tion (µ

g/m

L) Sepabeads EC-HFA

Sepabeads EC-EP

Eupergit C

Figure 3.5. Protein content in the filtration supernatant after each step of the beads washing procedure.

Error bars calculated using standard deviation of the two duplicate experiments.


45

Finally, it should be noted that both Sepabeads supports were easily handled, leading

to minimal support loss during the immobilization of arginase in these supports. However,

Eupergit C 250 L originates viscous suspensions, making the manipulation of the beads more

difficult. This led to a considerable loss of Eupergit C support during the washing steps. The

initial and recovered masses of all supports during the immobilization and blockage steps can

be consulted in Appendix G.

3.2.3. Recovered Activity of Immobilized Arginase

The recovered activity exhibited by the enzyme immobilized in the different supports

was compared with the soluble form. The support beads and soluble enzyme were assayed in

the presence of a 50 mM triethanolamide/acetate buffer (pH=8.0) with 1mM Mn2+

. This buffer

is utilized in order to guarantee that the pH and manganese concentration values are similar

in the different assays, as the soluble enzyme solution is already buffered and contains a

relatively high concentration of manganese ions.

For the calculation of the amount of enzyme immobilized in each milligram of support

the following assumptions were made: 100% of the enzyme in solution during incubation was

immobilized in the Sepabeads EC-HFA support, while only 95% was permanently

immobilized in the Sepabeads EC-EP and Eupregit C supports; 100% of the Sepabeads

supports were recovered after the washing steps, while only 95% of the initial mass of

Eupergit C support was recovered.

Recovered Activity of Immobilized Arginase

0

20

40

60

80

100

Sepabeads EC-HFA Sepabeads EC-EP Eupergit C 250 L

Re

co

vere

d A

ctiv

ity (

%) free epoxy groups

epoxy groups

blocked with glycine

Figure 3.6. Recovered activity of arginase immobilized in different epoxy supports. Percentages are

calculated by comparison with the soluble form. Error bars calculated using standard deviation of the

two duplicate experiments.

As expected, the covalent immobilization greatly influences the catalytic activity of

arginase (figure 3.6). The multi-point reaction between arginase and the support deforms the

enzyme, altering the shape of its active site and, possibly, affecting manganese uptake. The

values of recovered activity obtained for immobilization of arginase in the different epoxy


46

supports – from 43% to 61% – are low but acceptable. The results described in literature for

immobilization of other enzymes on similar supports are extremely irregular. For example, in

the case of ß-galactosidase values of recovered activity range from 15% to 100% depending

on the organism of origin and type of epoxy support utilized48

.

Despite the rather inconsistent results (see error bars), it is clear that the treatment

with glycine decreases the recovered activity of the immobilized arginase. The opposite

results were expected, as glycine was added with the objective of reacting with the epoxy

groups that remained free after incubation time. This would stop the covalent-binding

reaction, preventing excessive enzyme/support interaction that could destabilize the enzymes’

active site. One possible explanation for the abnormal results is that glycine might directly

interact with arginase’s active site, interfering with the catalytic mechanism.

The effect of the type of epoxy-activated support on the activity of immobilized

arginase is not clear. Previous studies show that different supports can have considerably

different effect on the immobilized enzyme activity48

. This may be related to the distinct

hydrophobicity of the surface of the various supports. The stronger or weaker hydrophobic

interactions between the support and the hydrophobic residues can affect the orientation of

the enzyme and of its active site, affecting the catalytic activity. The length of the epoxy

groups’ spacer arms (longer in Sepabeads EC-HFA) can also be a factor on the orientation of

the enzyme.

Finally, it is possible that the immobilization affects the quaternary structure of the

enzyme, resulting in the dissociation of arginase to its sub-units. If this is the case,

immobilized arginase monomers can still show residual activity and previous work26

indicates

that catalytic activity can be fully restored by the addition of soluble monomers. The

referenced work was performed using human arginase, further experimentation is necessary.

