Effects of different solid state fermentation substrate on biochemical properties of cutinase from Fusarium sp.

Article abstract of DOI:10.1016/j.molcatb.2011.06.003

The present results demonstrate that the catalytic characteristics of cutinase produced by the same strain differ depending on the culture medium used. This conclusion was possible after the study of biochemical characterization and enantioselective prope

Full text of DOI:10.1016/j.molcatb.2011.06.003

Journal of Molecular Catalysis B: Enzymatic 72 (2011) 181–186
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Effects of different solid state fermentation substrate on biochemical properties
of cutinase from Fusarium sp.
a,∗
b
a
Paula Speranza , Patrícia de Oliveira Carvalho , Gabriela Alves Macedo
a
Department of Food Science, State University of Campinas, Rua Monteiro Lobato 80, Caixa Postal 6121, CEP 13083-970, Campinas, SP, Brazil
University of São Francisco, Avenida São Francisco de Assis 218, CEP 12916-900, Bragan c¸ a Paulista, SP, Brazil
b
a r t i c l e i n f o
a b s t r a c t
Article history:
The present results demonstrate that the catalytic characteristics of cutinase produced by the same strain
differ depending on the culture medium used. This conclusion was possible after the study of biochem-
ical characterization and enantioselective properties of cutinases produced by Fusarium oxysporum in
four different culture mediums. The mediums were composed of wheat bran, soybean rind, rice bran
and Jatropha curcas seed cake, different Brazilian agricultural by-products. The largest difference can be
observed on cutinase produced by J. curcas seed cake. This enzyme has been activated in most metal ions
tested and exhibited excellent stability in organic solvent, especially hexane. The cutinase produced in
rice bran showed greatest activity in the presence of p-nitrophenyl butyrate as a substrate, whereas the
other enzymes showed greatest activity in the presence of p-nitrophenyl caprilate. Regarding enantios-
elective properties the cutinase produced in soybean rind showed the best result compared to enzymes
produced in wheat bran.
Received 17 February 2011
Received in revised form 1 June 2011
Accepted 2 June 2011
Available online 13 June 2011
Keywords:
Cutinase
Solid state fermentation
Biochemical properties
Enantioselectivity
©
2011 Elsevier B.V. All rights reserved.
1
. Introduction
mediums for the production of enzymes can lead to the formation
of isoforms with different catalytic characteristics [3]. The study of
these differences, as well as of the factors that affect the activity of
the enzymes is important in order to understand the action of such
enzymes and the alterations they go through in different conditions
of storage and use. Cutinases have been presented as a versatile
enzyme showing several noteworthy properties for application in
industrial products and processes [4]. Finding and characterizing
new cutinases allow further expansion of the possible applications
of such enzymes. The objective of this study was to biochemically
characterize and compare cutinases produced by Fusarium oxys-
porum in four different culture medium composed of wheat bran,
soybean rind, rice bran and Jatropha curcas seed cake and evalu-
ate the differences of these enzymes in the resolution of racemic
mixtures.
Cutinase (E.C. 3.1.1.74) is an enzyme that catalyzes the hydroly-
sis of esters bonds in cutin. Cutin is a biopolymer insoluble in water
composed of fatty acids C16 and C18 and found in the outside sur-
face of the plants’ aerial parts. Cutinases are also able to hydrolyse
a large variety of synthetic esters and show activity on short and
long chains of triacylglicerols [1].
Studies have been conducted with the aim of biochemically
characterizing the cutinases produced by different fermentation
conditions, microorganisms and medium compositions. Solid state
fermentation (SSF) holds tremendous potential for the production
of enzymes. It can be of special interest in those processes where
the crude fermented products may be used directly as enzyme
source [2]. A major advantage of microbial enzymes is that it shows
different biochemical characteristics depending on the genetic
variability of the specie. In addition, previous work performed in
our laboratory demonstrates that the enzymes produced from the
same microorganism and different culture medium, such as from
by-products or residues rich in carbon sources do not necessarily
presented the same biochemical properties. The use of different
2. Materials and methods
2.1. Materials
p-Nitrophenyl butyrate (p-NPB) was purchased from
Sigma–Aldrich Brazil Co. (São Paulo, SP, Brazil). Triton X-100,
tetrahydrofuran and other reagents were purchased from Merck
∗ Corresponding author at: Rua Monteiro Lobato 80, Caixa Postal 6121, CEP 13083-
62, Campinas, SP, Brazil. Tel.: +55 19 3521 2175; fax: +55 19 3521 2153.
E-mail addresses: paulasperanza@hotmail.com, paula@fea.unicamp.br
P. Speranza), gmacedo@fea.unicamp.br (G.A. Macedo).
(São Paulo, SP, Brazil). The agricultural by-products used as the
8
solid culture mediums were donated by different companies from
Campinas, SP, Brazil.
(
1
381-1177/$ – see front matter © 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.molcatb.2011.06.003

