CATALYTIC HYDROGELS
Ângela Tomaz Francisco Neves
Thesis to obtain the Master of Science Degree in
Biotechnology
Supervisor: Dr. Pedro Carlos de Barros Fernandes
Examination Committee
Chairperson: Prof. Arsénio do Carmo Sales Mendes Fialho
Supervisor: Dr. Pedro Carlos de Barros Fernandes
Members of the Committee: Prof. Maria Henriques Lourenço Ribeiro
November 2015
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Acknowledgements
I would like to thank all the people who contributed to the development of this work. First I would
like to thank my supervisor Doctor Pedro Fernandes for accepting me for this project, for the constant
guidance, support, patience and constructive critiques through the learning process of this master
thesis.
I would like to show my gratitude to my colleagues and friends from the lab for providing a
friendly and cooperative atmosphere at work and also by the useful feedback and insightful comments
on my work.
To my MSc colleagues, especially Alexandra Salvado, Mafalda Cavalheiro e Sara Gomes,
thanks for the constant encouragement, for cheer me up when I have needed it the most, for his love
and comprehension. Finally, words cannot express how grateful I am to my mom, dad, sister,
grandparents and boyfriend for the unequivocal support throughout my academia life, for the love and
encouragement.
Thank you all.
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Abstract
The advantages of enzyme immobilization in biotechnology impelled scientific community to
search for suitable immobilization matrices. Hydrogels have been playing an important role in enzyme
entrapment, due to their chemical and biological properties. Within such scope, this work aimed to
produce and characterize resilient polyvinyl alcohol and chitosan hydrogels, displaying high
operational and thermal stability for inulinase entrapment.
Chitosan beads and polyvinyl alcohol films were obtained, yielding an immobilization efficiency
of roughly 98% and 97%, respectively. The optimum pH of the immobilized enzyme in both matrices
was slightly more basic (5.0) than the one observed for the free enzyme (4.5). An optimum
temperature of 50ºC was displayed for free inulinase, whereas chitosan and polyvinyl alcohol
immobilized inulinase exhibited at 65ºC and 55ºC, respectively. The Michaelis constant (KM) of
inulinase for inulin had a 1.5 and 1.6 fold-increase after immobilization in chitosan and polyvinyl
alcohol, respectively, suggesting an apparent decrease in affinity of inulinase towards inulin.
Nonetheless, the maximum reaction rate (Vmax) did not significantly change after immobilization in
chitosan, whereas 2-fold decrease was observed upon immobilization in polyvinyl alcohol.
Also, thermal stability increased considerably after immobilization in both matrices. Moreover,
encouraging results were obtained for continuous operation. After 47 days at 55ºC a product yield of
91% was achieved for inulinase immobilized in chitosan. Continuous operation of inulinase
immobilized in polyvinyl alcohol obtained 63% of product yield after 31 days at 50ºC.
Overall, relatively simple, robust and low-cost immobilization procedures were established, with
appealing results towards commercial preparations.
Keywords: enzyme immobilization, inulinase, polyvinyl alcohol based hydrogel, chitosan based
hydrogel, hydrolytic activity, continuous operation.
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Resumo
As vantagens da imobilização de enzimas na biotecnologia levaram a comunidade científica à
procura de matrizes adequadas. Os hidrogéis têm evidenciado relevância na oclusão enzimática,
devido às suas características químicas e biológicas. Assim, este trabalho visa produzir e caracterizar
hidrogéis à base de álcool polivinílico e quitosano, que demonstrem uma elevada estabilidade
operacional e térmica para a oclusão da inulinase.
As partículas de quitosano e os filmes de álcool polivinílico produzidos evidenciaram uma
eficiência de imobilização de cerca de 98% e 97%, respectivamente. O pH óptimo da enzima
imobilizada em ambas as matrizes revelou-se ligeiramente mais básico (5,0) do que o observado para
a enzima livre (4,5). Uma temperatura óptima de 50ºC foi atingida para a inulinase livre, enquanto na
imobilização em quitosano e álcool polivinílico foi de 65ºC e de 55ºC, respectivamente. A constante
de Michaelis (KM) para a inulina aumentou 1,5 e 1,6 após a imobilização em quitosano e álcool
polivinílico, traduzindo uma aparente diminuição da afinidade para o substrato. A imobilização em
quitosano não alterou significativamente a velocidade máxima da reacção (Vmax), enquanto a
imobilização em álcool polivinílico promoveu uma diminuição para metade.
A estabilidade térmica aumentou consideravelmente após a imobilização em ambas as
matrizes. Os resultados da estabilidade operacional foram bastante promissores. Após 47 dias à
temperatura de 55ºC registou-se um rendimento de produto de 91% para a inulinase imobilizada em
quitosano, enquanto para o álcool polivinílico registou-se um rendimento de 63% após 31 dias à
temperatura de 50ºC.
Em suma, foram estabelecidos procedimentos relativamente simples, robustos e eficientes,
com resultados promissores num contexto comercial.
Palavras-chave: imobilização de enzimas, inulinase, hidrogel de álcool polivinílico, hidrogel de
quitosano, actividade hidrolítica, operação em contínuo.
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Table of contents
Acknowledgements ...................................................................................................................................i
Abstract.................................................................................................................................................... iii
Resumo ....................................................................................................................................................v
Table of contents .................................................................................................................................... vii
List of figures ........................................................................................................................................... xi
List of tables ........................................................................................................................................... xv
List of abbreviations .............................................................................................................................. xvii
Thesis Outline .......................................................................................................................................... 1
Chapter I. Introduction .......................................................................................................................... 3
I.1. Enzymes as biocatalysts ............................................................................................................... 3
I.1.1. History and importance of enzymes ........................................................................................ 3
I.1.2. Enzyme properties - advantages and constraints ................................................................... 4
I.1.3. Immobilization methods .......................................................................................................... 4
I.1.4. Immobilization matrices ........................................................................................................... 7
I.1.5. Applications of immobilized enzymes and markets perspective ............................................. 9
I.2. Hydrogels ..................................................................................................................................... 10
I.2.1. From characteristics to applications ...................................................................................... 10
I.2.2. Technologies adopted in hydrogel preparation ..................................................................... 12
I.2.3. Polyvinyl alcohol (PVA) based hydrogels ............................................................................. 13
I.2.3.1. Structure and properties of PVA ..................................................................................... 13
I.2.3.2. Synthesis and properties of PVA hydrogels ................................................................... 15
I.2.4. Chitosan (CS) based hydrogels ............................................................................................ 19
I.2.4.1. Structure and properties of CS ....................................................................................... 19
I.2.4.2. Synthesis and properties of CS hydrogels ..................................................................... 20
I.2.5. Applications of PVA and CS hydrogels ................................................................................. 21
I.3. Protein model system - Inulinase enzyme ................................................................................... 22
I.3.1. Inulinase - general considerations ........................................................................................ 22
I.3.2. From inulin to fructose ........................................................................................................... 23
I.3.3. Inulin hydrolysis for fructose production ................................................................................ 24
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I.3.4. Inulinase immobilization - review .......................................................................................... 25
I.4. Types of Reactors ........................................................................................................................ 25
I.4.1. Batch operation ..................................................................................................................... 26
I.4.2. Continuous operation ............................................................................................................ 26
I.4.3. Reactors used with immobilized inulinase ............................................................................ 27
Chapter II. Objectives .......................................................................................................................... 29
Chapter III. Materials and Methods .................................................................................................... 31
III.1. Chemicals and Materials ........................................................................................................... 31
III.2. Analytical methods ..................................................................................................................... 32
III.2.1. Quantification of reducing sugars by DNS method ............................................................. 32
III.2.2. Quantification of reducing sugars by HPLC method ........................................................... 32
III.2.3. Bradford protein quantification ............................................................................................ 32
III.2.4. Protein characterization by SDS-PAGE .............................................................................. 33
III.3. Immobilization procedures ......................................................................................................... 33
III.3.1. Enzyme immobilization in polyvinyl alcohol (PVA) film ....................................................... 33
III.3.2. Enzyme immobilization in chitosan (CS) beads .................................................................. 34
III.4. Immobilization efficiency, protein entrapment and immobilization yield .................................... 35
III.5. Bioconversion studies ................................................................................................................ 36
III.6. pH and temperature profiles ...................................................................................................... 36
III.7. Optimal mass of matrix for conversion ...................................................................................... 37
III.8. Determination of kinetic parameters .......................................................................................... 37
III.9. Stability studies .......................................................................................................................... 37
III.9.1. Thermal stability .................................................................................................................. 37
III.9.2. Storage stability ................................................................................................................... 38
III.9.3. Operational stability ............................................................................................................. 38
III.9.3.1. Determination of flow rate and reactor volume............................................................. 38
III.9.3.2. Continuous production of fructose ................................................................................ 38
Chapter IV. Results and Discussion .................................................................................................. 41
IV.1. Polyvinyl alcohol (PVA) film immobilized inulinase ................................................................... 41
IV.2. Chitosan (CS) beads immobilized inulinase .............................................................................. 47
IV.3. Immobilization parameters ........................................................................................................ 50
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IV.4. Effect of pH on the activity of free and immobilized inulinase ................................................... 51
IV.5. Effect of temperature on the activity of free and immobilized inulinase .................................... 53
IV.6. Mass of support ......................................................................................................................... 54
IV.7. Kinetic study of enzymatic inulin hydrolysis .............................................................................. 55
IV.8. Stability of immobilized enzyme ................................................................................................ 56
IV.8.1. Thermal stability .................................................................................................................. 57
IV.8.2. Storage stability .................................................................................................................. 60
IV.8.3. Operational stability ............................................................................................................ 62
Chapter V. Conclusions and Future Work ........................................................................................ 67
References ............................................................................................................................................ 69
Annexes ................................................................................................................................................. 81
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List of figures
Figure 1. Formation of concentration gradients in porous enzyme carriers. The analyte’s (substrate or
product) concentration in the well-mixed liquid bulk (Cbulk) often differs from the concentration inside
the carrier (Cin). For substrates, Cbulk may be higher than Cin, whereas for products, the situation may
be reversed. From [24]. ........................................................................................................................... 6
Figure 2. Materials used for fabrication of immobilization matrices. Adapted from [26]. ........................ 7
Figure 3. Representative images of each type of immobilization technique, enzyme is represented as
blue circle: covalent immobilization (A), non-covalent immobilization by physical adsorption (B),
entrapment (C), encapsulation (D) and cross-linking (E). ....................................................................... 8
Figure 4. Representative image of a hydrogel. Adapted from [46]. ...................................................... 10
Figure 5. Classification of hydrogels based on the different properties. Adapted from [51]. ................ 11
Figure 6. General hydrogel preparation diagram. ................................................................................ 12
Figure 7. Schematic of the reaction of the PVAc to PVA. In red are the acetate groups whereas in
green are the hydroxyl groups. .............................................................................................................. 14
Figure 8. Degree of hydrolysis (DH) and polymerization (DP) of PVA. ................................................ 14
Figure 9. Effect of molecular weight and % of hydrolysis in the properties of PVA. The variations in
properties with molecular weight are for a constant degree of hydrolysis (mol %), and the effect of
hydrolysis is at a constant molecular weight. From [76]. ....................................................................... 15
Figure 10. Mechanism of the reaction between PVA and GA. From [85]. .......................................... 16
Figure 11. Network structure of PVA gels cross-linked by hydrogen bonds. From [92]. ..................... 17
Figure 12. Reaction of N-deacetylation of chitin (A). The repeating structural unit of CS, showing the
functional groups at positions C-2, C-3 and C-6 (B). Adapted from [107]. ........................................... 19
Figure 13. Interactions of CS with GA (A) and TPP (B). Adapted from [117]. ...................................... 20
Figure 14. Representative crystal structure of an exo-inulinase from Aspergillus awamori. The
structure is a monomer with two domains and glycerol and N-acetyl-D-glucosamine are represented
as ligands. PDB ID 1Y4W...................................................................................................................... 22
Figure 15. Full inulin hydrolysis. Adapted from [152]. .......................................................................... 23
Figure 16. The miniature tubular reactor configuration used for continuous flow operation. ............... 31
Figure 17. Schematic representation of enzyme immobilization in PVA. ............................................. 33
Figure 18. Schematic representation of enzyme immobilization in CS. ............................................... 35
Figure 19. Representation the set-up assembled for the continuous production of fructose. The
miniature tubular reactor was immersed in a temperature-controlled batch and feeding was provided
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through the peristaltic pump. (A) Inlet – 5% (w/v) inulin solution in acetate buffer 100 mM pH 5.0; (B)
Peristaltic pump; (C) Thermostat ECO E4 where is immersed the reactor and is covered to avoid heat
loss (D) Outlet – for DNS and protein analysis...................................................................................... 39
Figure 20. Representative image of PVA film formed (61 kDa with 400 µL of GA) concerning the
conditions previous described, inside the screw vessel (A) or removed from the vessel (B);
Representative image of PVA solution of 61 kDa with 50 µL GA, which was not solidified (C)............ 41
Figure 21. Influence of the volume of GA added on the concentration of fructose (g.L-1
) formed from
inulin hydrolysis at 40ºC. The assay was performed for PVA films of 50 kDa (A), 61 kDa (B) and 145
kDa (C) and different volumes of GA (50 µL, 100 µL, 200 µL and 400 µL). Bioconversion runs were
performed in 5% (w/v) inulin in acetate buffer pH 4.5. 100 mg of immobilized inulinase was used as
biocatalyst. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the beginning of
incubation. Standard deviation did not exceed 5%. .............................................................................. 43
Figure 22. Influence of the volume of GA added on the concentration of fructose (g.L-1
) formed from
inulin hydrolysis at 45ºC (A), 50ºC (B) and 55ºC (C). The assay was performed for PVA films of 50
kDa, 61 kDa and 145 kDa and different volumes of GA (50 µL, 200 µL and 400 µL). Bioconversion
runs were performed in 5% (w/v) inulin in acetate buffer pH 4.5. 100 mg of immobilized inulinase was
used as biocatalyst. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the beginning
of incubation. Standard deviation did not exceed 5%. .......................................................................... 44
Figure 23. Influence of the volume of GA added on the concentration of fructose (g.L-1
) formed from
inulin hydrolysis at 60ºC. The assay was performed for PVA films of 50 kDa and 61 kDa with 400 µL of
GA. Bioconversion runs were performed in 5% (w/v) inulin in acetate buffer pH 4.5. 100 mg of
immobilized inulinase was used as biocatalyst. Samples (10 µL) were collected at 3.5, 6.5 and 24
hours after the beginning of incubation. Standard deviation did not exceed 5%. ................................. 45
Figure 24. Influence of the volume of GA added on concentration of fructose (g.L-1
) formed from inulin
hydrolysis at 65ºC. The assay was performed for PVA of 50 kDa with 400 µL of GA. Bioconversion
was performed in 5% (w/v) inulin in acetate buffer at pH 4.5. 100 mg of immobilized inulinase was
used as biocatalyst. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the beginning
of incubation. Standard deviation did not exceed 5%. .......................................................................... 46
Figure 25. Representative images of chitosan beads. No formation of beads through the original
procedure (A); Beads formed with chitosan solutions of 35 g.L-1
(B), 48.3 g.L-1
(C) and 68.3 g.L-1
(D).
After beads formation they were divided into three portions, to: stabilization in acetate buffer,
dehydration or reinforcement with GA. .................................................................................................. 47
Figure 26. Influence of different treatments on the concentration of fructose (g.L-1
) formed from inulin
hydrolysis. Each assay was performed at different temperatures 50ºC, 55ºC, 60ºC and 65ºC. Chitosan
beads stabilized in acetate buffer (absence), dehydrated in desiccator during 15 minutes or reinforced
with 500 µL of glutaraldehyde during 1 hour were studied. Bioconversion runs were performed in 5%
(w/v) inulin in acetate buffer at pH 4.5. 100 mg of immobilized inulinase was used as biocatalyst.
Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the beginning of incubation.
Standard deviation did not exceed 5%. ................................................................................................. 48
Figure 27. Studies performed on PVA film immobilized inulinase and CS beads immobilized inulinase.
............................................................................................................................................................... 50
Figure 28. Coomassie Blue stained SDS-PAGE gel of inulinase samples. Lane M - Precision Plus
Protein™ Dual Color Standard. Lanes 1 and 2 are duplicates of inulinase samples from Fructanase
Mixture. Lanes 3 and 4 are duplicates of inulinase 10-fold diluted in acetate buffer 100 mM pH 4.5. . 51
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Figure 29. pH profile of free inulinase and inulinase immobilized in CS beads and PVA film.
Bioconversion runs were performed in 5% (w/v) inulin solution at 50ºC at different pH values (3.6 –
5.5). 60 mg of immobilized inulinase in CS beads, 100 mg of immobilized inulinase in PVA film or 50
µL of a 100–fold diluted preparation of free inulinase were used as biocatalyst. ................................. 52
Figure 30. Temperature profile of free inulinase and inulinase immobilized in CS beads and PVA film.
Bioconversion runs were performed in 5% (w/v) inulin solution at pH 4.5 at different temperatures
(45ºC – 70ºC). 60 mg of immobilized inulinase in CS beads, 100 mg of immobilized inulinase in PVA
film or 50 µL of a 100–fold diluted preparation of free inulinase were used as biocatalyst. .................. 53
Figure 31. Influence of matrix weight on activity of immobilized enzyme, either in PVA film or CS
beads. Bioconversion runs were performed in 5% (w/v) inulin solution at pH 5.0 at 55ºC and 65ºC for
PVA and CS immobilization, respectively. Different matrix weight (40, 80, 120 and 200 mg) from both
types of immobilization were used as biocatalyst.................................................................................. 54
Figure 32. Michaelis-Menten kinetics of inulin hydrolysis expressed by free and immobilized inulinase
in PVA and CS matrices. Bioconversion reactions were carried out with concentrations of inulin
ranging from 15 g.L-1
to 150 g.L-1
in acetate buffer 100 mM pH 4.5 (for the free enzyme) or pH 5.0 (for
the immobilized enzyme) at optimum reaction conditions. .................................................................... 55
Figure 33. Thermal deactivation profile for free inulinase at 40ºC, 50ºC, 60ºC and 70ºC incubation
temperatures. The lines represent the trend of the three-parameter bi-exponential model.
Bioconversion runs for initial activity were performed in 5% (w/v) inulin solution 100 mM pH 4.5 with 50
µL of 100-fold diluted enzyme solution in acetate buffer 100 mM pH 4.5. ............................................ 58
Figure 34. Thermal deactivation profile for immobilized inulinase in CS beads (A) or in PVA film (B) at
40ºC, 50ºC, 60ºC and 70ºC incubation temperatures. The lines represent the trend of the three-
parameter bi-exponential model. Bioconversion runs for initial activity were performed in 5% (w/v)
inulin solution 100 mM pH 5.0 at optimum conditions. 60 mg of CS beads or 100 mg of PVA film were
used as biocatalyst. ............................................................................................................................... 59
Figure 35. Storage deactivation profile predicted through three-parameter bi-exponential model for
storage stability of free and immobilized biocatalyst. The lines represent the trend of the three-
parameter bi-exponential model. Bioconversion runs were performed with 50 µL of 100-fold enzyme
preparation in acetate buffer 100 mM pH 4.5, 60 mg of CS beads or 100 mg of PVA film. The
reactions were performed in 5% (v/w) inulin solution 100 mM at optimum conditions.......................... 61
Figure 36. Reactor filled with CS beads used for continuous-flow operation (A). Appearance of the
beads after continuous operation (B). ................................................................................................... 63
Figure 37. Reactor filled with PVA film used for continuous-flow operation (A). Appearance of the film
used after continuous operation. Note that the film was cut into cubes before implemented in the
reactor (B). ............................................................................................................................................. 64
Figure 38. Operational stability at the packed bed reactor for inulin hydrolysis, based on the relative
product yield. The 1.206 g of immobilized inulinase in CS were used for the hydrolysis of 5% (w/v)
inulin solution in acetate buffer 100 mM pH 5.0 at 55°C. At day one the initial concentration of fructose
was 55.5 (± 0.37) g.L-1
, with volumetric productivity of 22.6 g.L-1
.h-1
. Moreover, no deactivation model
fit to the experimental data. ................................................................................................................... 65
Figure 39. Operational stability at the packed bed reactor for inulin hydrolysis, based on the relative
product yield. Predicted values (red line) were estimated assuming a linear inverted model. The 3.450
g of inulinase in immobilized in PVA were used for the hydrolysis of 5% (w/v) inulin solution in acetate
xiv
buffer 100 mM pH 5.0 at 50°C. At day one the initial concentration of fructose was 50 ± 2.5 g.L-1
, with
volumetric productivity of 18.3 g.L-1
.h-1
. ................................................................................................. 65
Figure A1. Schematic representation of the reaction between DNS reagent and reducing sugar. In the
beginning of the assay the DNS reagent exhibits a yellow color, which shifts to orange–red if the
reaction occurs. ..................................................................................................................................... 81
Figure A2. DNS calibration curve for fructose concentrations ranging from 0 to 5 g.L-1
. ..................... 81
Figure A3. HPLC calibration curve for fructose concentrations ranging from 0 to 60 g.L-1
. ................. 82
Figure A4. Calibration curve used for total protein quantification, obtained from BSA standards with
concentrations ranging from 2.5 to 20 µg.mL-1
...................................................................................... 83
Figure A5. Flow rate (mL.min-1
) calibration curve for rotations per minute (rpm) ranging from 0.5 to
10 rpm. Calibration was performed with distilled water. ........................................................................ 83
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List of tables
Table 1. Advantages and drawbacks of enzymes as catalysts. Adapted from [9] .................................. 4
Table 2. Examples of immobilized enzymes with enhanced activity. ..................................................... 5
Table 3. Main characteristics of the five basic immobilization methods [9]. ........................................... 8
Table 4. Industrial applications of immobilized enzymes. ....................................................................... 9
Table 5. Examples of PVA based hydrogels for enzyme immobilization .............................................. 18
Table 6. Examples of CS hydrogels for enzyme immobilization ........................................................... 21
Table 7. Studies of inulinase immobilization in different matrix composition ........................................ 25
Table 8. Classification of enzyme reactors. Adapted from [196]. ......................................................... 25
Table 9. Effect of different PVA molecular weight and volume of cross-linker (GA) on film formation . 41
Table 10. Immobilization of inulinase in PVA and CS based matrices. The immobilization parameters
were determined as described in Methods section. .............................................................................. 50
Table 11. Kinetics constants obtained for inulin hydrolysis with free and immobilized inulinase. Data
were processed through Hyper32 software. ........................................................................................ 56
Table 12. Deactivation models and half-time equations. Where K(t) stands for activity at a given time,
Ki is the initial activity, Kd is the deactivation rate constant and t1/2 is the half time, A stands for a
complex function of individual rate constants, and α and β are apparent first-order rate constants.
Adapted from [207][15] .......................................................................................................................... 58
Table 13. Calculated parameters for the thermal deactivation of free and immobilized inulinase using a
three-parameter bi-exponential model. ................................................................................................. 60
Table 14. Rate constants for immobilized inulinase in both forms and free enzyme for storage
deactivation using a three-parameter bi-exponential equation. ............................................................ 61
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List of abbreviations
CS
CSTR
DD
DH
DNS
DP
FOS
GA
HFCS
HPLC
KM
MW
PBR
PFR
PVA
rpm
SDS
STR
TPP
v/v
Vmax
w/v
Chitosan
Continuous stirred tank reactor
Degree of deacetylation
Degree of hydrolysis
3,5-dinitrosalicylic acid
Degree of polymerization
Fructooligosaccharides
Glutaraldehyde
High fructose corn syrup
High performance liquid chromatography
Michaelis constant
Molecular weight
Packed bed reactor
Plug flow reactor
Polyvinyl alcohol
Rotations per minute
Sodium dodecyl sulfate–polyacrylamide gel electrophoresis
Stirred tank reactor
Tripolyphosphate
Volume per volume
Maximum reaction rate
Weight per volume
xviii
1
Thesis Outline
This dissertation is organized in five chapters.
The first chapter offers an overview on the increasing relevance of enzymes in biocatalysis,
followed by some insight into enzyme catalytic properties, advantages and constrains inherent to its
use, together with enzyme immobilization techniques. Special attention is given to hydrogels, which
are polymeric matrices used to entrap enzymes. A subchapter is focused on the description of
hydrogels, namely their characteristics and classification, their applications and the methods to
produce them. It is also important to have some insights about the protein model system (inulinase), in
particular the hydrolytic reaction, the substrate and the product. Examples of inulinase immobilization
into hydrogels are also mentioned. Finally, the main characteristics of the different types of bioreactors
are pointed out.
The second chapter comprises the motivation and the main goal of the master’s project.
The third chapter contains all the materials and methods used during the fulfillment of this work.
The fourth chapter describes the results attained with this study and the subsequently
discussion and comparison with the current knowledge. This chapter is divided into two subchapters.
In the first one the influence of some parameters on immobilized enzyme was tested, as follows: i)
different polymer molecular weights, ii) volumes of cross-linker, iii) temperatures and treatments
applied to the matrices. The second group of results comprises the characterization of free an
immobilized enzyme in terms of pH and temperature profiles, kinetic parameters, thermal and storage
stabilities and operational stability through the continuous production of fructose.
In the fifth and last chapter, final remarks considering the work developed and future
perspectives are made, together with references to what contributions this work offered in the field of
catalytic hydrogels.
2
3
Chapter I. Introduction
I.1. Enzymes as biocatalysts
Enzymes are the catalysts responsible for cell metabolism and/or substrate conversion in
various chemical reactions. Each reaction taking place in the cell is catalyzed by its own particular
enzyme, so that in a given cell there are a large number of enzymes. In the absence of enzymes most
of the reactions of cellular metabolism would not occur even over a time period of years. So far, more
than 3,000 enzymes have been identified with specific functions. Due to advances in the areas of
genomic & proteomic research, their number is further expected to increase [1].
I.1.1. History and importance of enzymes
The word enzyme is derived from the Greek meaning “in yeast” and was first used by Kühne in
1878. At this time it was used to distinguish between what were referred to as “organized ferments”
(meaning whole organism) and “unorganized ferments” (meaning extracts or secrets from whole
organism). The nomenclature of enzymes began in 1883, when Duclaux named enzymes by adding
the suffix – ase in the name of the substrate catalyzed, substance upon enzyme acts [2]. According to
the International Union of Biochemistry and Molecular Biology (IUBMB) nomenclature system, all
enzymes are classified into six classes, as follows: oxidoreductases, transferases, ligases, hydrolases,
isomerases and lyases. In 1894, Emil Fisher proposed the “lock-and-key” model for enzymatic
reaction, according to which a substrate molecule fits into the pocket of an enzyme. In a further
modification of this concept, Linus Pauling cited that a reaction is accelerated if a catalyst stabilizes
the transition state. The transition state stabilization concept is still used nowadays. From this time
ahead the biocatalysis basis was launched allowing a new field to develop and emerge from the
others at the time [3].
Enzymes can be produced from any living organism, either by extracting them from their
harboring cells or by recovering from cell exudate. In 1960, about 70% of the enzymes were extracted
from plant tissues or exudates and animal organs. Twenty years later the situation had reversed and
most industrial enzymes were produced from microbial sources, almost 90% of the total market [4].