3.2.4. Thermal Stability of Immobilized Arginase

The stability of the immobilized arginase derivatives was analyzed by following the

residual activity of the enzyme during incubation at 60°C. The activity of arginase immobilized

in the three tested supports (with and without blockage with glycine) and of soluble arginase

was assayed after periods of incubation at 60°C up to 24 hours. The results are illustrated in

figure 3.4. To prevent minor pH variations that may have a significant effect in the decrease of

specific activity at high temperatures, the beads were incubated in the presence of a 50 mM

triethanolamide/acetate buffer (pH=8.0) with 1mM Mn2+

. The addition of manganese ions

slows the deactivation of the enzyme (results no shown). In the absence of the metal ion the

deactivation is too fast, not allowing accurate comparison of the influence of immobilization in

the different supports.


47

Figure 3.7. Evolution of residual activity of arginase immobilized in different epoxy supports

compared to soluble arginase during the course of 6 hours incubation at 60°C. The percentages are

based on the initial (t=0) enzyme activity.

Initial interpretation of the results suggests that the blockage of the non-reactive

epoxy groups with glycine has a positive effect in immobilized enzyme stability. The blocked

supports with short-chained epoxy groups (Sepabeads EC-EP and Eupergit 250 C) appear to

retain their activity during the first 6 hours of incubation at 60°C. However, the graphics are

misleading. Even though the activity in these supports remains constant during the monitored

time period, its initial values are low when compared with the non-blocked supports. The low

ornithine concentrations produced not only aggravate the inherent experimental error, but

also difficult HPLC analysis and chromatogram integration. Thus, the residual activity values


48

obtained for the blocked Sepabeads EC-EP and Eupergit 250 C supports are not reliable and

should not be taken in much consideration. Possible solutions for this problem are the use of

more support during activity assays, less dilution of the samples and duplicate experiments.

Concerning the effect of the other tested supports, only the covalent-binding to

Sepabeads EC-EP without glycine blockage shows a mild positive effect in arginase thermal

stability. The immobilization in this supports leads to a 20 % increase in residual activity when

compared with the soluble enzyme after 6 hours of incubation at 60°C.

The immobilization of arginase in Sepabeads EC-HFA and Eupergit C 250 L appears

to have a negative effect in the enzyme’s stability. The deactivation of the immobilized

derivatives for both these supports is faster than the deactivation observed for the free

enzyme, although the blockage of Sepabeads EC-HFA with glycine seems to lead to an

increase in the derivatives stability.

The results are not in accordance with the results reported by Mateo et al.48

. This

author claims that the immobilization of different enzymes in all the three tested supports

consistently increases the enzymes’ thermal stability. Again, the results showed in the present

study are not extremely reliable and future duplicate experiments are essential to draw

objective conclusions.

3.2.4. Analytical Methods

The fluorescamine derivatization and HPLC analysis method utilized were suitable for

the experiments performed. The peeks obtained in the chromatogram show good separation

and are of easy integration (figure 3.5). The concentrations of arginine and ornithine in the

analyzed samples were calculated without difficulties using a linear standard curve.

Figure 3.8. Example of a typical HPLC chromatogram obtained for an activity assay sample.


49

Concerning the Bradford assay, it was verified that high buffer concentrations

interfere with the absorbance reading. This interference was observed during the analysis of

samples from immobilization of arginase in Sepabeads EC-EP and Eupergit C 250 (0.8 M

sodium phosphate buffer). The results were corrected accordingly.

Conclusions and Future Perspectives

51

4. Conclusions and Future Perspectives


The results obtained show that the chemical catalysis of arginine hydrolysis is not, at

this moment, a suitable method for ornithine production on an industrial scale. The maximum

yield of ornithine obtained was 15.2% at 125°C with pH initially adjusted to pH=12 with

sodium hydroxide. Not only were the yields obtained disappointing when compared to the

values mentioned in literature (90% by Murray et al.13

), but also the optimization of the

reaction would probably involve the use of buffers or the constant correction of pH with acid or

alkali, techniques undesirable in an industrial process. Nonetheless, on a lab scale study

these changes to the protocol could produce interesting results. The addition of small

quantities of acid or alkali in certain moments of the reaction for pH correction would require

the alteration of the experimental setting, as the reactors utilized do not allow this procedure.