1
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P. Speranza et al. / Journal of Molecular Catalysis B: Enzymatic 72 (2011) 181–186
2.2. Microorganism preservation and preparation of the
percentage residual activities were calculated by comparison to
pre-inoculums
untreated control enzyme. All the above tests were carried out in
duplicate.
A F. oxysporum strain isolated from soil and plants was selected
in a previous study with 400 strains of fungi as the best cutinase
producer. The culture medium used for this previous selection con-
tained apple cutin as the sole carbon source [5]. The strain was
maintained in potato dextrose Agar (PDA; Acumedia Manufactures
2
.5.3. Effect of metal ions
The effects of CaCl , KCl, HgCl , MnCl , CoCl , K HPO , NaNO ,
2
2
2
2
2
4
3
FeSO , MgSO , ZnSO , MnSO , K SO , Na SO and NaHSO3 on the
4
4
4
4
2
4
2
4
cutinolytic activity were investigated. Final concentrations of each
metal ion in the reaction mixture were 1 mM and 10 mM. The per-
centages of activities were determined by comparison with the
control mixture with no metal ion added. All the above tests were
carried out in duplicate.
◦
Inc. Lansing, MI, USA) slants and stored at 4 C. The pre-inoculum
was prepared by adding 2.5 mL of distilled water to remove the
7
spores, obtaining a suspension containing 7 × 10 spores/mL.
2.3. Cutinase production
2
.5.4. Substrate specificity
The influence of different substrates on the cutinolytic activ-
Three by-products were selected due to cutinase activity in
preliminary experiments and good development of F. oxysporum
strain: wheat bran, soybean rind and rice bran [3]. J. curcas seed
cake was also included in this study, because of its availability.
Brazil is a major producer of oil and the use in biotechnological
process of cake produced after extraction is very advantageous
from the environmental and economic point of views [6]. Crude
enzymes extracts were prepared as previously reported [3]. The
crude cutinases produced in each medium (wheat bran, soybean
rind, rice bran and J. curcas seed cake) were used in all experiments
of the biochemical characterization. The crude cutinases produced
in the mediums with wheat bran and soybean rind were used for
the enantioselective characterization.
ity was investigated replacing the p-NPB of the reaction described
above (cutinase assay) by the following substrates: p-nitrophenyl
caprilate (p-NPC), p-nitrophenyl laurate (p-NPL) and p-nitrophenyl
palmitate (p-NPP). All the above tests were carried out in duplicate.
2.5.5. Stability in organic solvent
The stability of the crude cutinases in acetone, methanol,
ethanol, butanol, propanol, hexane and octanol was tested. The
mixture of crude cutinases and 1 mL of each organic solvent
given above was incubated for 1 h at optimum temperature of the
enzymes. After this the organic solvents were dried with N and the
2
activities were determined by comparison with the control mixture
with no organic solvent added. All the above tests were carried out
in duplicate.
2.4. Cutinase assay
The activity against p-nitrophenyl butyrate (p-NPB) was deter-
2
.6. Esterification of (R) and (S)-2-octanol and enantioselective
mined as previously reported [5]. The hydrolysis of p-NPB
was spectrophotometrically monitored for the formation of p-
nitrophenol at 405 nm.
esterification of (R,S)-2-octanol
The reactions of esterification and enantioselective esterifica-
tion were performed with the crude cutinase produced in mediums
composed by wheat bran and soybean rind, these enzymes were
selected after preliminary tests (thin layer chromatography), that
indicated these enzymes were the most enantioselective (data not
shown).
One unit of cutinase activity was defined as the amount of