The first application of cell free enzymes was the use of rennin aspartic protease isolated from
calf or lamb stomach in cheese making. In that time, enzymes were mainly used in the production of
food products (cheese, sourdough, beer, wine and vinegar) and in the manufacture of commodities
(leather, indigo and linen). In 1914, Röhn prepared the first commercial enzyme preparation (trypsin),
isolated from animals and used in the detergent to degrade proteins [5]. Since then, enzymes are
exploited in innumerous fields viz. paper, textile, food, detergent and pharmaceutical industries, as
well as in bioremediation, (bio)medical applications and wastewater treatment [6].
4
I.1.2. Enzyme properties - advantages and constraints
Enzymes form an abundant class of very effective and precise (bio)catalysts owing to their
excellent functional properties. Similar to other catalysts, enzymes are characterized by two
fundamental properties. First, they increase the rate of chemical reactions without themselves being
consumed or permanently altered by the reaction. Second, they increase reaction rates without
altering the chemical equilibrium between reactants and products [7].
Potentials and drawbacks of enzymes as catalysts are summarized in Table 1.
Table 1. Advantages and drawbacks of enzymes as catalysts. Adapted from [8]
Advantages Drawbacks
Generally more efficient (lower concentration of enzyme needed); Can be modified to increase selectivity, stability, and activity; High specificity; Milder reaction conditions (pH range of 5 to 8 and temperature range of 20 to 40ºC); Environment friendly (completely degraded in the environment); Generally considered as natural products.
Susceptible to substrate or product inhibition; Solvent usually water (high boiling point and heat of vaporization); Enzymes found in nature in only one enantiomeric form; Limiting operating region (enzymes typically denatured at high temperature and pH); Enzymes can cause allergic reactions; High production costs.
Often, for a given biocatalytic process, the soluble enzyme does not meet the requirements for
large-scale application, and its properties thus need to be optimized or modulated. Numerous
examples of improved enzyme properties, based on recombinant DNA technology, high throughput
technology and protein engineering (through directed and particularly through random mutagenesis,
namely using directed evolution), have been reported [9]–[11]. Additionally, enzyme efficiency can be
improved by optimization of the reaction conditions, viz. pH, temperature, substrate concentration [12].
Beside the drawbacks described in Table 1, enzymes are complex molecular structures that are
intrinsically labile and in some cases costly to produce. Additionally, enzyme biocatalysis is often
hampered by a lack of long-term operational stability and difficult recovery and re-use of the enzyme
[13]. In order to overcome these drawbacks, enzyme immobilization has been emerging as a viable
and reliable solution.
I.1.3. Immobilization methods
The term “immobilized enzymes” refers to enzymes physically confined or localized in a certain
defined region of space with retention of their catalytic activities, and which can be used repeatedly
and continuously [14]. Immobilized enzymes are being used since 1916, when Nelson and Griffin
discovered that invertase when absorbed to charcoal has the ability to hydrolyze sucrose [15]. The
first industrial application of an immobilized enzyme is the Tanabe process for the production of
L-amino acids by resolution of racemic acylamino acids using an aminoacylase from Aspergillus
oryzae [16].
5
Immobilized biocatalysts allow the possibility to work in repeated batch mode or under
continuous mode of operation, the integration in cascade of several catalytic steps and easy
separation of enzyme from the product (determinant factors in food and pharmaceutical industries).
Enzyme recycling is the best option that immobilized enzymes can provide, reducing the costs of
operation and increasing, at the same time, the catalyst productivity (kg product/kg enzyme). If
properly design, immobilization can contribute to improve key enzyme properties viz. activity,
selectivity, substrate specificity, stability and decrease inhibition (examples of enhanced enzyme
activity are given in Table 2) [13], [17].
Stability is one of the most important features of an immobilized enzyme used in a large scale
operation. Most enzymes when immobilized show a higher stability against pH, temperatures,
contaminants, shear stress or different organic solvents [18]. Different types of stability are
distinguished, such as operational, storage and thermal. Among them, operational is the most relevant
engineering parameter.
Table 2. Examples of immobilized enzymes with enhanced activity.
Enzyme Applications Kinetic parameters References
Keratinase Synthesis of keratin Immobilized: specific activity = 129.0 U·mg
−1;
Soluble: specific activity = 37 U·mg−1
[19]
Laccase Textile wastewater
treatment
Immobilized: KM = 0.0717 mM, Vmax = 0.247 mM·min−1
;
Soluble: KM = 0.0044 mM, Vmax = 0.024 mM·min−1
[20]
Glucose
oxidase
Estimation of
glucose level
Immobilized: KM = 2.7 mM, Vmax = 28.6 U·μg−1
;
Soluble: KM = 9 mM, Vmax = 6.2 μmol·min−1
mg−1
[21]
Cellobiase Bioethanol
production
Immobilized: KM = 0.30 mM, Vmax = 6.77 μM·min−1
;
Soluble: KM = 2.48 mM, Vmax = 2.38 μM·min−1
[22]
* U stands for the amount of protein needed to convert 1 µmol of substrate in 1 minute
Nonetheless, the performance of enzyme immobilization may be affected by conformational
effects, partition effects and mass transfer limitations [23].
Conformational effects refer to the structural changes produced in the enzyme molecule as a
consequence of the immobilization procedure. Alteration of the native three-dimensional structure of
the enzyme protein and steric effects due to its close proximity to the surface of the matrix are
conformational changes that may produce differences in kinetic behavior with respect to the free
enzyme [4]. The most common partition effects are electrostatic interactions between charged species
and fixed charges in the carrier that typically lead to a shift in the pH-activity profile; or when the carrier
has a hydrophobic/hydrophilic nature that differs from that of the bulk medium. In some cases, the
effect can be used advantageously. Regarding mass transfer limitations, these are often termed as
diffusional restrictions since the transport of the substrate from the bulk medium to the
microenvironment of the biocatalyst and the transport of the product formed back to the bulk medium
is ultimately limited by molecular diffusion (Figure 1).
6
Due to internal structural changes and restricted access to the active site, kinetic constants (e.g.
KM, Vmax) of enzymes may be altered. This phenomenon has been observed in several studies.
Figure 1. Formation of concentration gradients in porous enzyme carriers. The analyte’s (substrate or product)
concentration in the well-mixed liquid bulk (Cbulk) often differs from the concentration inside the carrier (Cin). For
substrates, Cbulk may be higher than Cin, whereas for products, the situation may be reversed. From [24].
More than one hundred techniques for immobilizing enzymes have been developed, which can
be divided into five principal methods, as follows: immobilization by covalent binding and non-covalent
binding (which includes physical adsorption, ionic binding, affinity binding and hydrophobic binding),
immobilization by entrapment, immobilization by encapsulation and immobilization by cross-linking
[25], [26]. Table 3 summarizes the main characteristics of each technique.
Despite the huge variety of immobilization techniques, there is no general universally
immobilization method and no method is perfect for all purposes [8]. Adsorption of the enzyme onto a
surface is the easiest and the oldest method of immobilization. Entrapment and cross-linking tend to
be more laborious enzyme fixation methods, but they do not require altering the enzyme as much as
other techniques. The formation of the covalent linkage often requires harsh conditions, which can
result in a loss of activity because of conformational changes of the enzyme.
Immobilization by entrapment, using polymeric gels, has been gained a huge importance.
Hydrogels are being used for enzyme and cell immobilization, due to their biocompatibility, low
coefficient of friction and high water content [27]. Further details regarding hydrogels will be discussed
in Chapter I.2.
More recently new types of enzyme immobilization have been emerging, viz. single-enzyme
nanoparticles (SENs), enzymatic immobilization of enzymes, microwave irradiation technique and
photoimmobilization [28]. Also, nanobiocatalysis has emerged as a rapidly growing area, since
nanomaterials (nanofibers, nanotubes and nanoparticles) have manifested great efficiency in the
manipulation of the nanoscale environment of the enzyme [29]. Also, they can maximize surface area
(overcoming mass transfer limitations) and improve the enzyme activity relative to the classical matrix
structure [30].
7
I.1.4. Immobilization matrices
The characteristics of the matrix will be of major importance for the performance of the
immobilized systems and will determine the type of reactor used under technical conditions (i.e.,
stirred tank, tubular reactors with either fluidized or fixed beds). The most important factors to take into
account in order to choose the best matrix for immobilization are the existence of functional reactive
groups, the definition of hydrophobic or hydrophilic character, the chemical, mechanical or biological
resistance, the geometry and the cost or availability.
Regarding the shape of the matrix, it can be classified into two types, i.e. irregular and regular
shapes such as: beads, fibers, hollow spheres, thin films, discs or membranes.
Matrices can be classified as inorganic (natural mineral or processed materials) and organic
(natural or synthetic polymers) according to their chemical composition (Figure 2) and, as porous and
not porous according to their morphology. This last parameter gives an idea of the available area for
binding the biocatalyst, which determines the amount of enzyme that can bind to the matrix [8]. The
organic matrices are preferable because of its adaptability to different methods of immobilization. Its
main disadvantage is the relatively low chemical and mechanical resistance, which impair its use in
aggressive environments and limits the possibilities for regeneration. The synthetic polymers have the
advantage of being inert to microbial attack, allowing the variation of the degree of porosity [14]. By
contrast, the inorganic matrices have the advantage of being chemically inert and having a good
mechanical and thermal stability, relatively to organics. However, the functional groups are almost
always hydroxyl groups, which limit the types of binding [14].
Selection of the immobilization technique and matrix is largely dependent on the peculiarity of
certain applications, the enzyme and the economic features. The design of new protocols which fulfils
the requirements of industry is still an exciting goal. Moreover, bearing in mind that the final use will be
as industrial catalyst, the ideal immobilization processes should limit the use of toxic or highly unstable
reagents, be very simple, robust and cost-effective [8].
Figure 2. Materials used for fabrication of immobilization matrices. Adapted from [26].
Organic Matrices
Natural polymers
Alginate, chitosan, chitin, gelatin, cellulose, starch, pectin,collagen
Synthetic polymers
Epoxy resin, silicone, polyvinyl alcohol, nylon, polyurethane, polyester
Inorganic Matrices
Natural minerals
Silica, bentonite, charcoal, sand, diatomaceous earth
Processed materials
Glass (nonporous and controlled pore), metals, controlled pore metal oxides, zeolites, activated carbon
8
Table 3. Main characteristics of the five basic immobilization methods [8].
Principle Main characteristics
Covalent
immobilization
(Figure 3.A)
Formation of strong and stable interaction. Chemical binding between functional groups present on the surface of the matrix material (glass, polyacrylamide, cellulose, agarose) and those on the enzyme surface (amino or carboxyl groups). Advantages: i) no leakage of enzyme, ii) enzyme can be easily in contact to substrate and iii) increase of the thermal stability. Drawbacks: i) loss of enzyme activity, ii) steric effects and iii) good matrices are expensive. Immobilization in the presence of the substrate or competitive inhibitor of the enzyme, the use of a space-arm or/and a reversible immobilization trough disulfide bonds are possible solutions to overcome those drawbacks.
Non-covalent
immobilization
(Figure 3.B)
Involves hydrophobic and Van der Walls forces, hydrogen bonds, electrostatic and ionic binding. The procedure consists of mixing together the biological components and the matrix with suitable properties, under specific conditions. Advantages: i) reversibility and simplicity, ii) immobilization yields are high and no obnoxious reagents are involved and iii) cheap and quick method. Drawbacks: i) the enzyme can be easily desorbed from its matrix by subtle changes in the reaction medium, ii) contamination of product, iii) overloading on the matrix, iv) steric hindrance by the matrix and v) non-specific binding.
Immobilization by
entrapment
(Figure 3.C)
Consists in confining the enzyme within the inner cavities of a solid polymeric matrix compact enough to retain the enzyme molecules within it. Most popular matrices for entrapment are: alginate, polyacrylamide, polyurethane, polyvinyl alcohol and қ-carrageenan. Entrapment could be in film, fiber or capsules/beads. Advantages: i) enzyme loading is very high and ii) protection from direct contact with the environment thereby minimizing the effects of gas bubbles, mechanical sheer and adverse solvents. Drawbacks: ii) mass transfer limitations and ii) enzyme leakage from the matrix. This could be overcame by increasing gel strength which it turns magnifies mass transfer limitation; another approach involves the formation of cross-linked enzyme aggregates prior to entrapment.
Immobilization by
encapsulation
(Figure 3.D)
Is similar to entrapment in that the enzyme is free in solution, but restricted in space. Most popular membrane materials: nylon and cellulose nitrate. Advantages: i) the enzyme could be encapsulated inside the cell and ii) possibility of co-immobilization. Drawbacks: i) diffusion limitations and ii) membrane rupture.
Immobilization by
cross-linking
(Figure 3.E)
Comprises the reaction between NH2 groups of enzymes with bifunctional chemical cross-linker, in order to form a large and three-dimensional complex structure. The most use cross-linker is glutaraldehyde, since it is expensive and readily available in commercial quantities. There are three major types of cross-linking: CLEs (cross-linked solution enzyme), CLECs (cross-linked enzyme crystals) and CLEA (cross-linked enzyme aggregates). Advantages: i) matrix-free and ii) cross-linking between same enzyme molecules stabilizes the enzymes by improving the rigidity of the structure. Drawbacks: i) harshness of reagents and ii) possibility of crosslinking the active site of enzyme.
Figure 3. Representative images of each type of immobilization technique, enzyme is represented as blue circle: covalent immobilization (A), non-covalent immobilization by
physical adsorption (B), entrapment (C), encapsulation (D) and cross-linking (E).
A B C D E
9
I.1.5. Applications of immobilized enzymes and markets perspective
In spite of the long history and obvious advantages of enzyme immobilization Straathof et al.
(2002) estimated that only 20% of biocatalytic processes involve immobilized enzymes [31]. However,
over the last few years a number of interesting new developments have been reported in the literature
and patent applications, indicating that enzyme immobilization has entered an exciting new phase.
Currently, enzyme immobilization is widely exploited technique in various industries: food,
pharmaceutical, detergent, textile, fuel and chemicals (Table 4). Aside from these applications,
immobilized enzymes are presently used in the diagnosis and treatment of various diseases and in the
development of biosensors and analytical devices [32].
Table 4. Examples of industrial applications of immobilized enzymes.
Industry Enzyme Class Applications References
Detergent
Lipase Fatty acid and oily stain removal
[33] Cellulase Color clarification, cleaning
Peroxidase Dye removal
Fuel Lipase Biodiesel [FAME] production [34]
Textile Cellulase Scouring/biopolishing/desizing [35]
Food
Glucose isomerase Production of HFCS [36]
- Galactosidase Hydrolysis of lactose in dairy products [37]
Lipase Dairy, Baking, Fats/Oils [38]
Pharmaceuticals Penicillin acylase Synthesis of 6-APA for production of penicillin [39]
Chemicals Lipase Resolution of chiral alcohols and amines [40]
FAME: Fatty Acid Methyl Ester; HFCS: High Fructose Corn Syrup
According to Binod et al. (2013), the global market for industrial enzymes is estimated at 3.3
billion US dollar in 2010 and expected to reach 4.5 to 5 billion US dollar by 2015 at a compounded
annual growth rate of 7–9%. This market is dominated by products containing non-immobilized
enzymes, predominantly hydrolases (e.g. amylases, proteases, cellulases and lipases), which
represent 63% of the sales [5], [41]. Enzymes for use in non-industrial markets, primarily for
pharmaceutical, diagnostic and research applications, accounted for around 2.4 billion US dollar in
2010. Sales of enzymes for biocatalysis, many of which are used in immobilized form, were valued at
160 US dollar million in 2010 and projected to increase to 230 million US dollar by 2015 [42].
Glucose isomerase is one of the most commercialized enzyme (107 ton per year), which is
especially useful for the production of high fructose corn syrup. Since 1967, many methods for the
immobilization of glucose isomerase have been developed [43]. Alongside, and due to diverse
applications, the global market of immobilized lipase still increasing and represents a product scale of
1011
ton per year [42].
Major enzyme producers are located in Europe, USA and Japan. Nearly 75% of the total
enzymes are produced by three top enzyme companies, i.e. Denmark-based Novozymes, US-based
DuPontand and Switzerland-based Roche. The market is highly competitive, has small profit margins
and is technologically intensive [44].
10
I.2. Hydrogels
Hydrogels have been found in nature since life on Earth and play an important role in
entrapment. Many reports have been published about new chemical and physical structures,
properties and innovative restricted applications of polymer hydrogels, demonstrating their importance
beyond enzyme immobilization.
According to SciFinder®, the first reference of hydrogel was in 1894 to describe a colloidal gel of
inorganic salts. The first hydrogel polymer was synthesized in 1954, by Lim and Wichterle and it was
made by synthetic poly-2-hydroxyethyl methacrylate. The development of the first soft hydrogel
contact lenses by Wichterle in 1961 represented the first successful clinical application of hydrogel
polymers and remains one of the most important uses of hydrogels today. In the 1980s hydrogels
were modified for other applications. Lim and Sun (1980) obtained calcium alginate microcapsules for
cell encapsulation, and Yannas et al. (1989) incorporated natural polymers such as collagen and
shark cartilage into hydrogels for use as artificial burn dressings [45], [46].
I.2.1. From characteristics to applications
Hydrogel is a three-dimensional cross-linked hydrophilic polymer network capable of swelling
and de-swelling reversibly in water and retaining large volume of liquid in swollen state (Figure 4).
Swelling effect is quite a complicated process and consists of a number of steps [47].
In general, the amount of water is at least 20% of the total weight [48]. If water is composed of
more than 95% of the total weight, then the hydrogel is called superabsorbent [49].
The ability of hydrogels to absorb water arises from hydrophilic functional groups (NH2, COOH,
OH, CONH2, CONH and SO3H) attached to the polymeric backbone, while their resistance to
dissolution arises from crosslinks between network chains [50]. In the absence of crosslinking points,
the hydrophilic linear polymer chains dissolve in water due to the polymer chain and water
thermodynamic compatibility [50].
Figure 4. Representative image of a hydrogel. Adapted from [46].
The literature reports a number of classifications of hydrogels, and presents several views.
According to the source, hydrogels can be divided into those formed from natural polymers (i.e.
11
chitosan (CS), alginate, hyaluronan, starch) and those formed from synthetic polymers (i.e. polyvinyl
alcohol (PVA), polyvinylpyrrolidone, polyacrylamide) [51]. Synthetic polymers have been replaced
natural polymer hydrogels because of their purity, high absorption capacity, well-defined structure and
functionality, degradation and stability in varying ranges of pH, temperature and pressure [52]. One
polymer from each source (PVA and CS) will be discussed in Chapter I.2.3 and I.2.4.
On the basis of the cross-linking method, hydrogels can be divided into chemical, physical or
radiation cross-linking. Also, hydrogels are classified into conventional or smart hydrogels. Smart
hydrogels respond to environmental stimuli and experience unexpected changes in their growth
actions, network structure, mechanical strength and permeability, hence called environmentally
sensitive. Physical stimuli include temperature, electric or magnetic field, light, pressure and sound,
while the chemical stimuli include pH, solvent composition, ionic strength and molecular species [53].
A biochemical stimulus involves the responses to ligand, enzyme, antigen and other biochemical
agents [54]. Figure 5 represents the different classifications for hydrogels.
Figure 5. Classification of hydrogels based on the different properties. Adapted from [51].
There are several methods to characterize hydrogels. An easy way to quantify the presence of
hydrogel in a system is to disperse the polymer in water using a cylindrical vial and visually observe
the formation of insoluble material. Morphology, particle size, cell toxicity are some of the features to
characterize hydrogels. In order to achieve this characterization some methods can be used such as:
solubility identification, swelling measurement, Fourier Transform Infrared Spectroscopy, Scanning
Electron Microscopy, Light Scattering, sol-gel analysis or Nuclear Magnetic Resonance [50].
Besides water holding capacity and permeability, biocompatibility is the third most important
characteristic property. Generally, hydrogels possess a good biocompatibility since their hydrophilic
surface has a low interfacial free energy when in contact with body fluids, which results in a low
tendency for proteins and cells to adhere to these surfaces (antifouling property) [50], [53].
The cross-links between the different polymer chains results in viscoelastic and sometimes pure
elastic behavior and gives a gel its structure (hardness), elasticity and contribute to stickiness [50].
HYDROGELS
- Natural - Synthetic - Hybrid
Source
Degradability
- Biodegradable - Non-degradable
Polymeric composition
Polymeric configuration
Physical properties
Ionic charge
-Homopolymeric hydrogels -Copolymeric hydrogels -Multipolymer interpenetrating polymer (IPN)
-Crystalline -Amorphous (non-crystalline) -Semicrystalline
-Conventional -Smart
Appearance Cross-linking
-Nonionic -Ionic -Amphoteric -Zwitterionic
-Fibers -Films -Beads/capsules
-Chemical -Physical -Radiation
12
Furthermore, hydrogels, due to their significant water content possess a degree of flexibility similar to
natural tissue and low coefficient of friction. It is possible to change the chemistry of the hydrogel by
controlling their polarity, surface properties, mechanical properties and swelling behavior [50].
Due to these wonderful properties, hydrogels provide a platform for a range of applications.
Park and Park (1996) have reviewed the potential of hydrogels in bioapplications [48]. Regenerative
medicine and tissue engineering [48], [53], drug delivery [36],[42],[43], biosensors [57], [58], cancer
therapy [59], uterine artery embolization procedure [60], embolization of spinal tumors [61] and
pharmaceuticals [62] are some of the bioapplications reported. Also, hydrogels may be applied to the
development of protein microarrays and microfluidics [63], [64], food additives [65], agriculture as a
fertilizer release system to encourage the soil [66], wound dressing [67], cosmetic industry [68] and
wastewater treatment [69].
In addition, hydrogels are known to be suitable materials for enzyme immobilization. The
successfully immobilization of enzymes in hydrogels are now well documented. The fact that such gels
with a water content of about 96% provide a microenvironment for the immobilized enzyme close to
that of the soluble enzyme with minimal diffusion restrictions is a significant advantage. However,
every coin has two sides. Hydrogels are impaired by low mechanical strength/stability (particularly
those from natural polymers) and enzyme leakage trough the pores of the gel lattice (Table 3) [70].
I.2.2. Technologies adopted in hydrogel preparation
Polymer chains per se are not useful in the majority of applications, since they do not have
suitable properties; therefore the production of hydrogels is required.
The main steps of production of hydrogels are represented in Figure 6. If synthetic polymers are
used, a polymerization step is usually necessary in early stages. Depending on the type of polymer
could be necessary to perform a chemical treatment in order to have proper functional groups. Then,
the cross-linking between polymer chains and the solidification process are performed. Last process
includes a drying process or addition of solvents or salts, which will promote the solidification. Each
step should be chosen depending on the type of polymer, the immobilized enzyme, the matrix
appearance, the application intended, among others.
Figure 6. General hydrogel preparation diagram.
Regarding cross-linking, it could be physical cross-linking, chemical cross-linking and radiation
cross-linking. While physical hydrogels are reversible due to the conformational changes, chemical
Polimerization
Chemical treatment
Cross-linking process
Solidification process
13
hydrogels are irreversible [50]. Hence, the physical cross-linking is not as strong or as stable as the
chemical one.
Radiation cross-linking is widely used technique since it does not involve the use of chemical
additives and therefore retaining the biocompatibility of the biopolymer. However, it requires specific
equipment [71]. The technique mainly relies on producing free radicals in the polymer following the
exposure to the high energy source such as gamma ray, x-ray or electron beam. The action of
radiation (direct or indirect) will depend on the polymer environment (i.e. dilute solution, concentrated
solution, solid state) [72].
Chemical cross-linking can be achieved through the reaction of polymer functional groups (such
as OH, COOH, and NH2) with cross-linkers as aldehydes (e.g. glutaraldehyde, boric acids, boronic
acids) [50]. Glutaraldehyde (GA) has been used more frequently as a cross-linking agent than any
other reagent, as it is less expensive, readily available, and highly soluble in aqueous solution [73].
The high reactivity of the aldehyde groups, which readily form imine bonds (Schiff’s base) with amino
groups and acetal bonds with hydroxyl groups provides the efficiency of GA on the cross-linking of
several polymers [74]. Due to the proved toxicity of these compounds, physical cross-linking is
preferable for pharmaceutical, medical or food applications. However, GA remains an important
component of current commercial glucose isomerase products including Sweetzymetm
and
GenSweet IGI, marketed by Novozymes A/S and Genencor (now DuPont Industrial Biosciences),
respectively [42].
Physical cross-linking of polymer chains can be achieved without the need for chemical
modification or the addition of cross-linking entities. This method uses a variety of environmental
triggers as pH, temperature or ionic strength and, a variety of physicochemical interactions, as follows:
hydrogen bonding, ionic interaction or Van der Walls interactions [75].
I.2.3. Polyvinyl alcohol (PVA) based hydrogels
Polyvinyl alcohol (PVA) is a biodegradable synthetic polymer, odorless and tasteless,
translucent, white or cream colored granular powder. PVA was first prepared by Haehnel and
Herrmann in 1924 by hydrolyzing polyvinyl acetate in ethanol with potassium hydroxide. The first
scientific reports on PVA were published in 1927. PVA is the largest volume synthetic resin produced
in the world [76].
I.2.3.1. Structure and properties of PVA
PVA is a linear polymer which has a relatively simple chemical structure with a pendant
hydroxyl group. PVA is not prepared by polymerization of the corresponding monomer (vinyl alcohol),
14
since it does not exist in a stable form (occurring rearrangements with its tautomer). Instead, PVA is
commercially produced through the polymerization of vinyl acetate to polyvinyl acetate (PVAc),
followed by partial or complete hydrolysis of PVAc to PVA, removing acetate groups (Figure 7) [71].
Figure 7. Schematic of the reaction of the PVAc to PVA. In red are the acetate groups whereas in green are the
hydroxyl groups.
The polymerization step controls the length (called degree of polymerization) of the vinyl acetate
molecular chains, and therefore the molecular weight. The PVA membrane strength, aqueous solution
viscosity, and other properties vary greatly depending on the degree of polymerization. It is also
possible to produce PVA with different characteristics by copolymerizing other monomers during this
process [71].
Whereas the hydrolysis step controls the amount of hydroxyl groups, which determine the
degree of hydrolysis (Figure 8) [71]. These hydroxyl groups are prompt to suffer simple modifications,
which is an advantage for hydrogel production. The degree of polymerization (DP) and the degree of
hydrolysis (DH) determine physical, chemical and mechanical properties of PVA (Figure 9) [76]–[78].
Commercial PVA grades are available with high degrees of hydrolysis (above 98.5%) [71].
Figure 8. Degree of hydrolysis (DH) and polymerization (DP) of PVA.
From http://www.j-vp.co.jp/english/product/pva/index.html.
Solubility is one of the properties which will be pointed out. PVA is only soluble in highly polar
and hydrophilic solvents, such as water, Dimethyl Sulfoxide (DMSO), Ethylene Glycol (EG), and
N-Methyl Pyrrolidone (NMP) and insoluble in organic solvents. The solubility in water is a function of
the DP and DH, and solution temperature [78]. Fully hydrolyzed PVA is only completely soluble in hot
+ Methanol
Vinyl acetate monomers
Polyvinyl acetate (PVAc)
Polymerization
Hydrolysis
Polyvinyl alcohol (PVA)
+ Sodium hydroxide
Degree of polymerization = n + m
Degree of hydrolysis = n/(n+m) × 100
15
to boiling water, since hydroxyl groups in PVA contribute to strong hydrogen bonding both intra- and
intermolecularly, which reduces solubility in water. The presence of residual acetate groups in partially
hydrolyzed PVA weakens these hydrogen bonds and allows solubility at lower temperatures. In
general, partial hydrolysis lower melting points, lower strength and lower water dissolution
temperatures and, are easier to process than those based on fully hydrolyzed polymers [76].