A potential alternative for industry could be manipulation of pH through the continuous

separation of the different components of the reaction mixture.

Unfortunately, the high temperatures employed do not allow the recovery of urea, a

considerably valuable side product. Nonetheless, the production of ammonia can still provide

a substantial source of income in an industrial scale process.

The tested metal salts showed no influence either on arginine consumption rate or

ornithine yield.

The solid catalyst utilized, NaY zeolite, showed limited catalytic effects even at high

concentrations (5 g/l). It also displays the tendency to accumulate in the reactor walls, which

would generate serious complications in industry. As an alternative, other materials with

catalytic properties could be utilized including: zeolites ion-exchanged with metal ions that

generate stronger basic sites (Cs+, Rb

+, K

+); alkaline earth oxides; diverse heterogeneous

superbasic catalysts19

.


The results obtained were promising, with Bacillus subtllis arginase being

successfully immobilized in three different supports suitable for industrial application. The

covalent binding to the three tested supports did not show a significant increase in arginase’s

thermal stability and the activity of the immobilized derivatives is considerably low when

compared with the soluble form. The blockage of the un-reactive epoxy groups with glycine

did not show significant increase in stability/activity. Similar results were obtained for all the

tested supports, however, Sepabeads supports are of easier handling and should be

preferred over Eupergit C 250 L.

Further research should focus on identifying the operational conditions that maximize

the production of ornithine and the stability of immobilized arginase. Bacillus subtilis

arginase’s properties have not been the subject of extensive studies. Thus, experimental work

should be realized to determine: optimum pH and temperature; effects of product inhibition;

Conclusions and Future Perspectives

52

optimum Mn2+

concentrations. The manganese concentration is a particularly important factor

when considering industrial-scale application, as high salt concentrations can interfere with

downstream processing. Naturally, the next research step would be the lab-scale simulation

of a batch or continuous process for ornithine production with recovery and re-utilization of the

immobilized arginase

Finally, it should be noted that any possible industrial application of arginase would

require the large scale production of this enzyme at affordable prices. Presently the enzyme is

only available in diagnostic quantities at high expenses, being usually purified from mammal

liver.

Final Remarks

The research done on the chemical and enzymatic catalysis of the hydrolysis of

L-arginine can not be objectively compared. While the experimental setup utilized to study the

chemically catalyzed reaction is similar to a potential industrial process, the work realized on

the enzymatic conversion was primarily focused on the immobilization of arginase in industrial

suitable supports and not on the optimization of L-ornithine production.

Nonetheless, the biocatalysis approach seems to be the be the most promising. The

arginase catalyzed reaction is very clean when compared to the alkali catalyzed reaction, with

the only secondary product produced being urea. Of course, urea is actually an economically

attractive side product that cannot be obtained from the alkali catalyzed conversion of

L-arginine due to the high temperatures employed. The major drawbacks of arginase

application to a large scale process are the already mentioned dependence on manganese

ions and limited availability.

References

53

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Guisan, J. M.; Tam, A.; Daminati, M., Epoxy Sepabeads: A novel epoxy support for stabilization of industrial enzymes via very intense multipoint covalent attachment. Biotechnology Progress 2002, 18, (3), 629-634.

44. Fernandez-Lafuente, R.; Rodri?uez, V.; Mateo, C.; Penzol, G.; Herna?dez-Justiz, O.;

Irazoqui, G.; Villarino, A.; Ovsejevi, K.; Batista, F.; Guisa?n, J. M., Stabilization of multimeric enzymes via immobilization and post-immobilization techniques. Journal of Molecular Catalysis - B Enzymatic 1999, 7, (1-4), 181-189.