Figure 9. Effect of molecular weight and % of hydrolysis in the properties of PVA. The variations in properties with
molecular weight are for a constant degree of hydrolysis (mol %), and the effect of hydrolysis is at a constant
molecular weight. From [76].
I.2.3.2. Synthesis and properties of PVA hydrogels
Due to its hydrophilicity, PVA must be cross-linking in order to be useful for a wide variety of
applications, specifically in medicine and pharmaceutical sciences. In presence of cross-links it
remains insoluble in solution, forming hydrogels. PVA crosslinking may be achieved by chemical,
radiation or physical mechanisms. Some examples of how these mechanisms influence enzyme
immobilization will be given.
PVA hydrogels obtained by radiation cross-linking have already been documented. Jha and
D’Souza (2005) prepared a polymer membrane of polyvinyl alcohol-polyacrylamide by γ-irradiation for
entrapment of urease [79]. Bourke et al. (2003) obtained PVA hydrogels designed for wound-healing
applications by ultraviolet photo-cross-linking of acrylamide-functionalized nondegradable PVA [80].
Nikolic et al. (2007) obtained PVA hydrogels membranes by gamma irradiation for hydrogen fuel
cells [81].
Concerning chemical cross-linking, there are a wide variety of cross-linking agents for PVA such
as glutaraldehyde (GA), acetaldehyde, formaldehyde, maleic acid and boric acid [82] [83]. Combining
the use of one of these cross-linking agents with sulfuric, acetic acid or with methanol, leads to the
16
formation of acetal bridges between the pendant hydroxyl groups of the PVA chains [84]. The reaction
modes can be adjusted by specific cross-linker agents and/or operational conditions, also the resulting
cross-linked matrix is readily permeable to both water and salts [85].
In present review only cross-linking with glutaraldehyde will be discussed. GA has gained
increasing attention as a PVA cross-linking agent because of the absence of thermal treatment
needed to drive the reaction. Also, is considered a bifunctional cross-linker, because it has two active
sites, thus it can bind molecules of PVA together. The main drawback of this reaction is the acidic
environment required, which could lead to the enzyme denaturation [85]. Moreover, GA is known to
bind nonspecifically to biomolecules, such as proteins, so that it can bind such biomolecules and PVA
together. The mechanism proposed for the reaction indicates that there is an optimum ratio between
the reactants, PVA and GA, that favors the crosslinking of the polymer; an excess of the aldehyde can
lead to the branching of PVA [86], as illustrated in Figure 10.
Figure 10. Mechanism of the reaction between PVA and GA. From [85].
Although the reaction is typically favored in an acidic environment, Figueiredo et al. (2008)
performed the crosslinking of PVA using GA in the absence of an acid catalyst and organic solvents.
In this study, membranes were prepared at 40ºC with a pH of 6.2 and a GA/PVA mass ratio of 0.01,
with the aim of the further entrapment of proteins in this matrix [85]. Additionally, Ca et al. (2001) show
that PVA hydrogels are not thermosensitive when is used ethylenediaminetetraacetic dianhydride
(EDTAD) as cross-linker [87]. Nunes et al. (2012) performed the immobilization of naringinase using
GA as cross-linker and dimethyl sulfoxide (DMSO), which decreased the toxicity of the immobilization
process and allowed the control of membrane porosity, respectively [70]. Mansur et al. (2008) studied
the influence of partially hydrolyzed PVA chemically crosslinked with GA for preparation pH sensitive
hydrogels [88].
The third mechanism of hydrogel preparation is physical cross-linking. PVAs are hydrophilic
polymers whose concentrated aqueous solutions are capable of gelling per se with the formation of a
non-covalent spatial network under prolonged storing. The gelation process could be done by
cryogenic gelation techniques or non-cryogenic gelation techniques [89].
17
Regarding cryogenic gelation techniques, this technique involves freezing and thawing cycles
and, consequently the formation of cryogels and small microcrystals. In cryoPVA gels, hydrogen
bonds are formed between hydroxyl groups of neighboring polymer chains (interpolymers) or between
hydroxyl groups in the same polymer chain (Figure 11) [75], [89]–[91]. In general, freeze-thawing
method is performed in water or in a dimethyl sulfoxide (DMSO)/water mixed solvent, where the
intermolecular bonds formed during the process act as efficient crosslinks. The use of DMSO reduces
the amount of ice formed and avoids phase separation, thus reducing porosity within the cryogel [89].
Figure 11. Network structure of PVA gels cross-linked by hydrogen bonds. From [92].
Concerning non-cryogenic gelation methods of PVA, a straightforward method to produce PVA
hydrogels was reported by Otsuka et al. [92]. The hydrogels were prepared by pouring a PVA solution
into a plastic dish and leaving the sample in a closed space at room temperature for 7 days during
which the water content decreased by 20 wt.-%. This incubation afforded formation of polymer intra-
and intermolecular hydrogen bonds and the resulting hydrogels remained stable. This method is
simple, safe, and inexpensive to obtain physical PVA gels, however is very time consuming when is
produced large sized hydrogels [89].
On the other hand, the gel formation by controlled drying at room temperature is already
performed by GeniaLab, a company which offers ready-to-use PVA solutions, known as
Lentikats [89]. Lentikats or lens-shape particles offer the following advantages: low costs for matrix
and production, easy preparation, excellent mechanical stability, easy separation from reaction
mixture and low diffusion limitation [93].
Several bioprocesses have been developed up to a commercial scale using Lentikats®. One of
the most successful involve the removal of nitrogen from wastewater, using immobilized Paracoccus
denitrificans in high concentrations (30 times higher biomass content in comparison to activated
sludge process), allowing the removal of nitrogen of up to 98% and reducing in 30% the operational
costs and energy savings [94]. Pharmaceutical and food industry also benefit from Lentikats®
technology in the production of fructose, through the sucrose hydrolysis using immobilized invertase
[95]. Ethanol production using immobilized Zymomonas mobilis is also one of the several applications
of Lentikats®
[96].
18
Although immobilization in Lentikats has already been successfully reported, enzyme
immobilization often fails due to the small size and the possibility to diffuse out of the gel matrix.
Therefore, some improvements to this methodology were performed. Gröger et al. (2001) reported no
catalyst leaching when enzymes are cross-linking with chitosan and glutaraldehyde, to obtain a supra
molecular structure prior to entrapment in Lentikats [97]. Fernandes et al. (2009) immobilized
inulinase in PVA particles using Lentikat liquid followed by extrusion directly into polyethylene glycol
(PEG) 600, which afforded an instantaneous gelation of PVA hemispheric particles, avoiding the
drying process [27]. Also, the use of a chemical cross-linking (such as GA) after the formation of PVA
lenses (Lentikats), leads to a supramolecular structure which provides more stability [98].
In addition, Lentikats exhibit physical instability at around 55ºC and above this temperature, the
PVA-based matrix lost their mechanical strength at these conditions and tended to melt [99]. The
temperature is a key parameter in immobilization; therefore an improvement in parameter is an open
field of research.
Immobilizations in lenticular shape and in microbeads are attempts to solve the problem of
mass transfer limitation, introducing new techniques that adjust the size and shape of PVA hydrogels.
Gong et al. (2010) demonstrated that the introduction of montmorillonite (MONT) and
dimethyldioctadecylammonium bromide (DDAB) into PVA matrices improve their limitation to organic
compounds containing hydrophobic groups [100].
Table 5 displays some examples of PVA hydrogels used for enzyme immobilization. Is
important to highlight that the majority of PVA based hydrogels are prepared with the commercial
preparation Lentikat liquid and the form of the matrix is mainly beads/capsules.
Applications of PVA hydrogels beyond enzyme immobilization will be discussed further.
Table 5. Examples of PVA based hydrogels for enzyme immobilization.
Enzyme Matrix and mode of preparation References
Lipase Freezing and thawing PVA aqueous solutions [101]
Invertase PVA-alginate solution was extruded into boric acid and calcium chloride
solution to form beads [102]
Invertase Lentikats capsules were formed and dried trough the standard method [95]
Lipase PVA particles were formed by crosslinking with boric acid [103]
Fructozyme L* Lentikat liquid was extruded into polyethylene glycol (PEG) [27]
Fructozyme L* Lentikat liquid was either partially dehydrated trough the standard method or
extruded into PEG [99]
Naringinase PVA particles were formed using GA as cross-linker and DMSO as co-solvent [70], [104]
ß-galactosidase Lentikat liquid was partially dried by standard method [105]
Fructozyme L Lentikat liquid was extruded into PEG and crosslinking with GA [98]
*commercial preparation of inulinase from Aspergillus niger and Bacillus stearothermophilus
19
I.2.4. Chitosan (CS) based hydrogels
CS is the second most abundant natural polysaccharide on Earth and it was discovered by C.
Rouget in 1859 when he treated chitin in boiling and concentrated potassium hydroxide. Rouget
named this structure “modified chitin”. This chitin-derivative was renamed chitosan by Hoppe-Seyler in
1894. Only in 1950 was the structure of CS finally resolved [106]. In 1934, both the production of CS
and the preparation of CS film and fibers were patented [107].
I.2.4.1. Structure and properties of CS
CS is a linear and natural polymer, which is composed of D-glucosamine and N-acetyl-
D-glucosamine monomers linked by β-glycosidic bonds. This polymer is obtained through the alkaline
N-deacetylation of chitin (Figure 12). Also, this polymer can be degraded by enzymes such as
lysozymes, some lipases and proteases. Additionally, it is insoluble in most solvents, which
determines its applications [108].
Figure 12. Reaction of N-deacetylation of chitin (A). The repeating structural unit of CS, showing the functional
groups at positions C-2, C-3 and C-6 (B). Adapted from [107].
As occur in PVA, the most important factor that determine physical, chemical and mechanical
properties of CS, and consequently the specific applications, is the degree of deacetylation (DD) [106].
The DD of the CS is more than 50%, which means that the majorities of the CS monomers is in the
deacetylated form and possess an amino group at the C-2 position. For commercial purposes the DD
is usually 70-95% [109].
The free amino can be chemically modified leading to the formation of diverse derivatives with
new bioactivity and biofunctionalities [110]. Additionally, amino groups make CS a cationic
polyelectrolyte (pKa ≈ 6.5), one of the few found in nature. This basicity gives CS singular properties:
CS is soluble in aqueous acidic media at pH < 6.5 and when dissolved possesses high positive charge
on NH3+ groups, it adheres to negatively charged surfaces, it aggregates with polyanionic compounds
A
B
20
and chelates heavy metal ions [106], [111]. These characteristics offer extraordinary potential in a
broad spectrum of CS applications. Furthermore, DD is an important factor on biocompatibility. It
appears that the higher DD is characteristic of a higher biocompatibility. Therefore, it is important to
choose CS with an appropriate DD for medical purpose [112].
I.2.4.2. Synthesis and properties of CS hydrogels
As PVA hydrogels, CS hydrogels can be produced mainly by chemical or physical crosslinking.
There are several chitosan chemical cross-linkers already described such as genipin [113], glyoxal
[114], epichlorohydrin [115] and ascorbic acid [116] and GA [117]. GA seems to be the most studied
cross-linker, in Figure 13A is represented the reaction between CS and GA.
Alternatively to covalent bonds, ionic agents promote ionic gelation and electrostatic interactions
between the positively charged chitosan chains and polyanions. Sodium tripolyphosphate (TPP) and
calcium phosphate are the most reported ones [118]. TPP have been employed in enzyme
immobilization matrices, since it reduces the gel pores size and is nontoxic [111]. The cross-linking
structure of the CS-TPP system is mainly determined by the reaction between the amino groups of CS
and TPP ions, and this reaction depends strongly on the associated pH (Figure 13B). Bhumkar and
Pokharkar (2006) described that in TPP at higher pH (pH 9) both the OH–
and phosphoric ions are
present, which compete with each other to interact with the –NH3+ sites of chitosan [119]. Alsarra et al.
(2004) studied the influence of the pH, TPP concentration and ionic strength of the gelling medium on
the entrapment efficiency, release, and activity of lipase in chitosan hydrogel beads [120].
Figure 13. Interactions of CS with GA (A) and TPP (B). Adapted from [117].
Table 6 summarizes some examples of CS hydrogels cross-linked by TPP and/or GA.
Additionally, several studies have been performed with a blend of polymers, one example are
chitosan-alginate hydrogels, which have been applied in diverse fields [121].
A B
21
Table 6. Examples of CS hydrogels for enzyme immobilization.
Enzyme Immobilization technique References
-galactosidase CS solution was extruded into TPP solution to form beads [122]
Porcine pepsin CS solution was extruded into a cross-linker solution and
reinforced with GA to form beads. [123]
α-amylase CS solution was cross-lined with GA to form microbeads [124]
Catalase CS solution was extruded into a cross-linker solution and
reinforced with GA to form beads. [125]
Tannase CS solution was extruded into TPP solution to form beads [126]
-L-Arabinofuranosidase CS solution cross-linked with GA to form films [127]
Lipase CS solution cross-linked with TPP to form beads [120]
I.2.5. Applications of PVA and CS hydrogels
PVA and CS based hydrogels, due to their unique physio-chemical and biological properties are
attractive materials for use in innumerous applications. Both polymers can be easily modified through
its free groups and they are commercially available in a wide range of molecular weights at low
price [128].
The respective hydrogels exhibit hydrophilicity, biodegradability, biocompatibility, bioadhesive
properties, non-toxicity and non-carcinogenic activity [129]. Nonetheless, PVA and CS hydrogels have
some distinctive characteristics. PVA synthetic gels exhibit a high degree of swelling in water, a
rubbery and elastic nature and excellent film-forming properties [129]. While CS natural gels display
antimicrobial activity and physiological inertness, remarkable affinity to proteins, hemostatic,
fungistatic, antitumoral and anticholesteremic properties [107]. Also, CS acts as a penetration
enhancer by opening epithelial tight-junctions [130].
The combination of CS and PVA should have beneficial effects on the biological characteristics
of blend hydrogels. Hence, by combining hydrophilic polymers of different sources, a class of hybrid
organic–organic network can be produced with properties not present in either one separately [131].
Furthermore, PVA and CS hydrogels are attractive materials for biocatalysis being used as
matrix networks for enzyme and cell immobilization. During this review several studies have been
pointed out, supporting this statement.
In addition, PVA and CS hydrogels have been employed in pharmaceutical and drug delivery
systems [132]–[134], tissue engineering [135]–[137], soil remediation [138], wastewater treatment
[139], food packaging [140] and cosmetic industry [141]. PVA gels have also been used for contact
lenses [142], in the lining for artificial hearts [143], tendon repair [144], artificial pancreas [145], topical
pharmaceutical and ophthalmic formulation [146], uterine embolization procedure [147]. Whereas CS
gels exhibit antitumor and anticoagulant activity [148], participate in delivery systems [149], artificial
cells and hemodialysis membranes [150].
22
I.3. Protein model system - Inulinase enzyme
I.3.1. Inulinase - general considerations
Microbial inulinases belong to an important class of industrial enzymes that have gained
increasing attention in the recent years, due to cost effective production and economically viable
process. Inulinases can be produced by plants and a host of microorganisms, including fungi, yeast,
and bacteria. Among them, however, Aspergillus sp. (filamentous fungus) and Kluyveromyces sp.
(diploid yeast) are apparently the preferred choices for commercial applications [151].
Inulinases belong to the glycoside hydrolase family 32 (GH32), promoting the hydrolysis of
O-glycosyl bonds, and have fructans as typical substrates, one example is inulin [152]. Microbial
inulinases are classified as endoinulinase and exoinulinase, depending on their mode of action. Few
inulinases have been their crystal structures available; one example is the exo-inulinase from
Aspergillus awamori (Figure 14) [152].
Figure 14. Representative crystal structure of an exo-inulinase from Aspergillus awamori. The structure is a
monomer with two domains and glycerol and N-acetyl-D-glucosamine are represented as ligands. PDB ID 1Y4W.
Endoinulinases (2,1--D-fructan fructanohydrolase, EC 3.2.1.7) are specific to inulin and
hydrolyze it by breaking the bonds between fructose units that are located away from the ends of the
polymer network to produce oligosaccharides, similar to fructooligosaccharides (FOS). This type of
reaction requires a strict control of reaction time to achieve the desired polymerization grade [152].
Exoinulinases (-D-fructan fructohydrolase, EC 3.2.1.80) split terminal fructose units from the
non-reducing end of the inulin molecule to liberate fructose (Figure 15). This reaction yields up to 95%
fructose. The property of having an exo or an endo action also depends on the microbial origin of the
enzyme [152].
Inulinases can be used in a wide range of industrial applications, for high fructose corn syrup
(HFCS) obtaining from inulin, bioethanol production, inulo-oligosaccharide production, single-cell oil
and single-cell protein production, some chemicals production like citric acid, butanediol, alcohols,
lactic acid etc. [153].
23
Figure 15. Full inulin hydrolysis. Adapted from [152].
I.3.2. From inulin to fructose
Inulin belongs to the class of carbohydrates known as fructans, which is a natural storage
polymer found widely in plants such as chicory, artichoke and banana [154]. Inulin consists of fructose
molecules linked by β-(2→1) glycosidic bonds, which are responsible for its nutritional characteristics.
It may contain either a terminal -D-fructose or a -D-glucose molecule [155].
The number of fructose units can vary from few units to about 70 units depending on the origin.
Figure 15 exemplifies the chemical structure of inulin, were “n” represents the number of fructose
units. Considering the enzymatic reaction through the use of inulinase the final product ratio is
dependent on the number of units of fructose and glucose in inulin.
Besides the potential source of fructose, inulin is used as an indigestible soluble dietary fiber
and thickener in foods. Also, it can be used for the production of fructooligosaccharides (FOS) with 3
to 8 units of monomers. FOS have a prebiotic activity in human health since they stimulate the growth
of a limited number of bacteria in the colon, increasing the effectiveness of probiotic products [156],
[157]. Also, inulin is associated with the decrease in glucose blood level and the improvement of
calcium absorption [158].
Concerning the product, fructose (C6H12O6), it is a monosaccharide present in a large number of
fruits, particularly in apples and tomatoes, in honey, and has been also detected in some mammal’s
semen. It can be observed in two enantiomers: D-fructose and L-fructose; the last one cannot be
found in nature [159]. Another important property is that fructose is 1.3 times sweeter than sucrose on
an equal solids weight basis [160]. This means that any current application were sucrose is used as
sweetener can be performed identically by using a quantity of fructose reduced by a factor of
1.3 [159]. Due to its sweetness, fructose is emerging in the world of sweeteners facing the competition
with sucrose, which causes problems related to corpulence, carcinogenicity, arteriosclerosis and
diabetes [161]. The use of fructose in food and pharmaceutical formulations has gained some
relevance because of its sweetness properties. In the food sector, fructose are used as sweetening
agents and in different formulations, viz. as stabilizing agent [159]. Also, the pharmaceutical sector
24
uses fructose in tablets, syrups, and solutions as a flavoring and sweetening agent. The market for
HFCS rose steadily until 2000, as evidenced by the production in the USA, which is increased from
2.2 million tons in 1980 to 9.4 million tons in 1999 [162]. From then on there has been a slow decline
[163] to around roughly the 90% of the peak value, but it is still an impressive market.
I.3.3. Inulin hydrolysis for fructose production
The hydrolysis of inulin can be acid or enzymatic. Although the enzymatic method tends to gain
relevance due to his high yields and minimal product contamination, the acid hydrolysis has been
mostly used until nowadays.
In the case of acid hydrolysis of inulin, it can occur at low pH and high temperatures (80-100°C),
using organic or mineral acids or through heterogeneous catalysis using solid acid catalysts, such as
acid-cation exchange resins, zeolites or oxidized activated carbon [162]. The disadvantages of this
chemical approach are the formation of unwanted by-products and of colored or color forming
compounds, which lower product yield and require a more demanding downstream processing [164].
The enzymatic hydrolysis of inulin can be carried out by inulinase preparations, typically
displaying both exo- and endo-activity. Until now, the most common commercially available enzyme
formulation was Fructozyme L, from Novozymes. It is a mixture, composed by approximately 10% of
endoinulinase from Aspergillus niger (EC 3.2.1.7) and 90% of exoinulinase from Bacillus
stearothermophilus (EC 3.2.1.80). However this preparation is no longer available. Recently, another
formulation of a mixture with exo and endoinulinase Aspergillus niger is commercialized by Megazyme
(Fructanase Mixture) [165]. Until now no immobilization studies were published using this enzymatic
formulation.
Pandey et al. (1999) noted that fungal inulinases exhibited an optimum pH between 4.5 and 7.0,
yeast inulinases between 4.4 and 6.5 and bacterial inulinases between 4.8 and 7.0 [166]. Optimal
temperature values were generally higher for bacteria and yeasts than for fungi. The values from pH
and temperature are not necessarily those that will be used in an industrial process because of inulin
solubility, enzyme stability or risk of contamination for instance [159]. Other properties, such as
molecular weight, kinetic parameters, type of substrate are strictly related to the producing micro-
organism.
According to Carniti et al. (1991) the rate of reaction increases with inulin concentration and no
inhibitory effect was observed at any concentration [167]. He also noticed that a significant inulin
concentration is desirable because the presence of the substrate stabilizes the enzyme [159].
Fructose inhibition has been hinted, but this only occurs at initial fructose concentrations over those
that are expected at the end of a typical bioconversion run, starting with a saturated inulin solution at
50ºC. This effect could be explained by the reaction equilibrium thermodynamics or by diffusion
competition between large substrates molecules and small products [155].
25
I.3.4. Inulinase immobilization - review
In the late 1970s, inulinase enzymes started to be immobilized for use in continuous systems.
One of the first works on inulinase immobilization was by Nahm et al. (1979), who immobilized
inulinase from Kluyveromyces fragilis on Tygon tubes by silanation in chloroform with 10%
glutaraldehyde [159]. Table 7 reports some immobilized studies of inulinase in different type of
matrices.
Table 7. Studies of inulinase immobilization in different matrix composition.
Type of inulinase Matrix composition References
Fructozyme L* Alginate beads [168]
Inulinase from Aspergillus fumigatus Chitin, casein and sodium alginate beads [169]
Fructozyme L* Amberlite IRC 50 particles [170]
Fructozyme L* PVA lenses (Lentikats) [27]
Endoinulinase from Aspergillus niger Chitosan beads [171]
Inulinase from Kluyveromyces marxianus
Sodium alginate beads cross-linking with GA and activated coal
[172]
Fructozyme L* PVA (Lentikats) lenses cross-linking with GA [98]
Endoinulinase from Aspergillus niger Niobium beads [173]
Crude inulinase from Aspergillus niger Chitosan beads [174]
Fructozyme L* Polyurethane foam [175]
Fructozyme L* Alginate–chitosan beads [176]
*commercial preparation of inulinases from Aspergillus niger and Bacillus stearothermophilus
I.4. Types of Reactors
Since one of the most important applications of immobilized enzymes is industry, is important to
discuss the main types of reactors used. The use of immobilized enzymes in industrial processes is
performed in chemical reactors. A classification of enzyme reactor based on the mode of operation
and the flow characteristics of substrate and product is summarized in Table 8.
Table 8. Classification of enzyme reactors. Adapted from [177].
Mode of operation Flow pattern Type of reactors
Batch Well mixed Batch stirred tank reactor (BSTR)
Plug flow Total recycle reactor
Continuous
Well mixed Continuous stirred tank reactor (CSTR)
CSTR with membrane
Plug flow
Packed bed reactor (PBR)
Fluidized bed reactor (FBR)
Hollow fiber reactor
26
I.4.1. Batch operation
Batch processes operate in closed systems; substrate is added at the beginning of the process
and products removed only at the end. Batch reactors are the most commonly used type of rector
when free enzymes are used as catalyst. In most cases no recovery of the enzyme is attempted.
On the other hand, when immobilized enzymes are used in a batch operation, an additional
separation is required to recover the enzyme preparation. During this recovery procedure appreciable
loss of immobilized enzyme can occur, as well as, inactivation of the enzyme due to repeated recovery
cycles. It is, therefore, obvious that batch reactors have a rather limited potential in industrial
immobilized enzyme catalysis. Furthermore, the use of a simple batch reactor does not take
advantage of the potential of continuous operation, a major feature of immobilized enzyme systems.
The stirred tank reactor (STR) is the geometry which allows batch operation and the control of
pH and temperature is straightforward. However, when immobilized enzymes are used is the shear
stress induced by the stirring which create a hazardous environment for immobilized biocatalysts,
particularly when hydrogels are used, since most of them are prone to abrasion. In this case, a
possible alternative to the STR is the basket tank reactor in which the enzyme is retained within a
“basket”, either forming the impeller “blades” or the baffles of the tank reactor. This process reduces
the shear stress in the immobilized catalysts, easing their application in a stirred tank reactor and
operation under continuous flow. However this type of reactor is seldom used due to significant mass
transfer resistances associated [178].
Additionally, if the initial concentration of substrate is either harmful or inhibitory to the enzyme,
then a fed batch mode may be used.
I.4.2. Continuous operation
The continuous mode exhibits advantages regarding batch mode, namely: easy of automatic
control, of automation, easy to operate and quality control of the products.
The most common enzymatic reactor configurations for continuous operation are the continuous
stirred tank reactor (CSTR) and the plug flow reactor (PFR). These reactors are fundamentally
different and represent two extremes, i.e. complete mixing in the tank reactor and no mixing at all in
the PFR.
Consequently, conditions within the CSTR are the same as the outlet stream, while in the PFR
the conditions vary with length from inlet to outlet. While the CSTR operates under uniform conditions
of low substrate and high product concentrations, the conditions of the PFR are never uniform. A
nearly ideal CSTR is readily obtained in practice, but an ideal PFR is very difficult to construct. Several
factors can give rise to deviations from a plug flow pattern in a packed-bed reactor. Temperature and
velocity gradients normal to the flow direction and substrate diffusion in the axial direction are the most
frequently occurring complications, and even small deviations from the idealized flow pattern can alter
27
the kinetics of the reaction considerably. In practice, tracing experiments should be carried out to
establish deviation from ideality.
The use of membrane bioreactors (MBR) is another configuration that allows continuous
operation, thus overcoming batch reactors drawbacks such as batch-to-batch variation, high labor
costs, frequent start-up and shut-down procedures. The basic feature of MBR is the separation of the
enzyme, products and substrates by a semi-permeable membrane that creates a selective
physical/chemical barrier. The components are separated through the action of a driving force (viz.
chemical, potential, pressure, electric field). On the other hand the enzyme is physically retained within
the system by the membrane, allowing the establishment of a continuous operation with a feed and
product withdrawal [179].
Micro structured reactors are the most recent advance in bioreactors. Given the advances in
miniaturization, it is possible to develop a chip-like device where the enzymes can be immobilized in a
structured path previously designed. Micro structured rectors have a specific surface area between
10000 and 50000 m2/m
3 and have a flow highly symmetric and mostly laminar [180]. This reduction,
as described already, has some advantages such as the lower volume and cost of equipment and
higher yields [181].
It is known that the use of immobilized enzymes in enzymatic bioreactors is perfectly well-
established. In order to choose the most appropriate reactor system for a certain immobilized enzyme,
several factors should be taken into account, viz. the type of immobilization that is used, the kinetic
parameters and the operational stability of the immobilized enzyme, the mass transfer limitations and
the mode of operation.
I.4.3. Reactors used with immobilized inulinase
Some authors reported on immobilized inulinases used in batch reactors, while others
evaluated the performances of continuous reactors packed with immobilized inulinases on different
supports.