45. www.resindion.com/sepabeadsec/sepabeadsec.html. 46. Boller, T.; Meier, C.; Menzler, S., EUPERGIT Oxirane Acrylic Beads: How to Make

Enzymes Fit for Biocatalysis. Organic Process Research & Development. 2002, 6, (4), 509-519.

47. Hilterhaus, L.; Minow, B.; Mu?ler, J.; Berheide, M.; Quitmann, H.; Katzer, M.; Thum,

O.; Antranikian, G.; Zeng, A. P.; Liese, A., Practical application of different enzymes immobilized on sepabeads. Bioprocess and Biosystems Engineering 2008, 31, (3), 163-171.

48. Mateo, C.; Torres, R.; Fernandez-Lorente, G.; Ortiz, C.; Fuentes, M.; Hidalgo, A.;

Lopez-Gallego, F.; Abian, O.; Palomo, J. M.; Betancor, L.; Pessela, B. C. C.; Guisan, J. M.; Fernandez-Lafuente, R., Epoxy-amino groups: A new tool for improved immobilization of proteins by the epoxy method. Biomacromolecules 2003, 4, (3), 772-777.

References

56

49. Krause, I., Simultaneous determination of amino acids and biogenic amines by reversed-phase high-performance liquid chromatography of the dabsyl derivatives. Journal of chromatography. B, Biomedical applications 1995, 715, (1), 67.

50. Duerr, B. F.; Czarnika, W., Copper (II)-catalysed hydrolysis of an unactivated amide.

Application of the groves' rule to the hydrolysis of acrylamide. Journal of the Chemical Society. Chemical communications 1990, (23), 1707-1709.

51. Meriwether, L., Metal Ion Promoted Hydrolysis of Glycine Amide and of

Phenylalanylglycine Amide. Journal of the American Chemical Society 1956, 78, (19), 5119.

52. Buckingham, D. A., Peptide bond formation and subsequent hydrolysis at a cobalt

(III) center. Journal of the American Chemical Society 1967, 89, (11), 2772. 53. Jenkinson, C. P.; Grody, W. W.; Cederbaum, S. D., Comparative properties of

arginases. Comparative Biochemistry and Physiology - B Biochemistry and Molecular Biology 1996, 114, (1), 107-132.

54. Kanda, M.; Ohgishi, K.; Hanawa, T.; Saito, Y., Arginase of Bacillus brevis Nagano:

purification, properties, and implication in gramicidin S biosynthesis. Arch Biochem Biophys 1997, 344, (1), 37-42.

Appendix A

57

Appendix A – Gradient Curves

During the elution of a sample through the HPLC system, the rate of change of

solvent composition over time depends on the curve number and the length of the gradient

segment. The gradient curve profile specified in each row of the gradient table (table 2.3)

affects both solvent composition and flow rate. The different curves identified by a specific

number are represented in figure A.1.

Figure A.1. Different gradient curves identified by the input number.

Appendix B

58

Appendix B – NMR Spectra of L-Arginine

177.0

880

159.7

801

57.0

779

43.4

372

30.4

308

26.8

074

-0.0

000

(ppm)

020406080100120140160180

*** Current Data Parameters ***

NAME : abg-acid

EXPNO : 2

PROCNO : 1

*** Acquisition Parameters ***

DATE_t : 10:08:47

DATE_d : Aug 07 2008

NS : 1024

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm

*** Processing Parameters ***

GB : 0.0000000

LB : 1.00 Hz

OFFSET : 222.3916 ppm

SI : 32768

*** 1D NMR Plot Parameters ***

Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

ARG-acid (13C NMR in D2O + D2SO4 + TSP)

Figure B.1. 13

C-NMR spectrum of L-arginine.