Rhee and Kim (1989) studied the hydrolysis of Jerusalem artichoke tuber juice by inulinase from
Aspergillus ficuum (Novozym 230TM
) in both batch and continuous reactors [182]. Kim et al. (1982)
conducted the hydrolysis of inulin in a PBR after proving that, with other conditions unchanged,
reactors with different height/diameter (H/D) ratios had different steady state conversions [183]. Gill et
al. (2006) performed PBR tests with an exoinulinase from Aspergillus fumigatus immobilized on chitin,
QAE-Sephadex and ConA linked-amino activated silica beads at 60ºC [169]. Wenling et al. (1999)
analyzed different operating conditions. A 4.5% (w/v) inulin solution was completely hydrolyzed at
50ºC by inulinase obtained from Kluyveromyces sp. Y-85 and immobilized on macroporous ionic
polystyrene beads (D201-GM resin) [184].
28
29
Chapter II. Objectives
The use of immobilized enzymes is becoming one of the most powerful tools in biotechnology,
having a profound impact on health, food supply, environmental protection and sustainable fuel
production. Major challenges are referred to the design of robust enzyme immobilization protocol at
reduced price. Also, overcame the low operational stability, still observed in most procedures.
The main goal of the master project presented here is to produce and characterize resilient
hydrogels, displaying high operational and thermal stability. A relatively simple, robust and low-cost
procedure is intended, as to make the process appealing for commercial application and, alongside
being competitive to the existing ones, namely Lentikat liquid preparation.
In order to achieve this goal, inulinase was selected to be the protein model system and, due to
their chemical and biological properties, PVA and chitosan polymers were chosen to hydrogel
production.
For both types of polymers, the initial approach involved:
The immobilization procedures which display the best results, considering concentration of
fructose and physical stability of the matrix, were singled out for detail characterization. The kinetic
parameters, pH and temperature profiles as well as the thermal and storage stabilities were
established for free and immobilized inulinase. Operational stability was also tested in continuous flow
in a miniature, custom-made, packed bed reactor.
In order to accomplish the results, analytical methods mostly spectrophotometric methods (DNS
for reducing sugars and Bradford for protein quantification) and HPLC analysis were performed. Also,
SDS-PAGE was performed to evaluate the size of inulinase(s).
Lastly, the results will be compared with literature and commercial preparations available.
Polivynil Alcohol (PVA)
Inulinase entrapment in PVA films
Different PVA molecular weights and
volumes of cross-linker were tested;
Enzymatic hydrolysis assays at different
temperatures were accessed.
Chitosan (CS)
Inulinase entrapment in CS beads
The effect of different treatments (upon
immobilization) was analyzed;
Enzymatic hydrolysis assays at different
temperatures were performed.
30
31
Chapter III. Materials and Methods
III.1. Chemicals and Materials
Fructanase Mixture (batch number 121001), a commercial mixture of endo and exoinulinase
from Aspergillus niger, was provided by Megazyme (Bray, Ireland). The enzyme was kept at -20°C as
suggested by the manufacturer. Chitosan (CS) (average MW 200 kDa) was from Acros Organics
(Geel, Belgium). Polyvinyl alcohol (PVA) (98.0%-98.8% hydrolyzed, average MW 50 kDa) from Acros
Organics. Mowiol 28-99 (PVA, average MW 145 kDa) and Mowiol
10-98 (PVA, average MW 61 kDa)
were from Sigma-Aldrich (St Louis, USA). Glutaraldehyde (GA), as 24% (w/w) aqueous solution, and
sodium tripolyphosphate (TPP), as 85% pure, were purchased from Acros Organics. Inulin (Fibruline
S-20 with an average polymerization degree of about 10-12 units) was an offer by Induxtra (Moita,
Portugal). All solutions were prepared in distilled water. All other chemicals were of analytical grade
from various suppliers.
Spectrophotometer SPECTRA MAX PLUS 384 was from Molecular Devices (Sunnyvale, USA).
Controlled temperature environment was promoted using either a Thermomix MM thermoregulator
from B. Braun (Melsungen, Germany), an immersion thermostat ECO E4 from LAUDA (Lauda-
Königshofen, Germany) or an orbital shaker Agitorb 160 E from Aralab Equipamentos de Laboratório
e Electromecânica Geral, LDA (Albarraque, Portugal). Peristaltic pump Watson Marlow 205S (ERT,
Lisboa, Portugal) and 2-stop tubing with an internal diameter of 0.48 mm and an external diameter of
0.91 mm from IDEX ISMATEC (Qlabo, Lisboa, Portugal). The reactor was designed and assembled in
acrylic at Acrilicos Fernando Gil (Lisboa, Portugal). The reactor (Figure 16) has 3 cm height for 1 cm
width and a pre-chamber at both ends with 3 mm height and 9 mm width. Filter paper was inserted
into inlet and outlet to avoid clogging.
Figure 16. The miniature tubular reactor configuration used for continuous flow operation.
32
III.2. Analytical methods
III.2.1. Quantification of reducing sugars by DNS method
Reducing sugars were quantified by the 3,5-dinitrosalicylic acid (DNS) method developed by
Miller [185], but adapted to a microplate format, as described by Nunes et al. (2010) [186]. Briefly,
10 µL of sample were added to 90 µL of distilled water. 100 µL of DNS was added to each standard or
unknown sample replicate. In order to have rapid analyze result, 10 µL of acetate buffer solution
100 mM at pH 4.5 or 5.0 was used as negative control, whereas 10 µL of fructose 50 g.L-1
was used
as positive control. Then, an incubation at 100°C for 5 minutes (the plate was covered to avoid
evaporation) was performed. After the plate was cooled at room temperature, 500 µL of distilled water
was added and 200 μL of the resulting solutions was transferred to 96 well microplates (ELISA plates).
Then, the absorbance at 540 nm was measured for the quantification of the reducing sugars, which
was performed with reference to a calibration curve previously established (Annex I). Standardization
was obtained with fructose concentrations ranging from 0 to 5 g.L-1
. All samples and diluted fructose
standards were analyzed in triplicate at least.
III.2.2. Quantification of reducing sugars by HPLC method
In order to establish a comparison between the results from DNS and to guarantee the use of
DNS as a viable and fast method, the quantification of fructose in high performance liquid
chromatography (HPLC) was also carried out. The samples were first diluted 1:2 (v/v) with a H2SO4 50
mM solution and centrifuged for 2 minutes at maximum speed. The supernatant was then diluted 1:10
(v/v) with a H2SO4 50 mM solution for a final volume of 1 mL in the HPLC vial. The HPLC run used 20
µL of sample. The RI-detector L-2490 Elite Lachrom was form Hitachi and the analytical HPLC column
Rezex ROA – organic acid 8% cross linking (30 mm) was from Phenomenex, USA. The area of the
samples was compared with the calibration curve (Annex I) that was prepared using the same
procedure.
III.2.3. Bradford protein quantification
Quantification of protein (inulinase) was carried according to Bradford (1976) [187], but adapted
to micromethod, as described by Nunes et al. (2010) [186]. In the Bradford microplate procedure,
150 µL of sample and of each diluted inulinase standards were added to 150 µL of Bradford reagent
(Sigma–Aldrich), in a 96 well microplate (ELISA plates). A solution of acetate buffer 100 mM pH 4.5 or
5.0 was used as blank. The solution was mixed and incubated during 10 minutes at room temperature.
Then, the absorbance at 595 nm was measured for the quantification of the protein, which was
performed with reference to a calibration curve previously established (Annex II). Standardization was
obtained with bovine serum albumin (BSA) (Thermo Scientific Pierce BSA Protein Assay Standards,
33
Thermo Fisher Scientific, USA) concentrations ranging from 2.5 to 20 µg.mL-1
. All samples and diluted
inulinase standards were analyzed in triplicate.
III.2.4. Protein characterization by SDS-PAGE
Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) was performed to
evaluate the size of inulinase(s). The buffer sample was prepared with 62.5 mM Tris–HCl pH 6.2,
2% (w/v) SDS, 0.01% (w/v) bromophenol blue, and 10% (v/v) glycerol. The samples were denaturated
in reducing conditions with 100 mM dithiothreitol (DTT) at 100°C for 5-10 minutes. Samples were
loaded in a 12% acrylamide gel, prepared from a 40% acrylamide/bis stock solution (29:1) (Bio-Rad),
and ran at 90 mV using a running buffer composed by glycine 192 mM, Tris 25 mM, and 0.1% SDS
pH 8.3. Gels were stained with Coomassie PhastGel (Pharmacia AB Laboratory Separations®). Gels
were de-stained using a 30% ethanol/10% acetic acid solution. The molecular markers used were
Precision Plus Protein Dual Color (BioRad).
III.3. Immobilization procedures
III.3.1. Enzyme immobilization in polyvinyl alcohol (PVA) film
In order to entrap inulinase in PVA film, a PVA solution (5% w/v in aqueous solution) was
prepared by heating at 100ºC and then cooled to 40ºC. Before the addition of enzyme suspension, the
pH was adjusted to 4.5, with hydrochloric acid 5 M or sodium hydroxide 2 M. The enzyme suspension
(500 µL of a 10-fold diluted preparation in acetate buffer 100 mM pH 4.5) was added to the PVA
solution and mixed under mild magnetic stirring. Then, pH was again adjusted to 4.5 and 2 mL of this
enzyme enriched solution was added to each of the four screw-capped vessels. After, different
volumes of GA (50 µL, 100 µL, 200 µL and 400 µL) were added to the corresponding vessel. The
solutions were incubated for 16 hours at 35ºC to promote a controlled dehydration. After film
formation, it was rinsed with distilled water and the excess of water was removed with filter paper. The
film was weighted and stored in acetate buffer 100 mM pH 4.5 at 4°C until use. Figure 17 represents
the enzyme immobilization in PVA.
Figure 17. Schematic representation of enzyme immobilization in PVA.
Incubation at 35ºC
PVA solution (pH 4.5)
Addition of enzyme diluted
solution
Enzyme enriched solution (pH 4.5)
2 mL to each vessel
Addition of GA 50µL 100µL
200µL 400µL
34
The immobilization procedure was performed for three different PVA molecular weights, 50 kDa,
61 kDa and 145 kDa.
To evaluate the effect of different volumes of cross-linker added as well as temperature on
immobilized biocatalyst, enzymatic hydrolysis assays were performed in 10 mL screw-capped vessels
at different incubation temperatures (45ºC, 50ºC, 55ºC, 60ºC and 65ºC) with a stirring rate of
1000 rpm. Approximately, 100 mg of each PVA film, with different MW and volume of GA, were
weighted. For each temperature, 1 mL of 5% (w/v) inulin substrate solution in acetate buffer 100 mM
pH 4.5 was added to the vessel. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the
beginning of incubation and immediately assayed for reduced sugars. Results are expressed as
concentration of fructose (g.L-1
) over time (hour).
III.3.2. Enzyme immobilization in chitosan (CS) beads
For the CS immobilization inulinase protocol, a methodology adapted from Wentworth et al.
(2003) [122] was used. Briefly, 1.5% (w/v) CS solution in 3% (v/v) acetic acid was prepared and stirred
for 2 hours. 100 µL of enzyme suspension (10-fold diluted preparation in acetate buffer 100 mM
pH 4.5) was added to 900 µL of CS solution and mixed under magnetic stirring. The resulting solution
was extruded through a 1 mL needle (Therumo Neolus, 20 G×2’’, Therumo Corp., Leuven, Belgium)
into a solution of 15% (w/v) sodium tripolyphosphate (pH 5.0) at a vertical distance of approximately
15 cm, under mild stirring. After extrusion and curing for 10 minutes the beads were removed from the
sodium tripolyphosphate solution and rinsed with distilled water. Excess of water in the beads was
removed with filter paper.
A second immobilization approach was executed using more concentrated CS solutions (35.0,
48.3 and 68.3 g.L-1
). 100 µL of enzyme suspension (10-fold diluted preparation in acetate buffer
100 mM pH 4.5) was added to 900 µL of CS solution and mixed under the vortex. The resulting
solution was extruded through a 1 mL needle (Therumo Neolus, 20 G×2’’, Therumo Corp., Leuven,
Belgium) into a solution of 15% (w/v) sodium tripolyphosphate (pH 5.0) at a vertical distance of
approximately 15 cm, under mild stirring. After extrusion and curing for 10 minutes the beads were
removed from the sodium tripolyphosphate solution and rinsed with distilled water. Excess of water in
the beads was removed with filter paper.
After immobilization procedure, CS beads were weighted and divided into three parts. One
portion for stabilization in acetate buffer 100 mM pH 4.5 at 4°C (absence of treatment), a second
portion of the beads were dehydrated in desiccator during 15 minutes and the remaining beads were
reinforced with 500 µL of GA during 1 hour under magnetic stirring (850 rpm). Then, they were
weighted and stored to stabilize in acetate buffer 100 mM pH 4.5 at 4°C until use.
In Figure 18 is the schematic representation of the immobilization of inulinase in chitosan
beads.
35
Figure 18. Schematic representation of enzyme immobilization in CS.
To study the influence of each applied treatment as well as temperature on immobilized
biocatalyst, enzymatic hydrolysis assays were performed in 10 mL screw-capped vessels at different
incubation temperatures (50ºC, 55ºC, 60ºC and 65ºC) with a stirring rate of 1000 rpm. Approximately,
100 mg of each type of CS matrix (absence of treatment, dehydrated or reinforced with GA) were
weighted. For each temperature, 1 mL of 5% (w/v) inulin substrate solution in acetate buffer 100 mM
pH 4.5 was added to the vessel. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24 hours after the
beginning of incubation and immediately assayed for reduced sugars. Results are expressed as
concentration of fructose (g.L-1
) over time (hour).
The procedures described henceforth were only performed with the matrices selected, namely
PVA film 50 kDa reticulated with 400 µL of GA and CS beads reinforced with GA.
III.4. Immobilization efficiency, protein entrapment and immobilization yield
The amount of protein loaded on the matrix was calculated from the difference of initially added
protein to the protein obtained in the washing plus supernatant. The protein content was calculated
using the Bradford micro-method optimized for these studies (c.f. III.2.3).
The immobilization efficiency was defined in the Equation 1:
𝐼𝑚𝑚𝑜𝑏𝑖𝑙𝑖𝑧𝑎𝑡𝑖𝑜𝑛 𝑒𝑓𝑓𝑖𝑐𝑖𝑒𝑛𝑐𝑦 (%) = 𝑇𝑜𝑡𝑎𝑙 𝐼𝐴 𝑎𝑑𝑑𝑒𝑑−𝐼𝐴 𝑖𝑛 𝑡ℎ𝑒 𝑠𝑢𝑝𝑒𝑟𝑛𝑎𝑡𝑎𝑛𝑡
𝑇𝑜𝑡𝑎𝑙 𝐼𝐴 𝑎𝑑𝑑𝑒𝑑× 100 (1)
whereas IA stands for inulinase activity.
Protein entrapment was determined as follows:
𝑃𝑟𝑜𝑡𝑒𝑖𝑛 𝑒𝑛𝑡𝑟𝑎𝑝𝑚𝑒𝑛𝑡 (%) = 𝑇𝑜𝑡𝑎𝑙 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑎𝑑𝑑𝑒𝑑 − 𝑃𝑟𝑜𝑡𝑒𝑖𝑛 𝑖𝑛 𝑡ℎ𝑒 𝑠𝑢𝑝𝑒𝑟𝑛𝑎𝑡𝑎𝑛𝑡
𝑇𝑜𝑡𝑎𝑙 𝑝𝑟𝑜𝑡𝑒𝑖𝑛 𝑎𝑑𝑑𝑒𝑑× 100 (2)
Sodium tripolyphosphate solution
Chitosan solution with inulinase
Beads
36
At immobilization step, the matrices were washed with distilled water to remove the excess of
cross-linker and unbound enzyme. The immobilization yield (%) was determined as defined in the
Equation 3:
𝐼𝑚𝑚𝑜𝑏𝑖𝑙𝑖𝑧𝑎𝑡𝑖𝑜𝑛 𝑦𝑖𝑒𝑙𝑑 (%) = 𝑆𝑝𝑒𝑐𝑖𝑓𝑖𝑐 𝑎𝑐𝑡𝑖𝑣𝑖𝑡𝑦 𝑜𝑓 𝑖𝑚𝑚𝑜𝑏𝑖𝑙𝑖𝑧𝑒𝑑 𝑒𝑛𝑧𝑦𝑚𝑒
𝑆𝑝𝑒𝑐𝑖𝑓𝑖𝑐 𝑎𝑐𝑡𝑖𝑣𝑖𝑡𝑦 𝑜𝑓 𝑓𝑟𝑒𝑒 𝑒𝑛𝑧𝑦𝑚𝑒× 100 (3)
Enzyme activity (U) is defined as the amount of protein needed to convert 1 µmol of reducing
sugars in 1 minute.
III.5. Bioconversion studies
Inulinase bioconversion studies were carried out in magnetically stirred (850 rpm) 10 mL screw-
caped vessels at 50ºC (free enzyme), 55ºC (enzyme immobilized in PVA) and 65ºC (enzyme
immobilized in CS). A 2 mL of a 5% (w/v) inulin prepared in acetate buffer 100 mM pH 4.5 (free
enzyme) or pH 5.0 (immobilized enzyme) with either: 50 µL of a 100-fold diluted preparation of
Fructanase Mixture; 60 mg of immobilized enzyme in CS beads or 100 mg of immobilized enzyme in
PVA film. When assessing inulinase activity in supernatants, 500 μL of supernatant were mixed with
3 mL of a 5.6% (w/v) solution of inulin in acetate buffer 100 mM pH 4.5 containing. The bioconversion
runs were performed at 50ºC in 10 mL screw-capped, magnetically stirred (850 rpm) reactors.
Samples were collected: every 5 minutes until 40 minutes for the immobilized enzyme; every 3
minutes until 15 minutes for the free enzyme. Then, samples (10 µL) were immediately quenched in
DNS reagent and assayed for quantification of reducing sugars. Enzyme activity was determined
through the initial reaction rates. Each bioconversion run was performed in duplicate, at least.
III.6. pH and temperature profiles
The effects of pH and temperature on enzyme activity were evaluated in batch runs by
incubating either form of the enzyme in inulin solutions, 5% (w/v) in acetate buffer 100 mM, in a pH
range of 3.6 to 5.5, and in a temperature range of 45ºC to 70ºC. The conditions of enzymatic assays
were performed as described in Bioconversion studies. Also, samples (150 µL) were collected from
supernatant for protein detection by Bradford method. All runs were performed in triplicate. Results
were represented as relative activity (expressed as percentage), as follows:
𝑅𝑒𝑙𝑎𝑡𝑖𝑣𝑒 𝑎𝑐𝑡𝑖𝑣𝑖𝑡𝑦 (%) = 𝐴𝑐𝑡𝑖𝑣𝑖𝑡𝑦
𝑀𝑎𝑥𝑖𝑚𝑢𝑚 𝑎𝑐𝑡𝑖𝑣𝑖𝑡𝑦 × 100 (4)
37
III.7. Optimal mass of matrix for conversion
For the determination of the effect of matrix weight in the reaction different amounts of matrix
were tested. For this assay PVA film and CS beads were used. Runs were performed in 10 mL
screw-capped vessels, filled with 2 mL of a 5% (w/v) inulin solution in acetate buffer 100 mM pH 5.0.
Matrix weights of 40 mg, 80 mg, 120 mg and 200 mg were used. The agitation was promoted by
magnetic stirring (850 rpm) at 55°C and 65ºC for PVA and CS immobilized inulinase, respectively. The
determination of enzyme activity was performed as described in Bioconversion studies. All the assays
were performed in duplicate. The results were converted in relative activity (Equation 4).
III.8. Determination of kinetic parameters
The effect of substrate concentration in the free and immobilized inulinase activity was tested
with different concentrations of inulin: 1.5%, 2.5%, 5.0%, 7.5%, 10.0%, 12.0% and 15.0% (w/v) in
acetate buffer 100 mM pH 4.5 (for the free enzyme) or pH 5.0 (for the immobilized enzyme). For each
run, 60 mg of CS beads and 100 mg of PVA film were weighted and 50 μL of 100-fold diluted
preparation of free enzyme were used. Initial reaction rates were carried out as described in
Bioconversion studies. The results were then processed and presented as specific activity (activity per
mgenzyme) vs. inulin concentrations (g.L-1
). Determination of kinetic parameters, Vmax and KM, was
carried out through Hyper32 software and confirmed by excel using non-linear regression analysis.
III.9. Stability studies
III.9.1. Thermal stability
The thermostability of the free and immobilized inulinase was determined under different
temperatures (40ºC, 50ºC, 60ºC and 70ºC) for different time spans, namely: 48 hours for the 40C
assay; 30 hours for the 50C assay; 10 hours for the 60C and 5 hours for 70ºC. 1.5 g of PVA film and
800 mg of CS beads were weighted and incubated in 25 mL screw capped vessels, filled with 10 mL
of acetate buffer 100 mM pH 5.0, with agitation promoted by magnetic stirring (850 rpm) at the
determined temperature. Also, free inulinase as 100-fold diluted preparation in acetate buffer 100 mM
pH 4.5 was incubated at defined temperatures with the same agitation. Biocatalyst, either in
immobilized or free form was recovered at different times depending on the temperature, and initial
activity assays were performed as described in Bioconversion studies. Simultaneously, samples from
the supernatant were taken to assess protein leakage. The results were then analyzed and processed
in order to determine the deactivation pattern according to the three-parameter bi-exponential model
[188]. All the assays were performed in duplicate.
38
III.9.2. Storage stability
Given amounts of immobilized biocatalyst PVA film and CS beads were stored (100 mg and
60 mg for each independent run, respectively) at 4ºC in 25 mL screw capped vessels filled with 10 mL
acetate buffer 100 mM pH 5.0. Also, 1 mL of free enzyme (100-fold diluted preparation in acetate
buffer 100 mM pH 4.5) was stored. The storage stability was evaluated by determining the enzyme
activity up to 180 days. Bioconversion studies and initial activity assays were performed as described
in Section III.5. At the same time, a supernatant sample was collected to access protein leakage from
immobilization matrices. The data was analyzed through the to the three-parameter bi-exponential
model [188]. All the assays were performed in duplicate.
III.9.3. Operational stability
Operational stability was evaluated under continuous flow operation using a miniature tubular
reactor for production of fructose. The reactor was immersed in a temperature-controlled bath and
feeding was provided through a peristaltic pump. The assay was performed for both immobilization
forms (PVA and CS).
III.9.3.1. Determination of flow rate and reactor volume
Distilled water was used for the calibration of the flow rates provided by the peristaltic pump
using the mentioned tubing. The assay was performed in duplicate. For a given speed (rpm), distilled
water samples were collected to an Eppendorf until 1 mL was achieved. The time to fulfill that volume
was measured using a chronometer and the flow rate was determined (mL.min-1
). The procedure was
applied to several speed values (0.5 to 10 rpm) and the calibration curve relating speed (rpm) and flow
rate (mL.min-1
) was established (Annex III).
The void volume was determined by filling the reactor with the correspondent biocatalyst, PVA
or CS, and adding acetate buffer 100 mM pH 5.0. The volume between CS beads or PVA film was
determined, which corresponded to the void volume (V).
The relation between total reactor volume (Vt), void volume (V) and volume of matrix (Vm) is
given by: Vt = V + Vm. The total reactor volume was 2.75 mL.
III.9.3.2. Continuous production of fructose
Continuous production of fructose was performed during 31 and 47 days at 50ºC and 55ºC for
immobilization in PVA film and CS bead, respectively. The reactor was filled with 3.450 g of PVA film,
cut into cubes with dimensions of 2×2×2 mm in average, and 1.206 g of CS beads. A 5% (w/v) inulin
solution in acetate buffer 100 mM pH 5.0 was fed to the reactor at a flow rate of 0.0053 mL.min-1
.
39
Samples were collected on a daily basis and assayed for reducing sugars as well as for protein
detection. Figure 19 represents the set-up assembled for the continuous production of fructose. As a
side note, the set-up and the reactor described were used for both types of immobilized biocatalyst.
Results were represented as product yield (expressed as percentage), as follows:
𝑃𝑟𝑜𝑑𝑢𝑐𝑡 𝑦𝑖𝑒𝑙𝑑 (%) = [ 𝐹𝑟𝑢𝑐𝑡𝑜𝑠𝑒 𝑓𝑜𝑟𝑚𝑒𝑑]
[𝐼𝑛𝑖𝑡𝑖𝑎𝑙 𝑓𝑟𝑢𝑐𝑡𝑜𝑠𝑒 𝑓𝑜𝑟𝑚𝑒𝑑] × 100 (5)
Figure 19. Representation the set-up assembled for the continuous production of fructose. The miniature tubular
reactor was immersed in a temperature-controlled batch and feeding was provided through the peristaltic pump.
(A) Inlet – 5% (w/v) inulin solution in acetate buffer 100 mM pH 5.0; (B) Peristaltic pump; (C) Thermostat ECO E4
where is immersed the reactor and is covered to avoid heat loss (D) Outlet – for DNS and protein analysis.
A
B C
D
40
41
Chapter IV. Results and Discussion
First part
Preliminary studies were carried out in order to identify the most promising matrices for the
immobilization of inulinase in PVA film and CS beads, which were afterwards characterized with more
detail.
IV.1. Polyvinyl alcohol (PVA) film immobilized inulinase
The feasibility of the proposed methodology for enzyme immobilization in PVA was assessed
primarily by evaluating the film formation. The effect of different MW and volume of cross-linker (GA)
on film formation is presented in Table 9. The formation of immobilization matrices was effectively
accomplished, only one solution did not solidify (PVA solution of 61 kDa with 50 µL) (Figure 20). The
matrices had a cylindrical shape with approximately 5 mm of height and 15 mm of diameter, in
average.
Table 9. Effect of different PVA molecular weight and volume of cross-linker (GA) on film formation
Polymer
(PVA) MW
Volume of
cross-linker (GA)
Solidification
Process Film Formation
Film appearance and
weight
50 kDa
50 µL
Incubation
during 16 hours
at 35ºC
Yes Transparent / 0.353 g
100 µL Yes Transparent / 0.527 g
200 µL Yes Transparent / 0.625 g
400 µL Yes Transparent / 1.229 g
61 kDa
50 µL No (liquid state) -
100 µL Yes Transparent / 1.340 g
200 µL Yes Transparent / 0.566 g
400 µL Yes Transparent / 1.338 g
145 kDa
50 µL Yes Transparent / 0.249 g
100 µL Yes Transparent / 0.370 g
200 µL Yes Transparent / 0.486 g
400 µL Yes Transparent / 0.663 g
Figure 20. Representative image of PVA film formed (61 kDa with 400 µL of GA) concerning the conditions
previous described, inside the screw vessel (A) or removed from the vessel (B); Representative image of PVA
solution of 61 kDa with 50 µL GA, which was not solidified (C).
A
B C
42
The process of cross-linking with GA presents several advantages, as already described,
among them the fact that the method is quite simple and highly efficient. Also, GA can bind
nonspecifically to proteins. The reaction mechanism of PVA with GA was described in Figure 10.
A relationship between the volume of GA and matrix weight may be established: it seemed that higher
volumes of GA led to an increase in film weight (Table 9).
In order to evaluate the effect of temperature and volume of GA on the enzyme activity (as
concentration of fructose over time) and on the resilience of catalytically active PVA films, incubation
of inulinase immobilized in PVA films at different controlled temperatures (40ºC, 45ºC, 50ºC, 55ºC,
60ºC and 65ºC) was performed.
In a first approach, all the PVA films formed were tested at 40ºC in order to evaluate inulin
hydrolysis and their endurance towards several hours of incubation. The results are expressed as
concentration of fructose (g.L-1
) over time (hour). Concerning the thermostability at 40ºC (Figure 21),
after 2 hours of reaction the PVA films of 50 kDa with 50 µL and 100 µL of GA were in a liquid state.
Therefore, they were not considered in the study.
From Figure 21 is possible to observe that in all cases the concentration of fructose increases
with the time, ultimately leading to close to or full hydrolysis of inulin after 24 hours. No trace of protein
was detected in the supernatant by Bradford method.
The catalytic activity of the immobilized enzyme formulations globally seems to decrease with
the increase in the MW of the polymer, a feature noticeable from 2.5 hours of incubation. The higher
polymer size may have resulted in an increase in mass transfer resistance, thus reducing the overall
reaction rate. Nonetheless, a higher PVA molecular weight has been associated with permanent
entrapment of enzymes in PVA [189].
Regarding the effect of GA on each PVA molecular weight, the enzyme activity seemed to
decrease with the increase in the volume of GA, an exception was observed for 61 kDa with 400 µL.
Such behavior could be justified by the GA toxicity effect and, it is in agreement with the results
obtained by Chui and Wan (1997) [190]. According to the authors, the enzymatic activity was inversely
proportional to the concentration of GA used, caused by enzyme structure distortion as result of
extensive cross-linking. Additionally, the increase volume of GA results in higher availability of
molecules to form linkages with the polymeric chains of PVA. This leads to an increase in the density
of hydrogels and to higher resistance to mass transfer [70].
Also, it is important to notice that the immobilization protocol was performed at pH 4.5, which is
not reported in the literature. Generally, the reaction of cross-linking is performed using acid catalysts,
which in most cases leads to enzyme denaturation. Figueiredo et al. (2009) performed a cross-linking
reaction at pH 6.2 (acid-free reaction) and 40ºC, however no biomolecules were immobilized [85].
43
5 0 k D A
T im e (h o u r )
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
2 0 0 L G A
4 0 0 L G A
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
6 1 k D A
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
1 0 0 L G A
2 0 0 L G A
4 0 0 L G A
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
5 0 L G A
1 0 0 L G A
2 0 0 L G A
4 0 0 L G A
1 4 5 k D A
Figure 21. Influence of the volume of GA added on the concentration of fructose (g.L-1
) formed from inulin
hydrolysis at 40ºC. The assay was performed for PVA films of 50 kDa (A), 61 kDa (B) and 145 kDa (C) and
different volumes of GA (50 µL, 100 µL, 200 µL and 400 µL). Bioconversion runs were performed in 5% (w/v)
inulin in acetate buffer pH 4.5. 100 mg of immobilized inulinase was used as biocatalyst. Samples (10 µL) were
collected at 2.5, 3.5, 6.5 and 24 hours after the beginning of incubation. Standard deviation did not exceed 5%.
Since the PVA films of 50 kDa with 200 µL and with 400 µL GA, 61 kDa with 400 µL GA and
145 kDa with 50 µL GA were physically stable at 40ºC (did not melt) and displayed higher hydrolytic
activity, the same study was performed at 45ºC, 50ºC and 55ºC (Figure 22).
A B
C
A
B
C
44
4 5 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
5 0 k D a 2 0 0 µ L
5 0 k D a 4 0 0 µ L
6 1 k D a 4 0 0 µ L
1 4 5 k D a 5 0 µ L
5 0 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
5 0 k D a 2 0 0 µ L
5 0 k D a 4 0 0 µ L
6 1 k D a 4 0 0 µ L
1 4 5 k D a 5 0 µ L
5 5 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
5 0 k D a 2 0 0 µ L
5 0 k D a 4 0 0 µ L
6 1 k D a 4 0 0 µ L
1 4 5 k D a 5 0 µ L
Figure 22. Influence of the volume of GA added on the concentration of fructose (g.L
-1) formed from inulin
hydrolysis at 45ºC (A), 50ºC (B) and 55ºC (C). The assay was performed for PVA films of 50 kDa, 61 kDa and
145 kDa and different volumes of GA (50 µL, 200 µL and 400 µL). Bioconversion runs were performed in 5% (w/v)
inulin in acetate buffer pH 4.5. 100 mg of immobilized inulinase was used as biocatalyst. Samples (10 µL) were
collected at 2.5, 3.5, 6.5 and 24 hours after the beginning of incubation. Standard deviation did not exceed 5%.
B
C
A
45
The temperature increase had disparate impacts on the performance of the different
formulations of the immobilized biocatalyst. Thus, under the operational conditions used, the behavior
of particles immobilized in 50 kDa and 61 kDa PVA reticulated with 400 µl GA was, in the overall,
somehow alike, eventually an outcome of the relatively close nature of the polymers used and GA
volume used for reticulation. In almost all cases, these allowed for the highest final titers in fructose
formed, peaking at close to 50.0 0.005 g.L-1
at 50ºC. The final fructose concentration was slightly
inferior (48.0 0.005 g.L-1
) at 45ºC, whereas a more noticeable decrease to around 40.0 0.005 g.L-1
was observed with an increase to 55ºC. On the other hand the increase in temperature to 55ºC slightly
improved enzyme activity in earlier stages (2.5 h) of hydrolysis.
For PVA film of 50 kDa with 200 µL GA, the final product concentration (again close to 50.0
0.005 g.L-1
) also peaked at 50ºC, to decrease at the remaining temperatures. At 55ºC, its activity was
particularly poor. It can be tentatively suggested that the formulations do not provide a stabilizing role
for the enzyme towards the effect of temperature, given the activity/temperature profile obtained for
the free enzyme. Still dedicated assays are required for confirmation. Also, the sample 145 kDa with
50 µL was not considered for further studies, considering the poor enzymatic activity displayed.
Comparing these results with literature, Cattorrini et al. (2009) reported 46.0 and 47.0 g.L-1
of
final product concentration at 50ºC for Lentikats and hemispheric particles [99]. Moreover, these
authors have already reported a loss of rigidity and change of appearance from white to opaque of
Lentikats at 55ºC, suggesting physical instability [27], [99]. Nonetheless, PVA film of 50 kDa did not
display this thermal instability, highlighting the feasibility of this approach when comparing to
commercial applications.
The films 50 kDa with 400 µL and 61 kDa with 400 µL were used for an inulin hydrolysis assay
at 60ºC (Figure 23).
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
5 0 k D a 4 0 0 µ L G A
6 1 k D a 4 0 0 µ L G A
Figure 23. Influence of the volume of GA added on the concentration of fructose (g.L
-1) formed from inulin
hydrolysis at 60ºC. The assay was performed for PVA films of 50 kDa and 61 kDa with 400 µL of GA.
Bioconversion runs were performed in 5% (w/v) inulin in acetate buffer pH 4.5. 100 mg of immobilized inulinase
was used as biocatalyst. Samples (10 µL) were collected at 3.5, 6.5 and 24 hours after the beginning of
incubation. Standard deviation did not exceed 5%.
46
Over the 24 hours incubation period the PVA film of 50 kDa produced a concentration of
fructose of 41.0 0.005 g.L-1
, whereas the film of 61 kDa achieved 32.5 0.005 g.L-1
. Due to this
relatively poor hydrolytic performance, this film was not considered for the assay at 65ºC.
A final study at 65ºC with the PVA film of 50 kDa with 400 µL of GA was performed (Figure 24).
The final product concentration (44.0 0.005 g.L-1
) achieved suggests that the increase in
temperature did not affect the enzyme activity. Also, no protein was detected in the reaction medium.
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
Figure 24. Influence of the volume of GA added on concentration of fructose (g.L-1
) formed from inulin hydrolysis
at 65ºC. The assay was performed for PVA of 50 kDa with 400 µL of GA. Bioconversion was performed in 5%
(w/v) inulin in acetate buffer at pH 4.5. 100 mg of immobilized inulinase was used as biocatalyst. Samples (10 µL)
were collected at 2.5, 3.5, 6.5 and 24 hours after the beginning of incubation. Standard deviation did not
exceed 5%.
Taking into consideration the physical stability of PVA films and the concentration of fructose
after 24 hours of inulin hydrolysis, the PVA with a molecular weight of 50 kDa seemed to display more
promising results. This fact could be explaining by the mass transfer limitations, since the higher the
molecular weight, more limitations of mass transfer were reported [191].
Although the volume of cross-linking was the highest added (400 µL), it seemed to be the most
appropriate in order to provide stability. In this context, a commitment between MW and volume of
cross-linker had to be achieved. Nonetheless, more studies with PVA 50 kDa and different volumes of
GA are essential in order to determine the optimal volume of GA.
47
IV.2. Chitosan (CS) beads immobilized inulinase
The methodology for the immobilization in chitosan beads was performed according to
Wentworth et al. (2003) [122]. However, even following strictly the procedure, the beads did not form
or when its formation occurred, they were not resistant to stress promoted by the stirring. Then, just a
yellow mixture was observed (Figure 25 A).
In this context, some alterations to the procedure had to be implemented. In this context, more
concentrated CS solutions were used (35.0 and 48.3 g.L
-1) and beads were effectively formed.
However neither of these two concentrations improved the consistency of the beads (Figure 25 B and
C). A last CS concentration solution of 68.3 g.L-1
was implemented. The formed beads seemed to be
more consistent (Figure 25 D) and were used for further analysis. Each bead had a diameter between
2 and 3 mm and a weight of 19 mg, in average. After the reinforcement with glutaraldehyde, the beads
appeared to be browner which was a consequence of the formation of Schiff’s base.
Concentration of chitosan solution
Original (A) 35.0 g.L-1
(B) 48.3 g.L-1
(C)
68.3 g.L-1
(D)
Figure 25. Representative images of chitosan beads. No formation of beads through the original procedure (A);
Beads formed with chitosan solutions of 35.0 g.L-1
(B), 48.3 g.L-1
(C) and 68.3 g.L-1
(D). After beads formation they
were divided into three portions, to: stabilization in acetate buffer, dehydration or reinforcement with GA.
Different treatments after immobilization procedure may have influence on the enzyme activity.
Thus, enzyme activity in chitosan beads stabilized in acetate buffer, dehydrated or reinforced with GA
was evaluated under temperatures within 50ºC to 65ºC (Figure 26).
After beads formation
Stabilization in acetate buffer
Reinforcement with GA
Dehydration
48
5 0 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
A b s e n c e D e h y d ra t io n G lu ta ra ld e h y d e
5 5 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
A b s e n c e D e h y d ra t io n G lu ta ra ld e h y d e
6 0 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
A b s e n c e D e h y d ra t io n G lu ta ra ld e h y d e
6 5 º C
T im e (h o u r )
Co
nc
en
tra
tio
n o
f fr
uc
tos
e (
g.L
-1)
0 .0 1 0 .0 2 0 .0 3 0 .0
0 .0
1 0 .0
2 0 .0
3 0 .0
4 0 .0
5 0 .0
6 0 .0
A b s e n c e D e h y d ra t io n G lu ta ra ld e h y d e
Figure 26. Influence of different treatments on the concentration of fructose (g.L-1
) formed from inulin hydrolysis.
Each assay was performed at different temperatures 50ºC, 55ºC, 60ºC and 65ºC. Chitosan beads stabilized in
acetate buffer (absence), dehydrated in desiccator during 15 minutes or reinforced with 500 µL of glutaraldehyde
during 1 hour were studied. Bioconversion runs were performed in 5% (w/v) inulin in acetate buffer at pH 4.5.
100 mg of immobilized inulinase was used as biocatalyst. Samples (10 µL) were collected at 2.5, 3.5, 6.5 and 24
hours after the beginning of incubation. Standard deviation did not exceed 5%.
For all temperature analyzed, the final concentration of fructose was higher in the case of beads
reinforced with glutaraldehyde (49.0 0.005 g.L-1
at 55ºC and 60ºC), whereas the lowest
concentration was observed in the case of beads stabilized in buffer (27.0 0.005 g.L-1
at 65ºC).
Regarding the beads stabilized in acetate buffer, the interaction between the cross-linker (TPP)
and chitosan chains is purely ionic and electrostatic. Chitosan is polycationic in acidic media (pKa 6.5)
and its amino groups can interact with negatively charged groups in TPP. Therefore, this reaction
depends strongly on the associated pH. Wentworth et al. (2004) stated an optimal pH between 6 to 7
for -galactosidase entrapped in chitosan cross-linked with TPP [119].
49
Nevertheless, the use of TPP as cross-linker failed to prove effective enough to achieve good
particle stability. Hence, some treatments after immobilization procedure were performed. One of the
most common is the use of glutaraldehyde [123]. Glutaraldehyde reacts with the hydrogels structure
composed by chitosan-TPP and, making it more rigid and stable. Also, the enzyme already entrapped
inside the matrix can react with glutaraldehyde. Altun and Cetinus (2007) reported a higher stability of
chitosan beads after exposed to glutaraldehyde solution as a complementary stability curing
treatment [123].
Additionally, the dehydration treatment may promote the formation hydrogen bonds among
chains. This type of cross-linking is physical, whereas glutaraldehyde promotes a chemical cross-
linking, which is stronger.
The effect of temperature on each type of treatment is also important to evaluate. In the
absence of treatment, and neglecting the temperature of 55°C, it was observed that exits a
relationship between temperature and final product concentration. Since for the lowest temperature,
the concentration of final product was higher. At 65°C, this effect was even more evidenced. The
same observation could be done for dehydrated chitosan beads, since the higher concentration of
fructose was achieved at 50ºC (ignoring the curve at 55ºC). These results suggest that for beads with
no treatment or dehydrated, the increasing in temperature affects the enzyme activity and, therefore
the concentration of fructose.
Nonetheless, for the treatment with glutaraldehyde the final product concentration is almost the
same for all temperatures analyzed. This result highlights that treatment with glutaraldehyde promotes
a more controlled environment in the matrix.
Also, chitosan beads appear to have good stability until 65ºC, since particles did not dissolve
and the size was similar to the fresh samples. To corroborate this, no trace of proteins was detected
by the Bradford method.
Besides the difference in matrix composition, no significant difference in fructose concentration
over 24 hours at 65ºC was observed between inulinase immobilized in CS beads reinforced with GA
(46.5 0.005 g.L-1
) and PVA film of 50 kDa with 400 µL of GA (44.0 0.005 g.L-1
).
In conclusion and considering the highest hydrolytic capacity of inulinase immobilized in CS
beads reinforced with GA, this procedure/matrix was chosen to detail characterization.
In order to improve the results, an optimization of immobilization conditions could be performed
(viz. immobilization temperature, dehydration period, volume of glutaraldehyde). Additionally, other
types of cross-linkers could be used, such as glucomannan and others type of aldehydes.
50
Second part
After the selection of the most promising inulinase immobilization procedures, a detail
characterization of those biocatalysts was performed (Figure 27). Henceforth the matrices will only be
denoted as PVA film or CS beads, corresponding to the selected ones.
Figure 27. Studies performed on PVA film immobilized inulinase and CS beads immobilized inulinase.
IV.3. Immobilization parameters
The feasibility of the proposed methodology for enzyme immobilization was also assessed by
evaluating the immobilization yield and efficiency and protein entrapment for the immobilization
procedures previously selected (Table 10).
The immobilization yield reported for Lentikats or PVA capsules reticulated with 1%(v/v) GA
were similar to the one obtained herein, being a satisfactory result [98][99]. Also, is important to
highlight that the difference in the shape of the matrix between Lentikats and PVA films produced,
seemed not affect the immobilization yield. The percentage of protein entrapped is also a good
achievement. The optimized chitosan immobilization protocol performed herein obtained a higher
value of protein entrapment than the original protocol (59%) [122]. The fact that the immobilization
efficiency achieved a value slightly higher than the protein entrapment may be a consequence of the
inulinase leakage but also due to the lack of specificity of the Bradford method. Fructanase Mixture
also contains other enzymes with low MW which are susceptible to leakage.
Table 10. Immobilization of inulinase in PVA and CS based matrices. The immobilization parameters were
determined as described in Methods section.
Matrix Protein
entrapment (%)
Immobilization
efficiency (%)
Immobilization
yield (%)
CS 96 4 98 6 29 3
PVA 94 5 97 3 25 4
PVA film of 50 kDa with
400 µL of GA
CS beads reinforced
with GA
Inulinase immobilization
procedures selected for
detailed characterization
Immobilization parameters
Temperature and pH profiles
Kinetics characterization
Thermal, storage and operational stabilities
51
M L1 L2 L3 L4
250
150
100
75
50
37
25
20
kDa
Enzyme leakage through the pores of the matrices is a consequence of the low molecular
weight of inulinases from Aspergillus sp. (in the range of 54 – 78 kDa) [192]. The entrapment of
enzymes in hydrogels is often characterized by some diffusion of the biocatalyst from the support,
particularly for enzymes with molecular weight less than 300 kDa [168].
The SDS-PAGE analysis revealed the presence of a more evident band in lane 3 between 50
and 70 kDa, suggesting approximately 60 kDa as molecular weight for inulinase. Since the
commercial inulinase studied is a combination of endo and exo-inulinase, it was expected to see more
than one band. This may be a consequence of the similar molecular weight of both inulinases. Kumar
et al. (2011) reported a molecular weight of 63.8 kDa for exo-inulinase from Aspergillus niger [193],
whereas a molecular weight between 69 or 64 kDa was stated for endo-inulinase from Aspergillus
niger [194].
Figure 28. Coomassie Blue stained SDS-PAGE gel of
inulinase samples. Lane M - Precision Plus Protein™ Dual
Color Standard. Lanes 1 and 2 are duplicates of inulinase
samples from Fructanase Mixture. Lanes 3 and 4 are
duplicates of inulinase 10-fold diluted in acetate buffer
100 mM pH 4.5.
IV.4. Effect of pH on the activity of free and immobilized inulinase
Immobilization procedures may impose shifts in optimal reaction conditions; consequently the
determination of pH and temperature profiles is of great importance.
Inulinases have an active site which contains imidazole and sulfhydryl groups. The hydrolysis of
inulin is catalyzed by the imidazole group by a nucleophile attack on carbon in the 2-position, yielding
a tetrahedral intermediate complex. The sulfhydryl group may act as hydrogen acceptor or donor so
the reaction medium pH plays a key role in the enzyme activity, as it is often found in
biocatalysis [155].
The effect of pH in the initial activity of free and immobilized inulinase was assessed at 50ºC, in
the pH range of 3.6 to 5.5 (Figure 29). Free inulinase showed its maximum activity at pH 4.5 whereas
the optimum pH of the immobilized enzyme in PVA film or CS beads was shifted to a less acidic pH
52
value of 5.0. For pH values below 4.5 no visible difference between free and immobilized forms was
observed, however the enhancement of activity following immobilization was observed only for higher
pH values (above 5.0). Therefore, immobilization slightly altered the enzymatic pH-activity profile, as
compared with the free form.
p H
Re
lati
ve
ac
tiv
ity
(%
)
3 .0 3 .5 4 .0 4 .5 5 .0 5 .5 6 .0
0 .0
2 0 .0
4 0 .0
6 0 .0
8 0 .0
1 0 0 .0
1 2 0 .0
F re e e n z y m e
C S b e a d s im m o b il iz e d e n z y m e
P V A f i lm im m o b il iz e d e n z y m e
Figure 29. pH profile of free inulinase and inulinase immobilized in CS beads and PVA film. Bioconversion runs
were performed in 5% (w/v) inulin solution at 50ºC at different pH values (3.6 – 5.5). 60 mg of immobilized
inulinase in CS beads, 100 mg of immobilized inulinase in PVA film or 50 µL of a 100–fold diluted preparation of
free inulinase were used as biocatalyst.
Inulinases from Aspergillus niger are stable within a pH range of 3.5–8.5 [155], thus the
optimum pH value achieved is in accordance with literature [155], [195]. The observed shift of
optimum pH to more basic value has been reported for other immobilized inulinases [170] [171].
Such pH shift upon immobilization (0.5 pH units variation), either in PVA or CS, are suggested
to be because of secondary interactions between the inulinase and the matrix, which could lead to
conformational changes in inulinase [196]. More specific, glutaraldehyde displays conformational
changes depending on the pH of the aqueous solution, suggesting different modes of cross-linking the
amino groups in enzymes and the functional groups in matrices [197]. Moreover, negatively charged
groups of the matrix, namely OH- from sodium tripolyphosphate, will tend to concentrate protons
(lowering the pH) around the enzyme. Therefore, the pH around the enzyme will be lower than that of
the bulk phase from which the measurement of pH is carried out. Boliver et al. (2013) have reviewed
some methodologies to measure more precisely the pH inside the matrix, giving a powerful tool to
understand how the pH is affected by immobilization [24].
Although many authors reported an acidic shift in the pH optimum upon immobilization in
chitosan matrices [122], [174], [198], [199], Yao et al. (2014) observed a basic shift upon
immobilization of tannase in chitosan beads cross-linked with GA [200]. Nguyen et al. (2011) also
reported that the optimum pH of endo-inulinase from Aspergillus niger was shifted to a more basic
value after immobilization in chitosan [171].
53
Concerning PVA immobilized inulinase, Anes and Fernandes (2013) reported no shift in pH
upon immobilization of inulinase mixture in PVA particles using Lentikat liquid [98]. Similar patterns,
where optimum pH profile is not significantly altered with immobilization, were reported previously by
Cattorini et al. (2009) [99] and Fernandes et al. (2009) [27].
IV.5. Effect of temperature on the activity of free and immobilized inulinase
The reaction temperature is a major factor to take into consideration in biocatalysis. Enzymes
are known to catalyze reactions in a specific range of temperature. Above or below a given
temperature value there is a progressive loss of activity due to physical changes in hydrogen bond
(weakest interactions), resulting thus in loss of the enzyme three-dimensional structure affecting,
therefore, the enzyme activity [3].
The influence of temperature on the initial activity of free and immobilized inulinase was
evaluated at pH 4.5, in the range of 45°C to 70°C (Figure 30). In this study, the optimum temperature
for the immobilized enzyme increased to 65°C for CS immobilized enzyme and 55ºC for PVA
immobilized enzyme, compared to 50°C for the free enzyme. Therefore, the enzymatic temperature-
activity profile displayed great differences with the immobilization.
T e m p e r a tu r e ( C )
Re
lati
ve
ac
tiv
ity
(%
)
4 0 .0 4 5 .0 5 0 .0 5 5 .0 6 0 .0 6 5 .0 7 0 .0 7 5 .0
0 .0
2 0 .0
4 0 .0
6 0 .0
8 0 .0
1 0 0 .0
1 2 0 .0
F re e e n z y m e
C S b e a d s im m o b il iz e d e n z y m e
P V A f i lm im m o b il iz e d e n z y m e
Figure 30. Temperature profile of free inulinase and inulinase immobilized in CS beads and PVA film.
Bioconversion runs were performed in 5% (w/v) inulin solution at pH 4.5 at different temperatures (45ºC – 70ºC).
60 mg of immobilized inulinase in CS beads, 100 mg of immobilized inulinase in PVA film or 50 µL of a 100–fold
diluted preparation of free inulinase were used as biocatalyst.
According to the literature, the optimum temperatures for free inulinase could range from 30ºC
to 60ºC and they are generally higher for bacteria and yeasts than for fungi [159]. The inulinase
optimum temperature obtained herein is consistent with reports by other authors [192], [201].
54
The temperature shift of 5ºC and 15ºC, when compared to the optimal temperature observed for
the free enzyme, could be due to the formation of a molecular cage around the enzyme, which
protected the enzyme molecules from the temperature, hence denaturation. For industrial application,
a relatively high temperature of 65°C (CS immobilization) is preferably used to avoid microbial
contamination, and it permits a greater concentration of the sugars, an ideal condition for
fructooligosaccharide production by inulinases [202].
Additionally, chitosan beads are robust even at 80ºC unlike some other synthetic hydrogels e.g.
PVA (Lentikat liquid) which lack physical stability even at 60ºC [98], [99].
Recently, Paripoorani et al. (2015) reported an optimum temperature of 70ºC when using
soluble inulinase in magnetite nanoparticles entrapped in chitosan [203]. Yewale et al. (2013) found
60°C as the best temperature for the immobilized inulinase on chitosan [174].
IV.6. Mass of support
The effect of the matrix weight in the initial activity of immobilized enzyme was evaluated by
using a matrix weight ranging from 40 mg to 200 mg (Figure 31). Assuming a uniform distribution of
the enzyme among immobilization matrix, a higher matrix weight is directly associated with a higher
amount of enzyme immobilized. Therefore is important to determine the range in which this effect
could be observed. In the case of CS, a linear effect was detected for beads with weights ranging from
40 mg to 80 mg, whereas in PVA matrix this effect was observed for weights ranging from 40 mg to
120 mg. Therefore, matrix weights inside these ranging were chosen to bioconversion studies.
M a t r ix w e ig h t ( m g )
Re
lati
ve
ac
tiv
ity
(%
)
0 .0 5 0 .0 1 0 0 .0 1 5 0 .0 2 0 0 .0 2 5 0 .0
0 .0
2 0 .0
4 0 .0
6 0 .0
8 0 .0
1 0 0 .0
1 2 0 .0
C S b e a d s im m o b il iz e d e n z y m e
P V A f i lm im m o b il iz e d e n z y m e
Figure 31. Influence of matrix weight on activity of immobilized enzyme, either in PVA film or CS beads.
Bioconversion runs were performed in 5% (w/v) inulin solution at pH 5.0 at 55ºC and 65ºC for PVA and CS
immobilization, respectively. Different matrix weight (40, 80, 120 and 200 mg) from both types of immobilization
were used as biocatalyst.
55
IV.7. Kinetic study of enzymatic inulin hydrolysis
Kinetics studies can be used to identify catalytic mechanisms, specifically the effect of substrate
caused by certain molecules. When immobilization is addressed, kinetic studies allow to further clarify
the effect of immobilization in enzyme performance. More specifically, they allow establishing whether
mass transfer limitation occurs and/or structural changes take place.
The trials for the determination of the kinetic constants (KM and Vmax), which fitted the enzymatic
hydrolysis of inulin, were performed at pH 4.5 and 50°C for free inulinase, at pH 5.0 and 55ºC for PVA
immobilized inulinase and at pH 5.0 and 65ºC for CS immobilized inulinase. The initial rates values
were converted into specific activities and, kinetic constants were obtained using Hyper32 software,
assuming Michaelis–Menten kinetics for inulin hydrolysis (either free or immobilized inulinase). This
assumption was previously demonstrated [3]. The Michaelis-Menten kinetics is giving by Equation 6:
𝑉 = 𝑉𝑚𝑎𝑥×[𝑆]
𝐾𝑀+[𝑆] (6)
where V is the initial reaction rate (gfructose.min-1
.mgenzyme-1
), [S] is the substrate concentration (g.L-1
),
Vmax is the maximum reaction rate achieved by the reaction (gfructose.min-1
.mgenzyme-1
) and KM is the
Michaelis constant (g.L-1
), which is the substrate concentration at which the reaction rate is half
of Vmax.
According to the results presented in Figure 32 and Table 11, a 1.5 and 1.6-fold increase in KM
was obtained for the immobilized inulinase in CS and PVA, respectively, when comparing with the free
inulinase. However, given the similarity of Vmax values for the free and immobilized biocatalyst in CS,
such immobilization effect is not considered significant. Nonetheless, a 2-fold decrease in Vmax was
outstanding for PVA immobilized inulinase.