57

.07

06

43

.42

99

30

.42

35

26

.80

01

(ppm)

020406080100120140160180


NAME : abg-acid

EXPNO : 3

PROCNO : 1


DATE_t : 10:24:32


NS : 256

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

ARG-acid (DEPT135 in D2O + D2SO4 + TSP)

Figure B.2. DEPT spectrum of L-arginine.

Appendix B

59

1.0

00

0

2.0

48

0

2.0

38

1

2.0

67

7

Inte

gra

l

3.8

18

1

3.8

03

0

3.7

87

4

3.2

55

5

3.2

38

6

3.2

21

0

1.9

39

8

1.9

30

4

1.9

14

1

1.8

98

4

1.7

67

9

1.7

51

0

1.7

34

7

1.7

10

9

1.6

87

7

1.6

70

7

1.6

53

8

1.6

44

4

(ppm)

1.61.82.02.22.42.62.83.03.23.43.63.8


NAME : ABG-ACID

EXPNO : 1

PROCNO : 1


DATE_t : 09:07:56


NS : 16

NUCLEUS : off

PARMODE : 1D

SW : 20.5503 ppm


GB : 0.0000000

LB : 0.30 Hz


SI : 32768


Height : 13.78 cm

Start : 4.00 ppm

Stop : 1.50 ppm

ppm_cm : 0.11

AQ_time : 3.9845890 sec

NUCLEUS : off

Figure B.3.

1H-NMR spectrum of L-arginine.

Appendix C

60

Appendix C – NMR Spectra of L-Ornithine

176.8

911

56.9

612

41.7

312

30.2

413

25.5

899

-0.0

000

(ppm)

020406080100120140160180


NAME : orn

EXPNO : 2

PROCNO : 1


DATE_t : 16:57:48

DATE_d : Jun 19 2008

NS : 1024

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -4.99 ppm

ppm_cm : 9.28

AQ_time : 1.3631490 sec

NUCLEUS : off

ORN (13C NMR in D2O + TSP)

Figure C.1. 13

C-NMR spectrum of L-ornithine.

56

.95

39

41

.72

39

30

.23

40

25

.57

53

(ppm)

102030405060708090100110120130140150160170180190


NAME : orn

EXPNO : 3

PROCNO : 1


DATE_t : 17:13:01

DATE_d : Jun 19 2008

NS : 256

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : 0.01 ppm

ppm_cm : 9.05

AQ_time : 1.3631490 sec

NUCLEUS : off

ORN (DEPT135 in D2O + TSP)

Figure C.2. DEPT spectrum of L-ornithine.

Appendix C

61

1.0

00

0

2.0

14

5

2.0

78

4

2.0

32

8

Inte

gra

l

4.8

17

8

4.7

65

7

4.7

18

7

4.1

09

7

4.0

94

0

4.0

78

4

3.0

89

3

3.0

70

5

3.0

52

3

2.1

00

3

2.0

65

8

2.0

50

8

2.0

35

7

2.0

28

2

2.0

02

5

1.9

87

4

1.9

68

0

1.9

52

3

1.9

27

2

1.9

12

8

1.8

94

0

1.8

80

2

1.8

47

6

1.8

28

8

1.8

13

7

1.7

99

9

0.0

00

0

(ppm)

1.02.03.04.05.06.07.08.09.0


NAME : OBN-ACID

EXPNO : 1

PROCNO : 1


DATE_t : 14:38:06


NS : 16

NUCLEUS : off

PARMODE : 1D

SW : 20.5503 ppm


GB : 0.0000000

LB : 0.30 Hz


SI : 32768


Height : 13.78 cm

Start : 10.00 ppm

Stop : -0.10 ppm

ppm_cm : 0.46

AQ_time : 3.9845890 sec

NUCLEUS : off

Figure C.3.

1H-NMR spectrum of L-ornithine.