In u l in c o n c e n t r a t io n (g .L-1
)
Sp
ec
ific
ac
tiv
ity
(g
fru
cto
se.m
in-1
.mg
en
zy
me
-1)
0 .0 2 0 .0 4 0 .0 6 0 .0 8 0 .0 1 0 0 .0 1 2 0 .0 1 4 0 .0 1 6 0 .0
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
F re e e n z y m e
C S b e a d s im m o b il iz e d e n z y m e
P V A f i lm im m o b il iz e d e n z y m e
Figure 32. Michaelis-Menten kinetics of inulin hydrolysis expressed by free and immobilized inulinase in PVA and
CS matrices. Bioconversion reactions were carried out with concentrations of inulin ranging from 15 g.L-1
to
150 g.L-1
in acetate buffer 100 mM pH 4.5 (for the free enzyme) or pH 5.0 (for the immobilized enzyme) at
optimum reaction conditions.
56
The increase in KM suggests less affinity of enzyme for inulin, which can be ascribed to the
impaired accessibility of the bulky inulin molecule to the active site of the immobilized enzyme, as a
result of structural changes of the enzyme upon immobilization or steric limitations created by the
immobilization matrix [14].
Comparing KM values between PVA and CS immobilization no difference was notable,
suggesting that KM was not affected by differences in polymers or matrix shape.
Lower value of Vmax observed after PVA immobilization may also result from loss of enzyme
active during immobilization procedure, caused by new interactions between enzyme and matrix [204].
The insignificant difference on Vmax value after immobilization in CS was not unusual, a behavior also
reported previously by Gill et al. (2006) [169].
The Michaelis constant of free inulinase from the commercial preparation Fructanase Mixture is
not reported in literature. However when comparing the results obtained with the free enzyme from
Fructozyme L, the KM achieved is lower [170][27][99][205].
The increase in KM values and decrease in Vmax values is a commonly feature reported when
inulinase is entrapped in hydrogels for inulin hydrolysis. Anes and Fernandes (2013) reported also a
1.6-fold increase in KM when inulinase is entrapped in PVA lenses (Lentikats) cross-linked with GA
[98]. Moreover, a 1.8-fold increase was cited by Fernandes et al. (2009), when inulinase was
entrapped using PVA-based hydrogel particles [27]. Altun and Cetinus (2007) reported a 1.6-fold
increase in KM when chitosan beads immobilized pepsin [123]. Yewale et al. (2013) entrapped
inulinase in chitosan beads, however no significant variation on KM and Vmax was observed [174].
Table 11. Kinetics constants obtained for inulin hydrolysis with free and immobilized inulinase. Data were
processed through Hyper32 software.
Enzyme Vmax
(gfructose.min-1
.mgenzyme-1
)
KM
(g.L-1
) r
2
Free inulinase 1.1 ± 0.1 12.8 ± 4.8 0.92
CS beads immobilized inulinase 1.0 ± 0.1 19.6 ± 6.9 0.88
PVA film immobilized inulinase 0.57 ± 0.04 20.4 ± 4.1 0.78
IV.8. Stability of immobilized enzyme
High thermal, storage and operational stabilities of an immobilized enzyme are the most
importance for the economic viability of a biosynthetic process, hence these issues were addressed.
57
IV.8.1. Thermal stability
Thermal deactivation of the biocatalyst is one of the major causes of activity decay. Therefore,
the stability of the enzyme was evaluated at four different incubation temperatures (40ºC, 50ºC, 60ºC
and 70ºC) and during long time periods (48 hours for the 40C assay; 30 hours for the 50C assay; 10
hours for the 60C and 5 hours for 70ºC). This enables to determine the deactivation constants of free
and immobilized biocatalyst and establish if immobilization improved thermal stability.
Through the analysis of Figure 33 is possible to conclude that free inulinase is very susceptible
to high temperatures, since its relative activity was 43% at 50ºC and 14% at 70ºC, suggesting intrinsic
thermal deactivation.
When the inulinase is entrapped in PVA or CS hydrogels, the thermal stability was improved
(Figure 34). At 50ºC around 71% of relative activity was observed for either immobilization. For
temperatures between 40ºC and 60ºC no significant differences in relative activity of both
immobilizations, at the end of the assay, were observed. However, at 70ºC a pronounced decay in the
activity of PVA immobilized inulinase was observed where 70% of the initial activity was lost. Below
70ºC no visible structural changes in the films or beads were observed, however at 70ºC the PVA film
melting becomes noticeable. This melting behavior occurred at a higher temperature than Lentikats,
which start to melt at 55ºC [99]. In the case of CS immobilization, the relative activity remained above
50% even at 70ºC. This result is higher than the one reported by Yewale et al. (20% of residual activity
at 70ºC) [174]. Hence, the activity of immobilized inulinase in PVA or CS displayed an improved
thermal stability when compared to free enzyme.
Overall, it is possible to observe that the decay in activity is more pronounced at 60ºC and 70ºC,
for each studied biocatalyst. Moreover, the immobilization had improved the thermal deactivation
profile for all temperatures. Also, it is interesting to note that for temperatures 40ºC and 50ºC the CS
immobilization exhibited decay in initial activity values and kept constant until the end of assay. For
free or immobilized in PVA, the activity tended to decay along time.
Additionally, no protein loss was noticed using the Bradford method in all the assays.
Several models have been developed in order to explain the enzymatic deactivation. Table 12
represents the most common deactivation models and equations for the determination of the half time.
The half-life time is the time required for the enzymatic activity decrease to one half of its initial value.
58
Table 12. Deactivation models and half-time equations. Where K(t) stands for activity at a given time, Ki is the
initial activity, Kd is the deactivation rate constant and t1/2 is the half time, A stands for a complex function of
individual rate constants, and α and β are apparent first-order rate constants. Adapted from [188][14].
Deactivation models Half-time
Exponential deactivation model 𝐾 (𝑡) = 𝐾𝑖 × 𝑒(−𝐾𝑑×𝑡)
(8)
𝑡21 =
ln 2
𝐾𝑑
(9)
Linear inverted model 𝐾(𝑡) =
𝐾𝑖
1 + (𝐾𝑑 × 𝑡)
(10)
𝑡21 =
1
𝐾𝑑
(11)
Three-parameter bi-exponential deactivation model
𝐾 (𝑡) = 𝐾𝑖 [𝐴𝑒(−𝛼𝑡) + (1 − 𝐴)𝑒(−𝛽𝑡)] (12)
-
The deactivation models presented above were applied to the experimental data obtained for
thermal deactivation studies. Taking into consideration the correlation factor, the three-parameter bi-
exponential model was the one that provided the best fit for experimental data. This model was
developed by Aymard et al. (2000) and is characterized by being independent of the enzyme
mechanism [188]. The fit of three-parameter bi-exponential model to the experimental data is
represented as lines in Figure 33 and Figure 34.
T im e (h o u r )
Re
lativ
e s
pe
cif
ic a
cti
vit
y
0 .0 2 0 .0 4 0 .0 6 0 .0
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
4 0 º C 5 0 º C 6 0 º C 7 0 º C
F r e e e n z y m e
Figure 33. Thermal deactivation profile for free inulinase at 40ºC, 50ºC, 60ºC and 70ºC incubation temperatures.
The lines represent the trend of the three-parameter bi-exponential model. Bioconversion runs for initial activity
were performed in 5% (w/v) inulin solution 100 mM pH 4.5 with 50 µL of 100-fold diluted enzyme solution in
acetate buffer 100 mM pH 4.5.
59
T im e (h o u r )
Re
lativ
e s
pe
cif
ic a
cti
vit
y
0 .0 2 0 .0 4 0 .0 6 0 .0
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
4 0 º C 5 0 º C 6 0 º C 7 0 º C
C S b e a d s
T im e (h o u r )
Re
lativ
e s
pe
cif
ic a
cti
vit
y
0 .0 2 0 .0 4 0 .0 6 0 .0
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
4 0 º C 5 0 º C 6 0 º C 7 0 º C
P V A f i lm
Figure 34. Thermal deactivation profile for immobilized inulinase in CS beads (A) or in PVA film (B) at 40ºC,
50ºC, 60ºC and 70ºC incubation temperatures. The lines represent the trend of the three-parameter bi-
exponential model. Bioconversion runs for initial activity were performed in 5% (w/v) inulin solution 100 mM
pH 5.0 at optimum conditions. 60 mg of CS beads or 100 mg of PVA film were used as biocatalyst.
The constants rates were determined using ‘‘Solver’’ add-in from Excel for Windows by
minimizing the residual sum of squares between the experimental data points and those estimated by
the respective model. Half times were predicted by knowing the constants rates of Equation 12 and
assuming K(t)
Ki= 0.5. All constants and half times are given in Table 13. The correlation coefficient, r,
was calculated using a Pearson's correlation according to previous publications [206].
A
B
60
Table 13. Calculated parameters for the thermal deactivation of free and immobilized inulinase using a three-
parameter bi-exponential model.
Enzyme Temperature
(ºC) A (-) (h
-1) (h
-1) r t1/2 (h)
Free
40 0.89 1.16×10-2
9.15 0.972 49
50 0.88 2.00×10-2
9.15 0.987 28
60 0.73 1.09×10-1
9.15 0.995 3.5
70 1.13 5.15×10-1
16.63 0.997 1.5
Immobilized in CS
40 0.89 2.00×10-3
17.23 0.901 285
50 0.74 3.09×10-3
17.23 0.920 124
60 0.92 4.13×10-2
17.23 0.994 15
70 0.73 8.37×10-2
17.23 0.997 5
Immobilized in PVA
40 0.95 4.18×10-3
16.85 0.999 155
50 0.98 1.22×10-2
16.85 0.982 54
60 0.95 4.86×10-2
16.85 0.964 13
70 0.82 0.29 16.85 0.966 1.7
The half time values corroborate what mentioned before. First observation is that for each type
of biocatalyst the half time values decrease with temperature increase. Second observation is that the
highest half time values are displayed for inulinase immobilized in CS and the lowest for the free
inulinase. Thus, immobilization in CS enhances the half-time of the biocatalyst as much as (roughly)
5.8-fold (at 40ºC) as compared to the free form, whereas PVA immobilization exhibited 3.2-fold
increase.
The half times are dependent of the type of enzyme and the shift in their values is also
dependent to the type of matrix used. Since no publications regarding half time of Fructanase Mixture
were reported, a comparison to other inulinases was made.
Anes and Fernandes (2013) were able to immobilized inulinase from Fructozyme L in PVA
lenses (Lentikats) with a half time of 50 hours at 40ºC, 23 hours at 50ºC and 0.52 hours at 60ºC [98].
Catana et al. (2007) immobilized inulinase from Fructozyme L in Amberlite IRC 50® with a half-life of
504 hours at 40°C, 134.4 hours at 50°C and 14.6 h at 60°C [170]. Gupta et al. (1992) immobilized
inulinase from Fusarium oxysporum on DEAE-cellulose and achieved a half-life of 45 min at 50ºC.
Therefore, the immobilized inulinase in PVA film proved to have better stability than those where
DEAE-cellulose and aminoethylcellulose was used as a matrix. However when compared with the
results with Amberlite IRC 50®, the PVA matrix prove to be less efficient in maintaining the inulinase
stability at different temperatures.
IV.8.2. Storage stability
Storage stability of an enzyme is of significant importance for correctly scheduling a production
batch of the immobilized biocatalyst aiming at its application in given reaction runs along time. During
storage of immobilized biocatalyst in solution the enzyme activity may decay along time [27]. This
behavior can be ascribed to the decay of the intrinsic activity to the biocatalyst or to enzyme leakage.
An exception was reported by Nunes et al. (2012), where 1.5-fold increase in initial activity was
observed.
61
The storage stability of free and immobilized biocatalyst, stored at 4ºC in acetate buffer 100mM
pH 4.5 or 5.0 during 180 days, was assessed. Again, the three-parameter bi-exponential model
(Equation 12) was used to predict the deactivation behavior under storage of each immobilized form of
inulinase (Figure 35). The data estimated for the parameters of the bi-exponential model, as well as
the predicted half time, are presented in Table 14.
T im e ( d a y )
Re
lativ
e s
pe
cif
ic a
cti
vit
y
0 .0 5 0 .0 1 0 0 .0 1 5 0 .0 2 0 0 .0
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
F re e e n z y m e
C S b e a d s im m o b il iz e d e n z y m e
P V A f i lm im m o b il iz e d e n z y m e
Figure 35. Storage deactivation profile predicted through three-parameter bi-exponential model for storage
stability of free and immobilized biocatalyst. The lines represent the trend of the three-parameter bi-exponential
model. Bioconversion runs were performed with 50 µL of 100-fold enzyme preparation in acetate buffer 100 mM
pH 4.5, 60 mg of CS beads or 100 mg of PVA film. The reactions were performed in 5% (v/w) inulin solution
100 mM at optimum conditions.
Regarding the results obtained for the free enzyme, the activity decays to values around 70%
after 60 days and seemed to be stable until 180 days of storage. This initial activity decay may be
caused by enzyme adaptations to the storage environment. Nonetheless, in the immobilized inulinase
the initial decay was not so noticeable, may be due to the matrix protection. CS immobilization
retained about 80% of the initial activity, whereas the immobilized inulinase in PVA retained about
76%, which is a promising result. Furthermore, no protein was detected during the storage time and
no contamination was observed.
Table 14. Rate constants for immobilized inulinase in both forms and free enzyme for storage deactivation using
a three-parameter bi-exponential equation.
Enzyme A (-) (h-1
) (h-1
) r t1/2 (days)
Free 0.76 5.61×10-4
16.83 0.998 740
Immobilized in CS 0.94 9.01×10-4
16.83 0.994 700
Immobilized in PVA 0.87 8.00×10-4
16.83 0.991 685
62
Taking into consideration the half time of free (740 days) and immobilized enzyme (700 and 685
days), it seems that immobilization may not have a direct effect in the storage stability for long term
studies. Nonetheless, it is important to highlight that the enzyme used was a commercial preparation
supplied with a stabilized solution containing 50% glycerol and 0.02% sodium azide (from
https://secure.megazyme.com/Fructanase-Mixture-purified-liquid). Hence, is difficult to guarantee that
the data obtained from the immobilized biocatalysts were either due to the immobilization or due to the
enzyme stabilizing agents. To corroborate this observation, when endoinulinase produced from
Aspergillus niger AUMC 9375 (without addition of stabilizing agents) a complete loss of activity for the
free enzyme after 24 days was observed [192]. A further study must be prepared with produced
inulinase in order to determine the effect of the PVA and CS in storage stability.
The storage stability half time for this particular enzymatic commercial preparation is not
reported in literature. Nonetheless, Karimi et al. (2014) cited 65.2 days for free endo-inulinase from
Aspergillus niger, assuming the exponential deactivation model [207]. Anes and Fernandes (2013)
reported a storage stability of inulinase from Fructozyme L in PVA lenses (Lentikats) with a half time
of 238 days [98].
IV.8.3. Operational stability
The operational stability of the immobilized enzyme is the main characteristic that limits a
potential application of immobilizates in industrial scale. A final study was performed to evaluate the
operational stability of both immobilized systems (PVA and CS), under continuous operation during 47
and 31 days.
According to Lilly et al. (1966), and assuming a piston flow through the reactor the residence
time depends on the void volume and flow rate used [208]. Therefore, residence time (𝜏) was
calculated according to Equation 13.
τ = V
F (13)
where F is the flow rate (mL.min-1
) and V is the void volume (mL).
Also, volumetric productivity (Qp) was calculated according to Singh et al. (2008), as follows:
Qp = Cp ×F
V (14)
where Cp is the concentration of fructose produced (g.L-1
), F is the flow rate (mL.min-1
) and V is the
void volume (mL).
63
From previous publications the optimum pH remained the constant after application in
continuous operation, in the present work the pH established remained at 5.0. Regarding temperature,
it was a choice not to perform the continuous operation at optimum temperatures.
Continuous production of fructose using CS immobilized inulinase
The feasibility of using inulinase immobilized in CS reinforced with GA for full inulin hydrolysis
under continuous flow was then assessed. The reaction took place for 47 days at 55°C where
5% (w/v) inulin solution pH 5.0 was being fed to the reactor at a flow rate of 0.0053 mL.min-1
and DNS
analysis were performed on a daily basis for fructose quantification. The reactor had a void volume of
0.78 mL and the correspondent residence time was 147 minutes approximately.
Figure 36. Reactor filled with CS beads used for continuous-flow operation (A). Appearance of the beads after
continuous operation (B).
Figure 38 displayed encouraging operational stability during the 47 days of operation, since the
product yield was not under 90% during the first twenty four days and a final product yield of 91% was
achieved. The initial volumetric activity was 22.6 g.L-1
.h-1
. Regarding deactivation constant and half
time, no deactivation model fit to the experimental data. Therefore more prolonged operation would be
needed to provide experimental data to suggest a trend. Product yield (%) seemed to remain roughly
constant along time.
The results suggest that the set-up presents a promising tool for the continuous productions of
fructose. Results were very similar to the ones recently presented by Yewale et al. (2013). These
authors operated a packed bed reactor with inulinase immobilized on chitosan for the continuous
hydrolysis of inulin at 60ºC and were able to maintain full conversion for 6 days but only for a 1% (w/v)
inulin solution. No publication was found regarding continuous operation of inulinase immobilized in
CS during 47 days.
A B
64
Continuous production of fructose using PVA immobilized inulinase
The feasibility of using inulinase immobilized in PVA film for full inulin hydrolysis under
continuous flow was also assessed. The reaction took place for 31 days at 50°C where 5% (w/v)
inulin solution pH 5.0 was being fed to the reactor at a flow rate of 0.0053 mL.min-1
and DNS analysis
were performed on a daily basis for fructose quantification. The void volume was 0.87 mL and the
correspondent residence time of 164 minutes approximately.
Figure 37. Reactor filled with PVA film used for continuous-flow operation (A). Appearance of the film used after
continuous operation. Note that the film was cut into cubes before implemented in the reactor (B).
Operational stability for PVA immobilized inulinase exhibits promising results, since the product
yield was not under 90% during the first eight days and a final product yield of 63% was achieved
(Figure 39). The initial volumetric activity was 18.3 g.L-1
.h-1
. Using as reference a linear inverted model
for enzyme decay (Equation 10), a deactivation constant of 0.02 day-1
and a half time of 49 days were
calculated.
One publication which reports the continuous production of fructose with inulinase
(Fructozyme L) immobilized in PVA (Lentikats) crosslinking with 1% (v/v) glutaraldehyde cited a
product yield of 75% over 20 days of hydrolysis reaction. This result is comparable to the one
achieved in this work, 72% at day twenty.
Figueira et al. (2013) reported the continuous operation of β-galactosidase entrapped in PVA
lenses (Lentikats®) during 100 hours (4 days) at 50ºC with a final conversion of 75% [209].
Rebroš et al. (2006) studied the continuous operation of glucoamylase into Lentikats® at 45ºC
during 63 days with 60% of conversion with no drop in enzyme activity [210].
A B
65
Figure 38. Operational stability at the packed bed reactor for inulin hydrolysis, based on the relative product yield. The 1.206 g of immobilized inulinase in CS were used for the
hydrolysis of 5% (w/v) inulin solution in acetate buffer 100 mM pH 5.0 at 55°C. At day one the initial concentration of fructose was 55.5 ± 0.37 g.L-1
, with volumetric productivity
of 22.6 g.L-1
.h-1
. Moreover, no deactivation model fit to the experimental data.
Figure 39. Operational stability at the packed bed reactor for inulin hydrolysis, based on the relative product yield. Predicted values (red line) were estimated assuming a linear
inverted model. The 3.450 g of inulinase in immobilized in PVA were used for the hydrolysis of 5% (w/v) inulin solution in acetate buffer 100 mM pH 5.0 at 50°C. At day one the
initial concentration of fructose was 50.0 ± 2.5 g.L-1
, with volumetric productivity of 18.3 g.L-1
.h-1
.
0
20
40
60
80
100
120
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47
Pro
du
ct
yie
ld (
%)
Time (day)
0
20
40
60
80
100
120
1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31
Pro
du
ct
yie
ld (
%)
Time (day)
66
67
Chapter V. Conclusions and Future Work
Hydrogels are being employed in innumerous fields, namely in enzyme immobilization. Polyvinyl
alcohol and chitosan are synthetic and natural polymers, respectively used for the production of
hydrogels due to their hydrophilicity, biocompatibility and biodegradability.
Inulinase from Fructanase Mixture was efficiently immobilized on chitosan beads and polyvinyl
alcohol. The best compromise between polyvinyl alcohol molecular weight and volume of cross-linker
was achieved, with the film of 50 kDa and 400 µL of glutaraldehyde, yielding 97% of immobilization
efficiency. The chitosan beads reinforced with glutaraldehyde were the ones with more stability under
a range of different temperatures, with an immobilization efficiency of 98%.
Free inulinase displays an optimum pH at 4.5 and an optimum temperature of 50ºC. Shifts of
the optimum pH and temperature to 5.0 and to 65ºC or 55ºC were observed for enzyme immobilized in
chitosan or polyvinyl alcohol, respectively, were assayed.
The KM values of free and immobilized inulinase for inulin were 13 g.L-1
and 20 g.L-1
,
approximately suggesting lower affinity towards substrate. No difference was detected of the Vmax
value concerning immobilization in chitosan, whereas a 2-fold decrease was detected for polyvinyl
alcohol immobilization.
Thermal stability was enhanced after immobilization on both matrices. This improvement was
mostly noticed at high temperatures. Nonetheless, immobilization seemed not affect storage stability
during the 180 days of storage.
Operational stability highlights the potentials of the matrices selected, since a product yield of
91% was achieved for inulinase immobilized in chitosan, after 47 days at 55ºC. Also, the continuous
operation of inulinase immobilized in polyvinyl alcohol obtained 63% of product yield, after 31 days at
50ºC. In the case of chitosan no deactivation model was possible to fit to the experimental data.
In conclusion, the aim of this master’s thesis was accomplished, since relatively simple, robust
and low-cost immobilization procedures were established.
The results present in this project are a significant achievement for the field of catalytic
hydrogels. The commercial enzyme preparation has never been reported in immobilization studies
and thus, its characterization was an asset to the field.
Concerning the immobilization in PVA films, it was performed at pH more basic (4.5) than the
commonly published (pH 1 or 2) being an improvement and an advantage for enzymes sensible to
acidic pH values. Also, the fact that the matrix shape (film form) is different from the usual (lenses) and
it is an open field of research.
68
Regarding chitosan immobilization, the optimum temperature reported was with no doubt a
great achievement. Additionally, the improvement in thermal stability was notable. Finally, the
continuous operation with inulinase immobilized in chitosan revealed no deactivation pattern.
As future work it is proposed the automation process for beads formation and films in order to
have all matrices with the same dimensions and thus more accurate results. Regarding each matrix,
different diameters, heights and shapes could also be a topic of research. Concerning immobilization
technique, other types of cross-linkers and polymer interactions could be studied.
A more precise protein quantification method is also of great importance. A simple methodology
to verify the pH inside the matrix could give some insights about the effect of immobilization on pH
profile.
It will be interesting to apply the immobilization techniques established for other protein model
systems, in order to observe if the trend in stability assays and kinetic characterization remain.
On the subject of reactor, different configurations are also suggested to analyze in order to
obtain higher product yields as much time as possible.
The use of non-commercial enzyme preparations is also of relevance. A sample without
stabilizing agents will probably have different performances when storage stability is studied.
69
References
[1] N. Price and L. Stevens, Fundamentals of Enzymology: The Cell and Molecular Biology of
Catalytic Proteins, 3rd ed. Oxford: Oxford University Press, 1999.
[2] K. Tipton and S. Boyce, “History of the enzyme nomenclature system,” Bioinformatics, vol. 16,
no. 1, pp. 34–40, Jan. 2000.
[3] M. M. Cox and D. Nelson, Principles of Biochemistry, 5th ed. New York: W. H. Freeman and
Company, 2008.
[4] A. Illanes, Enzyme Biocatalysis: Principles and Applications. Dordrecht: Springer, 2008.
[5] P. Binod, P. Palkhiwala, R. Gaikaiwari, K. M. Nampoothiri, A. Duggal, K. Dey, and A. Pandey,
“Industrial Enzymes - Present status and future perspectives for India,” J. Sci. Ind. Res.
(India)., vol. 72, no. May, pp. 271–286, 2013.
[6] M. Ayala and E. Torres, “Enzymatic activation of alkanes: constraints and prospective,” Appl.
Catal. A Gen., vol. 272, no. 1–2, pp. 1–13, Sep. 2004.
[7] A. Illanes, A. Cauerhff, L. Wilson, and G. R. Castro, “Recent trends in biocatalysis
engineering,” Bioresour. Technol., vol. 115, pp. 48–57, Jul. 2012.
[8] L. Cao, Carrier-bond Immobilized Enzymes: Principles, Applications and Design, 1st ed. Wiley-
VCH Verlag GmbH & Co KGaA, 2005.
[9] D. Wilkinson, N. Akumanyi, R. Hurtado-Guerrero, H. Dawkes, P. F. Knowles, S. E. V Phillips,
and M. J. McPherson, “Structural and kinetic studies of a series of mutants of galactose
oxidase identified by directed evolution.,” Protein Eng. Des. Sel., vol. 17, no. 2, pp. 141–148,
2004.
[10] C. Q. Zhong, S. Song, N. Fang, X. Liang, H. Zhu, X. F. Tang, and B. Tang, “Improvement of
low-temperature caseinolytic activity of a thermophilic subtilase by directed evolution and site-
directed mutagenesis,” Biotechnol. Bioeng., vol. 104, no. 5, pp. 862–870, 2009.
[11] S. Lee, D. G. Lee, M. K. Jang, M. J. Jeon, H. J. Jang, and S. H. Lee, “Improvement in the
catalytic activity of β-agarase AgaA from Zobellia galactanivorans by site-directed
mutagenesis,” J. Microbiol. Biotechnol., vol. 21, no. 11, pp. 1116–1122, 2011.
[12] U. T. Bornscheuer, G. W. Huisman, R. J. Kazlauskas, S. Lutz, J. C. Moore, and K. Robins,
“Engineering the third wave of biocatalysis.,” Nature, vol. 485, no. 7397, pp. 185–94, May
2012.
[13] R. A. Sheldon, “Enzyme Immobilization: The Quest for Optimum Performance,” Adv. Synth.
Catal., vol. 349, no. 8–9, pp. 1289–1307, Jun. 2007.
[14] J. M. S. Cabral, M. R. Aires-Barros, and M. Gama, Engenharia Enzimática, 1st ed. Lisboa:
Lidel-Edições Técnicas Lda., 2003.
[15] J. M. Nelson and E. G. Griffin, “Adsorption of invertase,” J. Am. Chem. Soc., vol. 38, no. 5, pp.
1109–1115, May 1916.
[16] T. Tosa, T. Mori, N. Fuse, and I. Chibata, “Studies on continuous enzyme reactions. I.
Screening of carriers for preparation of water-insoluble aminoacylase.,” Enzymologia, vol. 31,
no. 4, pp. 214–24, Oct. 1966.
[17] W. Tischer and F. Wedekind, “Immobilized Enzymes : Methods and Applications,” Curr. Chem.,
vol. 200, pp. 95–126, 1999.
[18] D. Brady and J. Jordaan, “Advances in enzyme immobilisation.,” Biotechnol. Lett., vol. 31, no.
11, pp. 1639–50, Nov. 2009.