Appendix D

62

Appendix D – NMR Spectra of 3-Aminopiperid-2-one

174.3

467

171.7

950

52.3

900

44.2

246

41.7

021

29.7

091

27.5

511

25.6

264

22.7

393

(ppm)

020406080100120140160180


NAME : lac-acid

EXPNO : 2

PROCNO : 1


DATE_t : 14:16:38


NS : 1024

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

LAC-acid (13C NMR in D2O + D2SO4 + TSP)

Figure D.1. 13

C-NMR spectrum of 3-aminopiperid-2-one.

55.1

167

52.3

682

(ppm)

020406080100120140160180


NAME : lac-acid

EXPNO : 3

PROCNO : 1


DATE_t : 14:32:05


NS : 256

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

LAC-acid (DEPT135 in D2O + D2SO4 + TSP)

Figure D.2. DEPT spectrum of 3-aminopiperid-2-one.

Appendix D

63

1.0

00

0

3.2

91

5

6.7

75

8

1.9

92

2

3.4

37

5

5.5

58

6

8.9

42

4

0.4

04

6

Inte

gra

l

4.8

64

2

4.8

12

1

4.7

62

0

4.1

68

0

4.1

52

4

4.1

36

7

4.0

05

0

3.9

78

0

3.3

50

9

3.0

93

7

3.0

74

9

3.0

56

7

2.3

36

1

2.3

07

3

2.0

84

7

2.0

55

2

2.0

25

1

1.9

60

5

1.9

04

7

1.8

80

8

1.8

58

2

1.8

27

5

1.2

36

1

0.0

00

0

(ppm)

1.02.03.04.05.06.07.08.09.0


NAME : LAC-ACID

EXPNO : 1

PROCNO : 1


DATE_t : 13:17:00


NS : 16

NUCLEUS : off

PARMODE : 1D

SW : 20.5503 ppm


GB : 0.0000000

LB : 0.30 Hz


SI : 32768


Height : 13.28 cm

Start : 10.00 ppm

Stop : -0.10 ppm

ppm_cm : 0.46

AQ_time : 3.9845890 sec

NUCLEUS : off

LAC-acid (1H NMR in D2O + D2SO4 + TSP)

Figure D.3.

1H-NMR spectrum of 3-aminopiperid-2-one.

Appendix E

64

Appendix E – NMR Spectra of Putrescine

43

.00

70

31

.70

67

0.0

00

0

(ppm)

020406080100120140160180


NAME : pvt-acid

EXPNO : 2

PROCNO : 1


DATE_t : 12:55:35


NS : 1024

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

PUT-acid 08-08-08 (13C NMR in D2O + D2SO4 + TSP)

Figure E.1. 13

C-NMR spectrum of putrescine.

42.9

998

31.6

994

(ppm)

020406080100120140160180


NAME : pvt-acid

EXPNO : 3

PROCNO : 1


DATE_t : 13:11:01


NS : 256

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

PUT-acid 08-08-08 (DEPT135 in D2O + D2SO4 + TSP)

Figure E.2. DEPT spectrum of putrescine.

Appendix E

65

1.0

00

0

1.0

08

8

Inte

gra

l

4.5

87

0

2.6

15

2

2.5

98

3

2.5

82

6

1.4

53

7

1.4

45

6

1.4

36

2

1.4

28

6

1.4

20

5

-0.0

00

0

(ppm)

1.02.03.04.05.06.07.08.09.0


NAME : pvt-acid

EXPNO : 1

PROCNO : 1


DATE_t : 11:54:59


NS : 16

NUCLEUS : off

PARMODE : 1D

SW : 20.5503 ppm


GB : 0.0000000

LB : 0.30 Hz


SI : 32768


Height : 13.28 cm

Start : 10.00 ppm

Stop : -0.10 ppm

ppm_cm : 0.46

AQ_time : 3.9845890 sec

NUCLEUS : off

PUT-acid 08-08-08 (1H NMR in D2O + D2SO4 + TSP)

Figure E.3. 1H-NMR spectrum of putrescine.