[19] R. Konwarh, N. Karak, S. K. Rai, and A. K. Mukherjee, “Polymer-assisted iron oxide magnetic
nanoparticle immobilized keratinase.,” Nanotechnology, vol. 20, no. 22, Jun. 2009.
[20] R. O. Cristóvão, S. C. Silvério, A. P. Tavares, A. I. Brígida, J. M. Loureiro, R. A. Boaventura, E.
A. Macedo, and M. A. Coelho, “Green coconut fiber: a novel carrier for the immobilization of
commercial laccase by covalent attachment for textile dyes decolourization.,” World J.
Microbiol. Biotechnol., vol. 28, no. 9, pp. 2827–38, Sep. 2012.
70
[21] N. Hashemifard, A. Mohsenifar, B. Ranjbar, A. Allameh, S. Lotfi, and B. Etemadikia,
“Fabrication and kinetic studies of a novel silver nanoparticles-glucose oxidase bioconjugate.,”
Anal. Chim. Acta, vol. 675, no. 2, pp. 181–4, Aug. 2010.
[22] F. Wang, R.-X. Su, W. Qi, M.-J. Zhang, and Z.-M. He, “Preparation and activity of bubbling-
immobilized cellobiase within chitosan-alginate composite.,” Prep. Biochem. Biotechnol., vol.
40, no. 1, pp. 57–64, Jan. 2010.
[23] B. Brena, P. González-Pombo, and F. Batista-Viera, “Immobilization of enzymes: a literature
survey.,” Methods Mol. Biol., vol. 1051, pp. 15–31, Jan. 2013.
[24] J. M. Bolivar, T. Consolati, T. Mayr, and B. Nidetzky, “Shine a light on immobilized enzymes:
real-time sensing in solid supported biocatalysts,” Trends Biotechnol., vol. 31, no. 3, pp. 194–
203, Mar. 2013.
[25] S. Nisha, A. K. S, and N. Gobi, “A Review on Methods, Application and Properties of
Immobilized Enzyme,” Chem. Sci. Reiew Lett., vol. 1, no. 3, pp. 148–155, 2012.
[26] S. Datta, L. R. Christena, and Y. R. S. Rajaram, “Enzyme immobilization: an overview on
techniques and support materials,” 3 Biotech, vol. 3, no. 1, pp. 1–9, Jun. 2012.
[27] P. Fernandes, M. P. C. Marques, F. Carvalho, and J. M. S. Cabral, “A simple method for
biocatalyst immobilization using PVA-based hydrogel particles,” J. Chem. Technol. Biotechnol.,
vol. 84, no. 4, pp. 561–564, Apr. 2009.
[28] M. M. M. Elnashar, “Review Article: Immobilized Molecules Using Biomaterials and
Nanobiotechnology,” J. Biomater. Nanobiotechnol., vol. 01, no. 01, pp. 61–77, 2010.
[29] K. Min and Y. J. Yoo, “Recent progress in nanobiocatalysis for enzyme immobilization and its
application,” Biotechnol. Bioprocess Eng., vol. 19, no. 4, pp. 553–567, Jul. 2014.
[30] J. Zhu and G. Sun, “Lipase immobilization on glutaraldehyde-activated nanofibrous
membranes for improved enzyme stabilities and activities,” React. Funct. Polym., vol. 72, no.
11, pp. 839–845, Nov. 2012.
[31] A. J. J. Straathof, S. Panke, and A. Schmid, “The production of fine chemicals by
biotransformations,” Curr. Opin. Biotechnol., vol. 13, pp. 548–556, 2002.
[32] A. Sassolas, L. J. Blum, and B. D. Leca-Bouvier, “Immobilization strategies to develop
enzymatic biosensors,” Biotechnol. Adv., vol. 30, no. 3, pp. 489–511, May 2012.
[33] A. Vasconcelos, C. J. S. M. Silva, M. Schroeder, G. M. Guebitz, and A. Cavaco-Paulo,
“Detergent Formulations for Wool Domestic Washings Containing Immobilized Enzymes,”
Biotechnol. Lett., vol. 28, no. 10, pp. 725–731, May 2006.
[34] J. C. Moreno-Pirajàn and L. Giraldo, “Study of immobilized candida rugosa lipase for biodiesel
fuel production from palm oil by flow microcalorimetry,” Arab. J. Chem., vol. 4, no. 1, pp. 55–62,
Jan. 2011.
[35] A. Dinçer and A. Telefoncu, “Improving the stability of cellulase by immobilization on modified
polyvinyl alcohol coated chitosan beads,” J. Mol. Catal. B Enzym., vol. 45, no. 1–2, pp. 10–14,
Apr. 2007.
[36] R. A. Messing and A. M. Filbert, “Immobilized glucose isomerase for the continuous conversion
of glucose to fructose,” J. Agric. Food Chem., vol. 23, no. 5, pp. 920–923, May 1975.
[37] Q. Husain, “β Galactosidases and their potential applications: a review,” Crit. Rev. Biotechnol.,
vol. 30, no. 1, pp. 41–62, Mar. 2010.
[38] J. Jeganathan, A. Bassi, and G. Nakhla, “Pre-treatment of high oil and grease pet food
industrial wastewaters using immobilized lipase hydrolyzation,” J. Hazard. Mater., vol. 137, no.
1, pp. 121–128, Sep. 2006.
[39] A. Illanes, L. Wilson, O. Corrotea, L. Tavernini, F. Zamorano, and C. Aguirre, “Synthesis of
cephalexin with immobilized penicillin acylase at very high substrate concentrations in fully
aqueous medium,” J. Mol. Catal. B Enzym., vol. 47, no. 1–2, pp. 72–78, Jun. 2007.
[40] I. Hoffmann, V. D. Silva, and M. da G. Nascimento, “Enantioselective resolution of (R,S)-1-
phenylethanol catalyzed by lipases immobilized in starch films,” J. Braz. Chem. Soc., vol. 22,
no. 8, pp. 1559–1567, Aug. 2011.
[41] P. Fernandes, “Miniaturization in biocatalysis.,” Int. J. Mol. Sci., vol. 11, no. 3, pp. 858–79, Jan.
2010.
71
[42] R. DiCosimo, J. McAuliffe, A. J. Poulose, and G. Bohlmann, “Industrial use of immobilized
enzymes,” Chem. Soc. Rev., vol. 42, no. 15, p. 6437, 2013.
[43] S. H. Hemmingsen, “Development of an Immobilized Glucose Isomerase for Industrial
Application,” in Applied Biochemistry and Bioengineering, Elsevier, 1979, pp. 157–183.
[44] S. Li, X. Yang, S. Yang, M. Zhu, and X. Wang, “Technology prospecting on enzymes:
application, marketing and engineering.,” Comput. Struct. Biotechnol. J., vol. 2, no. 3, Jan.
2012.
[45] S. C. Lee, I. K. Kwon, and K. Park, “Hydrogels for delivery of bioactive agents: a historical
perspective.,” Adv. Drug Deliv. Rev., vol. 65, no. 1, pp. 17–20, 2013.
[46] S. J. Buwalda, K. W. M. Boere, P. J. Dijkstra, J. Feijen, T. Vermonden, and W. E. Hennink,
“Hydrogels in a historical perspective: From simple networks to smart materials,” J. Control.
Release, vol. 190, pp. 254–273, Sep. 2014.
[47] M. H. Kim, S. An, K. Won, H. J. Kim, and S. H. Lee, “Entrapment of enzymes into cellulose–
biopolymer composite hydrogel beads using biocompatible ionic liquid,” J. Mol. Catal. B
Enzym., vol. 75, pp. 68–72, Mar. 2012.
[48] H. Park and K. Park, “Hydrogels in Bioapplications,” in Hydrogels and Biodegradable Polymers
for Bioapplications, no. 2, American Chemical Society, 1996, pp. 2–10.
[49] M. R. Guilherme, F. A. Aouada, A. R. Fajardo, A. F. Martins, A. T. Paulino, M. F. T. Davi, A. F.
Rubira, and E. C. Muniz, “Superabsorbent hydrogels based on polysaccharides for application
in agriculture as soil conditioner and nutrient carrier: A review,” Eur. Polym. J., Apr. 2015.
[50] S. K. H. Gulrez, S. Al-assaf, and G. O. Phillips, “Hydrogels : Methods of Preparation ,
Characterisation and Applications,” in Progress in Molecular and Environmental
Bioengineering, InTech, 2003, pp. 117–150.
[51] E. M. Ahmed, “Hydrogel: Preparation, characterization, and applications,” J. Adv. Res., Jul.
2013.
[52] F. Ullah, M. B. H. Othman, F. Javed, Z. Ahmad, and H. M. Akil, “Classification, processing and
application of hydrogels: A review,” Mater. Sci. Eng. C, vol. 57, pp. 414–433, Dec. 2015.
[53] R. Ulijn, N. Bibi, V. Jayawarna, P. Thornton, S. Todd, R. Mart, A. Smith, and J. Gough,
“Bioresponsive hydrogels,” Mater. Today, vol. 10, no. 4, pp. 40–48, 2007.
[54] E. Gil and S. Hudson, “Stimuli-reponsive polymers and their bioconjugates,” Prog. Polym. Sci.,
vol. 29, no. 12, pp. 1173–1222, Dec. 2004.
[55] T. R. Hoare and D. S. Kohane, “Hydrogels in drug delivery: Progress and challenges,” Polymer
(Guildf)., vol. 49, no. 8, pp. 1993–2007, Apr. 2008.
[56] M. Hamidi, A. Azadi, and P. Rafiei, “Hydrogel nanoparticles in drug delivery.,” Adv. Drug Deliv.
Rev., vol. 60, no. 15, pp. 1638–49, Dec. 2008.
[57] D. Buenger, F. Topuz, and J. Groll, “Hydrogels in sensing applications,” Prog. Polym. Sci., vol.
37, no. 12, pp. 1678–1719, Dec. 2012.
[58] B. Kim, Y. Lee, K. Lee, and W.-G. Koh, “Immobilization of enzymes within hydrogel
microparticles to create optical biosensors for the detection of organophosphorus compounds,”
Curr. Appl. Phys., vol. 9, no. 4, pp. e225–e228, Jul. 2009.
[59] R. Tian, J. Chen, and R. Niu, “The development of low-molecular weight hydrogels for
applications in cancer therapy.,” Nanoscale, vol. 6, no. 7, pp. 3474–82, May 2014.
[60] A. Laurent, “Microspheres and nonspherical particles for embolization.,” Tech. Vasc. Interv.
Radiol., vol. 10, no. 4, pp. 248–56, Dec. 2007.
[61] E. Ozkan and S. Gupta, “Embolization of spinal tumors: vascular anatomy, indications, and
technique.,” Tech. Vasc. Interv. Radiol., vol. 14, no. 3, pp. 129–40, Sep. 2011.
[62] N. Kashyap, N. Kumar, and M. N. V. R. Kumar, “Hydrogels for pharmaceutical and biomedical
applications.,” Crit. Rev. Ther. Drug Carrier Syst., vol. 22, no. 2, pp. 107–49, 2005.
[63] D. Kim and A. E. Herr, “Protein immobilization techniques for microfluidic assays.,”
Biomicrofluidics, vol. 7, no. 4, p. 41501, Jan. 2013.
[64] O. A. Zasedateleva, A. L. Mikheikin, A. Y. Turygin, D. V Prokopenko, A. V Chudinov, E. E.
Belobritskaya, V. R. Chechetkin, and A. S. Zasedatelev, “Gel-based oligonucleotide microarray
72
approach to analyze protein-ssDNA binding specificity.,” Nucleic Acids Res., vol. 36, no. 10, p.
e61, Jun. 2008.
[65] X. Chen, T. K. Neubauer, R. Linhardt, J. S. Dordick, and D. G. Rethwisch, “Enzymatic and
chemoenzymatic approaches to synthesis of sugar-based polymer and hydrogels,” Carbohydr.
Polym., vol. 28, pp. 15–21, 1995.
[66] T. Jamnongkan and S. Kaewpirom, “Potassium Release Kinetics and Water Retention of
Controlled-Release Fertilizers Based on Chitosan Hydrogels,” J. Polym. Environ., vol. 18, no.
3, pp. 413–421, Sep. 2010.
[67] P. Sikareepaisan, U. Ruktanonchai, and P. Supaphol, “Preparation and characterization of
asiaticoside-loaded alginate films and their potential for use as effectual wound dressings,”
Carbohydr. Polym., vol. 83, no. 4, pp. 1457–1469, Feb. 2011.
[68] M. E. Parente, A. Ochoa Andrade, G. Ares, F. Russo, and Á. Jiménez-Kairuz, “Bioadhesive
hydrogels for cosmetic applications,” Int. J. Cosmet. Sci., vol. 37, no. 5, pp. 511–518, Oct.
2015.
[69] G. Jing, L. Wang, H. Yu, W. A. Amer, and L. Zhang, “Recent progress on study of hybrid
hydrogels for water treatment,” Colloids Surfaces A Physicochem. Eng. Asp., vol. 416, pp. 86–
94, Jan. 2013.
[70] M. Nunes, P. Fernandes, and M. Ribeiro, “High-affinity water-soluble system for efficient
naringinase immobilization in polyvinyl alcohol-dimethyl sulfoxide lens-shaped particles.,” J.
Mol. Recognit., vol. 25, no. 11, pp. 580–94, Dec. 2012.
[71] C. M. Hassan and N. A. Peppas, “Structure and Applications of Poly ( vinyl alcohol ) Hydrogels
Produced by Conventional Crosslinking or by Freezing / Thawing Methods,” Adv. Polym. Sci.,
vol. 153, 2000.
[72] A. B. Lugao and S. M. Malmonge, “Use of radiation in the production of hydrogels,” Nucl.
Instruments Methods Phys. Res. Sect. B Beam Interact. with Mater. Atoms, vol. 185, no. 1–4,
pp. 37–42, Dec. 2001.
[73] A. Jayakrishnan and S. R. Jameela, “Glutaraldehyde as a fixative in bioprostheses and drug
delivery matrices,” Biomaterials, vol. 17, no. 5, pp. 471–484, Jan. 1996.
[74] T. Wang, M. Turhan, and S. Gunasekaran, “Selected properties of pH-sensitive, biodegradable
chitosan–poly(vinyl alcohol) hydrogel,” Polym. Int., vol. 53, no. 7, pp. 911–918, Jul. 2004.
[75] V. I. Lozinsky, I. Y. Galaev, F. M. Plieva, I. N. Savina, H. Jungvid, and B. Mattiasson,
“Polymeric cryogels as promising materials of biotechnological interest.,” Trends Biotechnol.,
vol. 21, no. 10, pp. 445–51, Oct. 2003.
[76] F. L. Marten, “Vinyl alcohol polymers,” in Encyclopedia of Chemical Technology, no. 7, 1927,
pp. 1–34.
[77] T. M. C. Maria, R. a. de Carvalho, P. J. a. Sobral, A. M. B. Q. Habitante, and J. Solorza-Feria,
“The effect of the degree of hydrolysis of the PVA and the plasticizer concentration on the
color, opacity, and thermal and mechanical properties of films based on PVA and gelatin
blends,” J. Food Eng., vol. 87, no. 2, pp. 191–199, Jul. 2008.
[78] R. Phosphorimetry, T. K. Itade, Y. W. Ada, S. T. Akegami, A. K. Uroda, and S. K. Ishi, “Effects
of Polymerization and Hydrolysis Degrees of Poly ( vinyl alcohol ) ( PVA ) on the Efficiency of
the PVA-Solid Substrate for,” Anal. Sci., vol. 18, no. March, pp. 337–341, 2002.
[79] S. K. Jha and S. F. D’Souza, “Preparation of polyvinyl alcohol-polyacrylamide composite
polymer membrane by γ-irradiation for entrapment of urease,” J. Biochem. Biophys. Methods,
vol. 62, no. 3, pp. 215–218, Mar. 2005.
[80] S. L. Bourke, M. Al-Khalili, T. Briggs, B. B. Michniak, J. Kohn, and L. A. Poole-Warren, “A
photo-crosslinked poly(vinyl alcohol) hydrogel growth factor release vehicle for wound healing
applications.,” AAPS PharmSci, vol. 5, no. 4, p. E33, 2003.
[81] V. M. Nikolic, A. Krkljes, Z. K. Popovic, Z. V. Lausevic, and S. S. Miljanic, “On the use of
gamma irradiation crosslinked PVA membranes in hydrogen fuel cells,” Electrochem.
commun., vol. 9, no. 11, pp. 2661–2665, Nov. 2007.
[82] L. Zhang, W. Wu, and J. Wang, “Immobilization of activated sludge using improved polyvinyl
alcohol (PVA) gel,” J. Environ. Sci., vol. 19, no. 11, pp. 1293–1297, Jan. 2007.
73
[83] H. Kurokawa, M. Shibayama, T. Ishimaru, S. Nomura, and W. Wu, “Phase behaviour and sol-
gel transition of poly (vinyl alcohol )-borate complex in aqueous solution,” vol. 33, no. 10, pp.
2182–2188, 1992.
[84] N. A. Peppas, J. Z. Hilt, A. Khademhosseini, and R. Langer, “Hydrogels in Biology and
Medicine: From Molecular Principles to Bionanotechnology,” Adv. Mater., vol. 18, no. 11, pp.
1345–1360, Jun. 2006.
[85] K. C. S. Figueiredo, T. L. M. Alves, and C. P. Borges, “Poly (vinyl alcohol) Films Crosslinked by
Glutaraldehyde Under Mild Conditions,” J. Appl. Polym. Sci., vol. 111, pp. 3074–3080, 2008.
[86] H. Chen, Q. Zhang, Y. Dang, and G. Shu, “The Effect of Glutaraldehyde Cross-Linking on the
Enzyme Activity of Immobilized β -Galactosidase on Chitosan Bead,” Adv. J. Food Sci.
Technol., vol. 5, no. 7, pp. 932–935, 2013.
[87] V. Ca, J. Ruiz, and A. Manteco, “Synthesis and properties of hydrogels from poly (vinyl alcohol)
and ethylenediaminetetraacetic dianhydride,” Polymer (Guildf)., vol. 42, pp. 6347–6354, 2001.
[88] H. S. Mansur, C. M. Sadahira, A. N. Souza, and A. A. P. Mansur, “FTIR spectroscopy
characterization of poly (vinyl alcohol) hydrogel with different hydrolysis degree and chemically
crosslinked with glutaraldehyde,” Mater. Sci. Eng. C, vol. 28, no. 4, pp. 539–548, May 2008.
[89] M.-H. Alves, B. E. B. Jensen, A. a a Smith, and A. N. Zelikin, “Poly(vinyl alcohol) physical
hydrogels: new vista on a long serving biomaterial.,” Macromol. Biosci., vol. 11, no. 10, pp.
1293–313, Oct. 2011.
[90] F. M. Plieva, M. Karlsson, M.-R. Aguilar, D. Gomez, S. Mikhalovsky, I. Y. Galaev, and B.
Mattiasson, “Pore structure of macroporous monolithic cryogels prepared from poly(vinyl
alcohol),” J. Appl. Polym. Sci., vol. 100, no. 2, pp. 1057–1066, Apr. 2006.
[91] V. I. Lozinsky and F. M. Plieva, “Poly(vinyl alcohol) cryogels employed as matrices for cell
immobilization. 3. Overview of recent research and developments,” Enzyme Microb. Technol.,
vol. 23, no. 3–4, pp. 227–242, Aug. 1998.
[92] E. Otsuka and A. Suzuki, “A simple method to obtain a swollen PVA gel crosslinked by
hydrogen bonds,” J. Appl. Polym. Sci., vol. 114, no. 1, pp. 10–16, Oct. 2009.
[93] M. Schlieker and K. Vorlop, “A Novel Immobilization Method for Entrapment LentiKats®,” in
Methods in Biotechnology: Immobilization of Enzymes and Cells, 2nd ed., Totowa, NJ:
Humana Press Inc., 2006, pp. 333–343.
[94] A. Boušková, J. Mrákota, R. Stloukal, J. Trögl, V. Pilařová, L. Křiklavová, and T. Lederer,
“Three examples of nitrogen removal from industrial wastewater using Lentikats
Biotechnology,” Desalination, vol. 280, no. 1–3, pp. 191–196, Oct. 2011.
[95] M. Rebroš, M. Rosenberg, Z. Mlichova, and L. Kristofikova, “Hydrolysis of sucrose by invertase
entrapped in polyvinyl alcohol hydrogel capsules,” Food Chem., vol. 102, no. 3, pp. 784–787,
2007.
[96] M. Rebroš, M. Rosenberg, R. Stloukal, and L. Kristofíková, “High efficiency ethanol
fermentation by entrapment of Zymomonas mobilis into LentiKats.,” Lett. Appl. Microbiol., vol.
41, no. 5, pp. 412–6, Jan. 2005.
[97] H. Gröger, E. Capan, A. Barthuber, and K. D. Vorlop, “Asymmetric synthesis of an (R)-
cyanohydrin using enzymes entrapped in lens-shaped gels.,” Org. Lett., vol. 3, no. 13, pp.
1969–72, Jun. 2001.
[98] J. Anes and P. Fernandes, “Towards the continuous production of fructose syrups from inulin
using inulinase entrapped in PVA-based particles,” Biocatal. Agric. Biotechnol., vol. 3, no. 4,
pp. 296–302, Oct. 2014.
[99] S. Cattorini, M. P. C. Marques, F. Carvalho, V. Chheub, J. M. S. Cabral, and P. Fernandes,
“Lentikat ® -based Biocatalysts : Effective Tools for Inulin Hydrolysis,” Chem. Biochem. Eng.,
vol. 23, no. 4, pp. 429–434, 2009.
[100] G.-H. Gong, Y. Hou, Q. Zhao, M.-A. Yu, F. Liao, L. Jiang, and X.-L. Yang, “A new approach for
the immobilization of permeabilized brewer’s yeast cells in a modified composite polyvinyl
alcohol lens-shaped capsule containing montmorillonite and dimethyldioctadecylammonium
bromide for use as a biocatalyst,” Process Biochem., vol. 45, no. 9, pp. 1445–1449, Sep. 2010.
74
[101] F. M. Veronese, C. Mammucari, F. Schiavon, O. Schiavon, S. Lora, F. Secundo, A. Chilin, and
A. Guiotto, “Pegylated enzyme entrapped in poly(vinyl alcohol) hydrogel for biocatalytic
application,” Farm., vol. 56, no. 8, pp. 541–547, Aug. 2001.
[102] A. Idris, N. A. M. Zain, and M. S. Suhaimi, “Immobilization of Baker’s yeast invertase in PVA–
alginate matrix using innovative immobilization technique,” Process Biochem., vol. 43, no. 4,
pp. 331–338, Apr. 2008.
[103] R. Dave and D. Madamwar, “Entrapment of lipase in polymer of polyvinyl alcohol-boric acid for
esterification in organic media,” vol. 5, no. July, pp. 368–372, 2006.
[104] M. Nunes, M. Rosa, P. Fernandes, and M. Ribeiro, “Operational stability of naringinase PVA
lens-shaped microparticles in batch stirred reactors and mini packed bed reactors-one step
closer to industry.,” Bioresour. Technol., vol. 164, pp. 362–70, Jul. 2014.
[105] R. Jovanovic-Malinovska, P. Fernandes, E. Winkelhausen, and L. Fonseca, “Galacto-
oligosaccharides synthesis from lactose and whey by β-galactosidase immobilized in PVA.,”
Appl. Biochem. Biotechnol., vol. 168, no. 5, pp. 1197–211, Dec. 2012.
[106] S. M. L. Silva, C. R. C. Braga, M. V. L. Fook, C. M. O. Raposo, L. H. Carvalho, and E. L.
Canedo, “Application of Infrared Spectroscopy to Analysis of Chitosan / Clay Nanocomposites,”
in Infrared Spectroscopy - Materials Science, Engineering and Technology, 2011, pp. 43–62.
[107] M. Rinaudo, “Chitin and chitosan: Properties and applications,” Prog. Polym. Sci., vol. 31, no.
7, pp. 603–632, Jul. 2006.
[108] M. G. Peter, “Applications and Environmental Aspects of Chitin and Chitosan,” J. Macromol.
Sci. Part A, vol. 32, no. 4, pp. 629–640, Apr. 1995.
[109] G. Lamarque, J. Lucas, C. Viton, and A. Domard, “Physicochemical Behavior of Homogeneous
Series of Acetylated Chitosans in Aqueous Solution : Role of Various Structural Parameters,”
vol. 46, pp. 131–142, 2005.
[110] N. M. Alves and J. F. Mano, “Chitosan derivatives obtained by chemical modifications for
biomedical and environmental applications,” Int. J. Biol. Macromol., vol. 43, no. 5, pp. 401–414,
Dec. 2008.
[111] E. Campos, P. Coimbra, and M. H. Gil, “An improved method for preparing glutaraldehyde
cross-linked chitosan–poly(vinyl alcohol) microparticles,” Polym. Bull., vol. 70, no. 2, pp. 549–
561, Sep. 2012.
[112] M. N. . Ravi Kumar, “A review of chitin and chitosan applications,” React. Funct. Polym., vol.
46, no. 1, pp. 1–27, Nov. 2000.
[113] V. Sinha, A. Singla, S. Wadhawan, R. Kaushik, R. Kumria, K. Bansal, and S. Dhawan,
“Chitosan microspheres as a potential carrier for drugs,” Int. J. Pharm., vol. 274, no. 1–2, pp.
1–33, Apr. 2004.
[114] Q. Yang, F. Dou, B. Liang, and Q. Shen, “Studies of cross-linking reaction on chitosan fiber
with glyoxal,” Carbohydr. Polym., vol. 59, no. 2, pp. 205–210, Jan. 2005.
[115] W. Wan Ngah, C. Endud, and R. Mayanar, “Removal of copper(II) ions from aqueous solution
onto chitosan and cross-linked chitosan beads,” React. Funct. Polym., vol. 50, no. 2, pp. 181–
190, Jan. 2002.
[116] J. Varshosaz, H. Sadrai, and R. Alinagari, “Nasal delivery of insulin using chitosan
microspheres.,” J. Microencapsul., vol. 21, no. 7, pp. 761–74, 2004.
[117] J. Berger, M. Reist, J. M. Mayer, O. Felt, N. A. Peppas, and R. Gurny, “Structure and
interactions in covalently and ionically crosslinked chitosan hydrogels for biomedical
applications.,” Eur. J. Pharm. Biopharm., vol. 57, no. 1, pp. 19–34, 2004.
[118] A. Mendes, P. Castro, and R. Giordano, “Application of chitosan as support for immobilization
of enzymes of industrial interest,” Quim. Nov., vol. 34, no. 5, pp. 831–840, 2011.
[119] D. R. Bhumkar and V. B. Pokharkar, “Studies on effect of pH on cross-linking of chitosan with
sodium tripolyphosphate: a technical note.,” AAPS PharmSciTech, vol. 7, no. 2, p. E50, Jan.
2006.
[120] I. A. Alsarra, S. H. Neau, and M. A. Howard, “Effects of preparative parameters on the
properties of chitosan hydrogel beads containing Candida rugosa lipase.,” Biomaterials, vol.
25, no. 13, pp. 2645–55, 2004.
75
[121] K. Baysal, A. Z. Aroguz, Z. Adiguzel, and B. M. Baysal, “Chitosan/alginate crosslinked
hydrogels: Preparation, characterization and application for cell growth purposes,” Int. J. Biol.
Macromol., vol. 59, pp. 342–348, Aug. 2013.