Appendix F

66

Appendix F – NMR Spectra of 30 h Reaction Mixture

17

1.8

38

7

46

.47

01

44

.22

46

43

.38

62

41

.79

68

30

.27

04

30

.05

90

27

.66

04

27

.44

90

26

.76

37

25

.63

36

22

.73

93

-0.0

00

0

(ppm)

020406080100120140160180


NAME : hydr-a~1

EXPNO : 2

PROCNO : 1


DATE_t : 16:58:48


NS : 1024

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

HYDR-ARG30-acid (13C NMR in D2O + D2SO4 +TSP)

Figure F.1. 13

C-NMR spectrum of 30h reaction mixture.

46

.46

28

44

.21

73

43

.37

16

41

.78

23

30

.05

17

27

.43

44

26

.74

91

22

.73

20

(ppm)

020406080100120140160180


NAME : hydr-a~1

EXPNO : 3

PROCNO : 1


DATE_t : 17:14:14


NS : 256

NUCLEUS : off

PARMODE : 1D

SW : 238.8728 ppm


GB : 0.0000000

LB : 1.00 Hz


SI : 32768


Height : 14.78 cm

Start : 200.00 ppm

Stop : -2.00 ppm

ppm_cm : 9.14

AQ_time : 1.3631490 sec

NUCLEUS : off

HYDR-ARG30-acid (DEPT135 in D2O + D2SO4 +TSP)

Figure F.2. DEPT spectrum of 30h reaction mixture.

Appendix F

67

0.4

14

9

0.6

78

4

1.0

00

0

1.2

61

6

16

.63

6

0.8

42

5

3.7

72

2

4.1

40

7

37

.88

9

Inte

gra

l

7.2

80

0

4.7

88

9

4.7

47

5

4.7

11

2

4.2

02

5

3.9

04

6

3.8

89

6

3.8

74

5

3.3

49

0

3.2

95

1

3.2

59

9

3.2

43

6

3.1

39

5

3.0

75

5

3.0

56

7

3.0

39

2

2.3

38

6

2.3

12

3

2.0

21

3

1.9

69

9

1.9

55

5

1.9

43

5

1.9

29

7

1.8

85

2

1.8

61

4

1.8

33

8

1.7

51

0

1.6

75

1

0.0

00

0

(ppm)

1.02.03.04.05.06.07.08.09.0


NAME : HYDR-A~1

EXPNO : 1

PROCNO : 1


DATE_t : 15:59:09


NS : 16

NUCLEUS : off

PARMODE : 1D

SW : 20.5503 ppm


GB : 0.0000000

LB : 0.30 Hz


SI : 32768


Height : 13.28 cm

Start : 10.00 ppm

Stop : -0.10 ppm

ppm_cm : 0.46

AQ_time : 3.9845890 sec

NUCLEUS : off

HYDR-ARG30-acid (1H NMR in D2O + D2SO4 +TSP)

Figure F.3.

1H-NMR spectrum of 30h reaction mixture.

Appendix G

68

Appendix G – Certificates of Analysis

Appendix G

69

Appendix H

70

Appendix H – Recovered Support Masses

Table H.1. Initial and recovered masses of the three tested epoxy-activated supports, including

duplicates, during the immobilization and epoxy groups blockage steps.

Initial Mass (mg) Support

Wet Support Water Content (%) Dry Weight Wet Weight

Recovered Mass (mg)

Sepabeads EC-HFA 54 Duplicate I 1002 2179 2152 Duplicate II 1005 2186 2123

Duplicate I Blocked 230 502 490 Duplicate II Blocked 232 505 510 Sepabeads EC-EP 58

Duplicate I 979 2331 2368 Duplicate II 979 2332 2345

Duplicate I Blocked 217 516 488 Duplicate II Blocked 213 506 569

Eupergit C 250 86

Duplicate I 993 - 3938 Duplicate II 990 - 4024

Duplicate I Blocked 496 - 579 Duplicate II Blocked 498 - 587

hydrolysis of l-arginine â€“ chemical and enzymatic catalysis

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