[122] D. S. Wentworth, D. Skonberg, D. W. Donahue, and A. Ghanem, “Application of Chitosan -
Entrapped B-Galactosidase in a Packed-Bed Reactor System,” J. Appl. Polym. Sci., vol. 91,
pp. 1294–1299, 2004.
[123] G. D. Altun and S. A. Cetinus, “Immobilization of pepsin on chitosan beads,” Food Chem., vol.
100, no. 3, pp. 964–971, Jan. 2007.
[124] M. I. González Siso, E. Lang, B. Carrenõ-Gómez, M. Becerra, F. Otero Espinar, and J. Blanco
Méndez, “Enzyme encapsulation on chitosan microbeads,” Process Biochem., vol. 32, no. 3,
pp. 211–216, Mar. 1997.
[125] Ş. Akkuş Çetinus and H. Nursevin Öztop, “Immobilization of catalase into chemically
crosslinked chitosan beads,” Enzyme Microb. Technol., vol. 32, no. 7, pp. 889–894, Jun. 2003.
[126] D. Boadi and R. Neufeld, “Encapsulation of tannase for the hydrolysis of tea tannins,” Enzyme
Microb. Technol., vol. 28, no. 7–8, pp. 590–595, May 2001.
[127] G. Spagna, R. N. Barbagallo, E. Greco, I. Manenti, and P. G. Pifferi, “A mixture of purified
glycosidases from Aspergillus niger for oenological application immobilised by inclusion in
chitosan gels,” Enzyme Microb. Technol., vol. 30, no. 1, pp. 80–89, Jan. 2002.
[128] K. Carbone, M. Casarci, and M. Varrone, “Crosslinked Poly (Vinyl Alcohol) Supports for the
Immobilization of a Lipolytic Enzyme,” J. Appl. Polym. Sci., vol. 74, no. June, pp. 1881–1889,
1998.
[129] V. G. Kadajji and G. V. Betageri, “Water Soluble Polymers for Pharmaceutical Applications,”
Polymers (Basel)., vol. 3, no. 4, pp. 1972–2009, Nov. 2011.
[130] K. Sonaje, E.-Y. Chuang, K.-J. Lin, T.-C. Yen, F.-Y. Su, M. T. Tseng, and H.-W. Sung,
“Opening of Epithelial Tight Junctions and Enhancement of Paracellular Permeation by
Chitosan: Microscopic, Ultrastructural, and Computed-Tomographic Observations,” Mol.
Pharm., p. 120413125623001, Apr. 2012.
[131] E. Parparita, C. Cheaburu, and C. Vasile, “Morphological, thermal and rheological
characterization of polyvinyl alcohol/chitosan blends,” Cellul. Chem. Technol., vol. 46, no. 9–
10, pp. 571–581, 2012.
[132] S. Jiang, S. Liu, and W. Feng, “PVA hydrogel properties for biomedical application.,” J. Mech.
Behav. Biomed. Mater., vol. 4, no. 7, pp. 1228–33, Oct. 2011.
[133] D. H. Reneker, A. L. Yarin, H. Fong, and S. Koombhongse, “Bending instability of electrically
charged liquid jets of polymer solutions in electrospinning,” J. Appl. Phys., vol. 87, no. 9, p.
4531, 2000.
[134] A. M. De Campos, A. Sánchez, and M. J. Alonso, “Chitosan nanoparticles: a new vehicle for
the improvement of the delivery of drugs to the ocular surface. Application to cyclosporin A,”
Int. J. Pharm., vol. 224, no. 1–2, pp. 159–168, Aug. 2001.
[135] R. H. Schmedlen, K. S. Masters, and J. L. West, “Photocrosslinkable polyvinyl alcohol
hydrogels that can be modified with cell adhesion peptides for use in tissue engineering,”
Biomaterials, vol. 23, no. 22, pp. 4325–4332, Nov. 2002.
[136] D. Archana, L. Upadhyay, R. P. Tewari, J. Dutta, Y. B. Huang, and P. K. Dutta, “Chitosan-
pectin-alginate as a novel scaffold for tissue engineering applications,” vol. 12, no. October, pp.
475–482, 2013.
[137] S. Hsu, S. W. Whu, C.-L. Tsai, Y.-H. Wu, H.-W. Chen, and K.-H. Hsieh, “Chitosan as Scaffold
Materials: Effects of Molecular Weight and Degree of Deacetylation,” J. Polym. Res., vol. 11,
no. 2, pp. 141–147, Jun. 2004.
[138] a Kamari, I. D. Pulford, and J. S. J. Hargreaves, “Chitosan as a potential amendment to
remediate metal contaminated soil - a characterisation study.,” Colloids Surf. B. Biointerfaces,
vol. 82, no. 1, pp. 71–80, Jan. 2011.
[139] M. Kumar, B. P. Tripathi, and V. K. Shahi, “Crosslinked chitosan/polyvinyl alcohol blend beads
for removal and recovery of Cd(II) from wastewater.,” J. Hazard. Mater., vol. 172, no. 2–3, pp.
1041–8, Dec. 2009.
76
[140] E. Portes, C. Gardrat, A. Castellan, and V. Coma, “Environmentally friendly films based on
chitosan and tetrahydrocurcuminoid derivatives exhibiting antibacterial and antioxidative
properties,” Carbohydr. Polym., vol. 76, no. 4, pp. 578–584, May 2009.
[141] V. Ospedale and P. De Mallorca, “Chitosan beads loaded with essential oils in cosmetic
formulations,” J. Cosmet. Sci., vol. 214, no. June, pp. 205–214, 2006.
[142] M. Kita, Y. Ogura, Y. Honda, S. H. Hyon, W. Cha, and Y. Ikada, “Evaluation of polyvinyl alcohol
hydrogel as a soft contact lens material.,” Graefes Arch. Clin. Exp. Ophthalmol., vol. 228, no. 6,
pp. 533–7, Jan. 1990.
[143] F. Gnanaprakasam Thankam, J. Muthu, V. Sankar, and R. Kozhiparambil Gopal, “Growth and
survival of cells in biosynthetic poly vinyl alcohol–alginate IPN hydrogels for cardiac
applications,” Colloids Surfaces B Biointerfaces, vol. 107, pp. 137–145, Jul. 2013.
[144] M. Kobayashi, J. Toguchida, and M. Oka, “Development of the shields for tendon injury repair
using polyvinyl alcohol-hydrogel (PVA-H).,” J. Biomed. Mater. Res., vol. 58, no. 4, pp. 344–51,
2001.
[145] M. Qi, Y. Gu, N. Sakata, D. Kim, Y. Shirouzu, C. Yamamoto, A. Hiura, S. Sumi, and K. Inoue,
“PVA hydrogel sheet macroencapsulation for the bioartificial pancreas.,” Biomaterials, vol. 25,
no. 27, pp. 5885–92, 2004.
[146] M. Abdallh, O. Hamood, and E. Yousif, “Study the Optical properties of Poly (vinyl alcohol)
Doped Cupper Chloride,” J. Al-Nahrain Univ., vol. 16, no. 1, pp. 17–20, 2013.
[147] T. Bilhim, J. M. Pisco, M. Duarte, and A. G. Oliveira, “Polyvinyl alcohol particle size for uterine
artery embolization: a prospective randomized study of initial use of 350-500 μm particles
versus initial use of 500-700 μm particles.,” J. Vasc. Interv. Radiol., vol. 22, no. 1, pp. 21–7,
Jan. 2011.
[148] H. Y. Yoon, S. Son, S. J. Lee, D. G. You, J. Y. Yhee, J. H. Park, M. Swierczewska, S. Lee, I. C.
Kwon, S. H. Kim, K. Kim, and M. G. Pomper, “Glycol chitosan nanoparticles as specialized
cancer therapeutic vehicles: sequential delivery of doxorubicin and Bcl-2 siRNA.,” Sci. Rep.,
vol. 4, p. 6878, Jan. 2014.
[149] H. T. Ta, C. R. Dass, I. Larson, P. F. M. Choong, and D. E. Dunstan, “A chitosan hydrogel
delivery system for osteosarcoma gene therapy with pigment epithelium-derived factor
combined with chemotherapy,” Biomaterials, vol. 30, no. 27, pp. 4815–4823, Sep. 2009.
[150] B. Krajewska, “Application of chitin- and chitosan-based materials for enzyme immobilizations:
a review,” Enzyme Microb. Technol., vol. 35, no. 2–3, pp. 126–139, Aug. 2004.
[151] A. Basso, P. Spizzo, V. Ferrario, L. Knapic, N. Savko, P. Braiuca, C. Ebert, E. Ricca, V.
Calabrò, and L. Gardossi, “Endo- and exo-inulinases: enzyme-substrate interaction and
rational immobilization.,” Biotechnol. Prog., vol. 26, no. 2, pp. 397–405, 2010.
[152] D. M. Lima, P. Fernandes, D. S. Nascimento, R. De Cássia, L. F. Ribeiro, and S. A. De Assis,
“Fructose Syrup : A Biotechnology Asset,” Food Technol. Biotechnol., vol. 9862, no. 4, pp.
424–434, 2011.
[153] R. B. Letters, “Establishing the optimal conditions for fructose production from chicory inulin,”
vol. 18, no. 3, pp. 8263–8270, 2013.
[154] K. Ohta, N. Suetsugu, and T. Nakamura, “Purification and properties of an extracellular
inulinase from Rhizopus sp. strain TN-96,” J. Biosci. Bioeng., vol. 94, no. 1, pp. 78–80, Jul.
2002.
[155] E. Diaz, R. Catana, B. Ferreira, S. Luque, P. Fernandes, and J. Cabral, “Towards the
development of a membrane reactor for enzymatic inulin hydrolysis,” J. Memb. Sci., vol. 273,
no. 1–2, pp. 152–158, Mar. 2006.
[156] M. F. Silva, D. Rigo, V. Mossi, S. Golunski, G. de Oliveira Kuhn, M. Di Luccio, R. Dallago, D.
de Oliveira, J. V. Oliveira, and H. Treichel, “Enzymatic synthesis of fructooligosaccharides by
inulinases from Aspergillus niger and Kluyveromyces marxianus NRRL Y-7571 in aqueous–
organic medium,” Food Chem., vol. 138, no. 1, pp. 148–153, May 2013.
[157] M. Losada and T. Olleros, “Towards a healthier diet for the colon: the influence of
fructooligosaccharides and lactobacilli on intestinal health,” Nutr. Res., vol. 22, no. 1–2, pp. 71–
84, Jan. 2002.
77
[158] N. Kaur and A. K. Gupta, “Applications of inulin and oligofructose in health and nutrition.,” J.
Biosci., vol. 27, no. 7, pp. 703–14, Dec. 2002.
[159] E. Ricca, V. Calabrò, S. Curcio, and G. Iorio, “The state of the art in the production of fructose
from inulin enzymatic hydrolysis.,” Crit. Rev. Biotechnol., vol. 27, no. 3, pp. 129–45, 2007.
[160] R. A. Forshee, M. L. Storey, D. B. Allison, W. H. Glinsmann, G. L. Hein, D. R. Lineback, S. A.
Miller, T. A. Nicklas, G. A. Weaver, and J. S. White, “A critical examination of the evidence
relating high fructose corn syrup and weight gain.,” Crit. Rev. Food Sci. Nutr., vol. 47, no. 6, pp.
561–82, Jan. 2007.
[161] E. J. Vandamme and D. G. Derycke, “Microbial inulinases: fermentation process, properties,
and applications.,” Adv. Appl. Microbiol., vol. 29, pp. 139–76, Jan. 1983.
[162] R. Catana, M. Eloy, J. R. Rocha, B. S. Ferreira, J. M. S. Cabral, and P. Fernandes, “Stability
evaluation of an immobilized enzyme system for inulin hydrolysis,” Food Chem., vol. 101, no. 1,
pp. 260–266, Jan. 2007.
[163] K. Parker, M. Salas, and V. C. Nwosu, “High fructose corn syrup : Production , uses and public
health concerns,” Biotechnol. Mol. Biol. Rev., vol. 5, no. December, pp. 71–78, 2010.
[164] Z.-M. Chi, T. Zhang, T.-S. Cao, X.-Y. Liu, W. Cui, and C.-H. Zhao, “Biotechnological potential
of inulin for bioprocesses.,” Bioresour. Technol., vol. 102, no. 6, pp. 4295–303, Mar. 2011.
[165] Megazyme, “Fructanase Mixture (purified-liquid),” http://secure.megazyme.com/Fructanase-
Mixture-purified-liquid. .
[166] A. Pandey, C. R. Soccol, P. Selvakumar, V. T. Soccol, N. Krieger, and J. D. Fontana, “Recent
Developments in Microbial Inulinases: Its Production, Properties, and Industrial Applications,”
Appl. Biochem. Biotechnol., vol. 81, no. 1, pp. 35–52, 1999.
[167] P. Carniti, P. L. Beltrame, and D. Guardione, “Hydrolysis of Inulin : A Kinetic Study of the
Reaction Catalyzed by an lnulinase from Aspergillus ficuum,” Biotechnol. Bioeng., vol. 37, pp.
575–579, 1991.
[168] R. Catana, B. S. Ferreira, J. M. S. Cabral, and P. Fernandes, “Immobilization of inulinase for
sucrose hydrolysis,” Food Chem., vol. 91, no. 3, pp. 517–520, Jul. 2005.
[169] P. K. Gill, R. K. Manhas, and P. Singh, “Hydrolysis of inulin by immobilized thermostable
extracellular exoinulinase from Aspergillus fumigatus,” J. Food Eng., vol. 76, no. 3, pp. 369–
375, Oct. 2006.
[170] J. R. Rocha, R. Catana, B. S. Ferreira, J. M. S. Cabral, and P. Fernandes, “Design and
characterisation of an enzyme system for inulin hydrolysis,” Food Chem., vol. 95, no. 1, pp. 77–
82, Mar. 2006.
[171] Q. D. Nguyen, J. M. Rezessy-Szabó, B. Czukor, and Á. Hoschke, “Continuous production of
oligofructose syrup from Jerusalem artichoke juice by immobilized endo-inulinase,” Process
Biochem., vol. 46, no. 1, pp. 298–303, Jan. 2011.
[172] A. Richetti, C. B. Munaretto, L. a Lerin, L. Batistella, J. Vladimir Oliveira, R. M. Dallago, V.
Astolfi, M. Di Luccio, M. a Mazutti, D. de Oliveira, and H. Treichel, “Immobilization of inulinase
from Kluyveromyces marxianus NRRL Y-7571 using modified sodium alginate beads.,”
Bioprocess Biosyst. Eng., vol. 35, no. 3, pp. 383–8, Mar. 2012.
[173] E. Aguiar-Oliveira, P. Fernandes, J. M. S. Cabral, and F. Maugeri, “Characterisation of
biocatalysts immobilised in niobium-a new inorganic solid support,” Can. J. Chem. Eng., vol.
91, no. 3, pp. 432–440, Mar. 2013.
[174] T. Yewale, R. S. Singhal, and A. a. Vaidya, “Immobilization of inulinase from Aspergillus niger
NCIM 945 on chitosan and its application in continuous inulin hydrolysis,” Biocatal. Agric.
Biotechnol., vol. 2, no. 2, pp. 96–101, Apr. 2013.
[175] M. F. Silva, D. Rigo, V. Mossi, R. M. Dallago, P. Henrick, G. D. O. Kuhn, C. D. Rosa, D.
Oliveira, J. V. Oliveira, and H. Treichel, “Evaluation of enzymatic activity of commercial
inulinase from Aspergillus niger immobilized in polyurethane foam,” Food Bioprod. Process.,
vol. 91, no. 1, pp. 54–59, Jan. 2013.
[176] J. Missau, A. J. Scheid, E. L. Foletto, S. L. Jahn, M. a Mazutti, and R. C. Kuhn, “Immobilization
of commercial inulinase on alginate–chitosan beads,” Sustain. Chem. Process., vol. 2, no. 1, p.
13, 2014.
78
[177] J. M. S. Ferreira, B.S., Fernandes, P., Cabral, “Design and Modelling of Immobilised
Biocatalytic Reactors,” in Multiphase Bioreactor Design, London: Taylor & Francis, 2001, pp.
85–114.
[178] B. S. Ferreira, P. Fernandes, and C. J.M.S., “Design and Modelling of Immobilised Biocatalytic
Reactors,” in Multiphase Bioreactor Design, London: Taylor & Francis, 2001, pp. 85–114.
[179] D. M. F. Prazeres and J. M. S. Cabral, “Enzymatic membrane reactors,” in Multiphase
bioreactor design, London: Taylor & Francis, 2001, pp. 135–151.
[180] J. M. Bolivar, J. Wiesbauer, and B. Nidetzky, “Biotransformations in microstructured reactors:
more than flowing with the stream?,” Trends Biotechnol., vol. 29, no. 7, pp. 333–42, Jul. 2011.
[181] M. P. C. Marques and P. Fernandes, “Microfluidic devices: useful tools for bioprocess
intensification.,” Molecules, vol. 16, no. 10, pp. 8368–401, Jan. 2011.
[182] C. H. Kim and S. K. Rhee, “Fructose production from Jerusalem artichoke by inulinase
immobilized on chitin,” Biotechnol. Lett., vol. 11, no. 3, pp. 201–206, Mar. 1989.
[183] W. Y. Kim, S. M. Byun, and T. B. Uhm, “Hydrolysis of inulin from Jerusalem artichoke by
inulinase immobilized on aminoethylcellulose,” Enzyme Microb. Technol., vol. 4, no. 4, pp.
239–244, Jul. 1982.
[184] W. Wenling, W. Wuguang Le Huiying, and W. Shiyuan, “Continuous preparation of fructose
syrups from Jerusalem artichoke tuber using immobilized intracellular inulinase from
Kluyveromyces sp. Y-85,” Process Biochem., vol. 34, no. 6–7, pp. 643–646, Sep. 1999.
[185] G. L. Miller, “Use of Dinitrosalicylic Acid Reagent for Determination of Reducing Sugar,” Anal.
Chem., vol. 31, no. 3, pp. 426–428, Mar. 1959.
[186] M. A. P. Nunes, H. Vila-Real, P. C. B. Fernandes, and M. H. L. Ribeiro, “Immobilization of
naringinase in PVA-alginate matrix using an innovative technique.,” Appl. Biochem.
Biotechnol., vol. 160, no. 7, pp. 2129–47, Apr. 2010.
[187] M. M. Bradford, “A rapid and sensitive method for the quantitation of microgram quantities of
protein utilizing the principle of protein-dye binding,” Anal. Biochem., vol. 72, no. 1–2, pp. 248–
254, May 1976.
[188] C. Aymard and A. Belarbi, “Kinetics of thermal deactivation of enzymes: a simple three
parameters phenomenological model can describe the decay of enzyme activity, irrespectively
of the mechanism.,” Enzyme Microb. Technol., vol. 27, no. 8, pp. 612–618, Nov. 2000.
[189] O. S. Veronese, C. Mammucari, P. Caliceti, “Influence of PEGylation on the release of low and
high molecular-weight proteins from PVA matrices,” J. Bioact. Compat. Polym., vol. 14, pp.
315–330, 1999.
[190] W. K. Chui, “Prolonged retention of cross-linked trypsin in calcium alginate microspheres.,” J.
Microencapsul., vol. 14, pp. 51–61, 1997.
[191] T. Siritientong and P. Aramwit, “Characteristics of carboxymethyl cellulose/sericin hydrogels
and the influence of molecular weight of carboxymethyl cellulose,” Macromol. Res., vol. 23, no.
9, pp. 861–866, Sep. 2015.
[192] M. M. Housseiny, “Production of an endoinulinase from Aspergillus niger AUMC 9375, by solid
state fermentation of agricultural wastes, with purification and characterization of the free and
immobilized enzyme.,” J. Microbiol., vol. 52, no. 5, pp. 389–98, May 2014.
[193] V. Vinoth Kumar, M. P. Premkumar, V. kumar Sathyaselvabala, S. Dineshkirupha, J.
Nandagopal, and S. Sivanesan, “Aspergillus niger exo-inulinase purification by three phase
partitioning,” Eng. Life Sci., vol. 11, no. 6, pp. 607–614, Dec. 2011.
[194] Marcin Skowronek and Jan Fiedurek, “Purification and Properties of Extracellular
Endoinulinase from Aspergillus niger 20 OSM,” Food Technol. Biotechnol., vol. 44, 2006.
[195] E. J. Vandamme and D. G. Derycke, “Microbial inulinases: fermentation process, properties,
and applications.,” Adv. Appl. Microbiol., vol. 29, pp. 139–76, 1983.
[196] Keerti, A. Gupta, V. Kumar, A. Dubey, and A. K. Verma, “Kinetic Characterization and Effect of
Immobilized Thermostable β -Glucosidase in Alginate Gel Beads on Sugarcane Juice,” ISRN
Biochem., vol. 2014, pp. 1–8, 2014.
79
[197] I. Migneault, C. Dartiguenave, M. J. Bertrand, and K. C. Waldron, “Glutaraldehyde: Behavior in
aqueous solution, reaction with proteins, and application to enzyme crosslinking,”
Biotechniques, vol. 37, no. 5, pp. 790–802, 2004.
[198] R. Bergamasco, F. J. Bassetti, F. F. de Moraes, and G. M. Zanin, “Characterization of free and
immobilized invertase regarding activity and energy of activation,” Brazilian J. Chem. Eng., vol.
17, no. 4–7, pp. 873–880, Dec. 2000.
[199] A. M. P. Santos, M. G. Oliveira, and F. Maugeri, “Modelling thermal stability and activity of free
and immobilized enzymes as a novel tool for enzyme reactor design.,” Bioresour. Technol., vol.
98, no. 16, pp. 3142–8, 2007.
[200] J. Yao, Q. Chen, G. Zhong, W. Cao, A. Yu, and Y. Liu, “Immobilization and characterization of
tannase from a metagenomic library and its use for removal of tannins from green tea
infusion.,” J. Microbiol. Biotechnol., vol. 24, no. 1, pp. 80–6, Jan. 2014.
[201] M. Dinarvand, A. Ariff, H. Moeini, M. Masomian, S. S. Mousavi, R. Nahavandi, and S. Mustafa,
“Effect of extrinsic and intrinsic parameters on inulinase production by Aspergillus niger ATCC
20611,” Electron. J. Biotechnol., vol. 15, no. 4, pp. 1–11, Jun. 2012.
[202] J. L. Bicas, J. C. Silva, A. P. Dionísio, and G. M. Pastore, “Biotechnological production of
bioflavors and functional sugars,” Ciência e Tecnol. Aliment., vol. 30, no. 1, Mar. 2010.
[203] K. S. Paripoorani, G. Ashwin, P. Vengatapriya, V. Ranjitha, S. Rupasree, V. Vasanth Kumar,
and V. Vinoth Kumar, “Insolubilization of inulinase on magnetite chitosan microparticles, an
easily recoverable and reusable support,” J. Mol. Catal. B Enzym., vol. 113, pp. 47–55, Mar.
2015.
[204] Z. Grosová, M. Rosenberg, M. Rebros, M. Sipocz, and B. Sedlácková, “Entrapment of beta-
galactosidase in polyvinylalcohol hydrogel.,” Biotechnol. Lett., vol. 30, no. 4, pp. 763–7, 2008.
[205] E. M. Ribeiro and P. Fernandes, “Coated-Wall Mini Reactor for Inulin Hydrolysis,” Curr.
Biotechnol., vol. 2, no. 1, pp. 47–52, 2013.
[206] M. J. C. Claudino, D. Soares, F. Van Keulen, M. P. C. Marques, J. M. S. Cabral, and P.
Fernandes, “Immobilization of mycobacterial cells onto silicone – Assessing the feasibility of
the immobilized biocatalyst in the production of androstenedione from sitosterol,” Bioresour.
Technol., vol. 99, no. 7, pp. 2304–2311, May 2008.
[207] M. Karimi, M. Habibi-Rezaei, M. Safari, A. A. Moosavi-Movahedi, M. Sayyah, R. Sadeghi, and
J. Kokini, “Immobilization of endo-inulinase on poly-d-lysine coated CaCO3 micro-particles,”
Food Res. Int., vol. 66, pp. 485–492, Dec. 2014.
[208] M. D. Lilly, W. E. Hornby, and E. M. Crook, “The kinetics of carboxymethylcellulose--ficin in
packed beds.,” Biochem. J., vol. 100, no. 3, pp. 718–23, 1966.
[209] J. A. Figueira, H. H. Sato, and P. Fernandes, “Establishing the Feasibility of Using β-
Glucosidase Entrapped in Lentikats and in Sol–Gel Supports for Cellobiose Hydrolysis,” J.
Agric. Food Chem., vol. 61, no. 3, pp. 626–634, Jan. 2013.
[210] M. Rebroš, M. Rosenberg, Z. Mlichová, L. Krištofíková, and M. Paluch, “A simple entrapment of
glucoamylase into LentiKats® as an efficient catalyst for maltodextrin hydrolysis,” Enzyme
Microb. Technol., vol. 39, no. 4, pp. 800–804, Aug. 2006.
80
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Annexes
Annex I. Quantification of reducing sugars
Most of the methods for determination of carbohydrase, inulinase or invertase activities are
based on the analysis of reducing sugars (RSs). The detection of RSs have been based on the well-
known 3,5-dinitrosalicylic acid (DNS) colorimetric method, since it is a sound, reliable, time- and cost-
efficient method (Figure A1). DNS method can react with substrates with five or six carbon reducing
sugars (fructose and glucose), however if the hydrolytic reaction yields disaccharides such as sucrose,
it will not be detected.
Figure A1. Schematic representation of the reaction between DNS reagent and reducing sugar. In the beginning
of the assay the DNS reagent exhibits a yellow color, which shifts to orange–red if the reaction occurs.
In the present work, the quantification of RSs relies on the specific quantification of fructose
upon hydrolysis of inulin. In this sense, the calibration curve was prepared using different fructose
concentrations ranging from 0 to 5 g.L-1
(Figure A2).
Figure A2. DNS calibration curve for fructose concentrations ranging from 0 to 5 g.L-1
.
82
In order to validate DNS methodology, HPLC analysis was also performed. The calibration
curve was prepared using fructose concentrations ranging from 0 to 60 g.L-1
(Figure A3).
Figure A3. HPLC calibration curve for fructose concentrations ranging from 0 to 60 g.L-1
.
Samples were taken from random assays were analyzed by both methods and the values
obtained were compared. The values obtained by the HPLC method are considered to be more
reliable given the high accuracy of the methodology. Nonetheless, DNS methodology proved to be
accurate when compared to the HPLC quantification. Taking this into account as well as the intrinsic
advantages already mentioned, the DNS method proved to be a reliable method for fructose
quantification.
Annex II. Protein quantification
For the total protein quantification by the Bradford method a calibration curve was also
prepared. Bovine serum albumin (BSA) was used a standard for protein calibration, and the calibration
curve were prepared using a set standards with concentrations ranging from 2.5 to 20 µg.mL-1
(Figure
A4). Inulinase quantification was determined by preparing a 100-fold dilute solution in acetate buffer
100 mM pH 4.5. An inulinase concentration of 1.02 mg.mL-1
was achieved, which is lower than the
expected. Nonetheless, this concentration was used in further analysis.
83
Figure A4. Calibration curve used for total protein quantification, obtained from BSA standards with
concentrations ranging from 2.5 to 20 µg.mL-1
.
Annex III. Flow rate calibration curve for continuous operation
Figure A5. Flow rate (mL.min-1
) calibration curve for rotations per minute (rpm) ranging from 0.5 to 10 rpm.
Calibration was performed with distilled water.