enzymes required to convert 1 ␮mol of p-NPB to p-nitrophenol per
minute, under the specified conditions [7].
2
2
.5. Biochemical characteristics of the crude cutinases
.5.1. Effects of temperature and thermal stability
2.6.1. Esterification of (R) and (S)-2-octanol
To determine the optimum temperature for cutinolytic activ-
The esterification of (R) and (S)-2-octanol was performed in a
ity, enzymatic reactions at various temperatures over the range of
◦
mixed reaction containing 40 mM of each enantiomerically pure
of 2-octanol, 40 mM of octanoic or hexanoic acid, 4 mL of hexane
and the crude cutinase of wheat bran and soybean rind. The reac-
2
5–75 C were performed by using the procedure described above
(
cutinase assay).
In order to determine the thermal stability, aliquots of each
◦
tion was carried out at 30 C, 168 h and 130 rpm. Experiments with
crude cutinase in Eppendorf tubes were incubated for 1 h at various
◦
no added enzyme were carried out to evaluate the percentage of
spontaneous esterification in the system. The ester formation was
followed using a Chrompack gas cromatograph (Chrompack Co.,
Holland) equipped with a flame ionization detector and a column
temperatures over 30 C to boiling temperature. After incubation,
the tubes were rapidly cooled in an ice bath and then brought to
room temperature. The activities were determined with the cuti-
nase assay described previously. The percentage residual activities
were calculated by comparison to untreated control enzyme. All
the above tests were carried out in duplicate.
(
i.d. 0.32 mm, length 30 m) CP-WAX 52 CB. The initial temperature
◦
of the column was 50 C for 2 min then the temperature increase
at a rate of 10 C/min up to 220 C. For the analysis, H was used as
the carrier gas and the detector and injector were set at 250 C and
2
◦
◦
2
◦
2.5.2. Effects of pH and pH stability
◦
20 C, respectively.
The effect of pH on cutinolytics activities was determined by
using the following buffers (all at 0.1 mM): acetate buffer pH 3.6,
4
8
.0, 5.0, 5.6; phosphate buffer pH 6.0, 6.5, 7.0; Tris–HCl buffer pH
.0, 8.5, 9.0 and borate–NaOH pH 9.5, 10.0. The activities were
2.6.2. Gas chromatography analysis
The reaction conditions of the resolution of (R,S)-2-octanol was
performed under the same conditions described above (esterifi-
cation of (R) and (S)-2-octanol). The enantiomeric excess (eep) of
the ester produced was determined using Chrompack gas chro-
matograph (Chrompack Co., Holland) equipped with a fused silica
determined with the cutinase assay described previously at opti-
mum temperature of the enzymes.
The same buffers were used to determine pH stability of the
crudes cutinases for cutinolytics activities. The mixture of each
crude cutinases and 1 mL of buffer given above was incubated
for 24 h at optimum temperature of the enzymes. The activities
were determined with the cutinase assay described previously. The
TM
capillary quiral column BETA DEX 120 (0.25 mm, 60 m, Supelco).
◦
The initial temperature of the column was 95 C for 30 min, and
then the temperature increased at a rate of 5 C/min up to 220 C.
◦
◦

P. Speranza et al. / Journal of Molecular Catalysis B: Enzymatic 72 (2011) 181–186
183
A100
A100
8
6
4
0
0
0
8
6
4
2
0
0
0
0
0
Wheat bran
Soybean rind
Wheat bran
Soybean rind
Rice bran
Rice bran
Jatropha curcas seed cake
Jatropha curcas seed cake
20
0
3
4
5
6
7
pH
8
9
10 11
1
0
20
30
40
50
60
70
80
Temperature (°C)
B 1
00
B 100
8
6
4
0
0
0
8
6
4
2
0
0
0
0
0
Wheat bran
Soybean rind
Rice bran
Wheat bran
Soybean rind
Jatropha curcas seed cake
Rice bran
20
0
Jatropha curcas seed cake
3
0
40 50 60 70 80 90 100
3
4
5
6
7
8
9
10
11
pH
Temperature (°C)
Fig. 2. Optima (A) and stability (B) pH values for crude cutinase produced by culture
mediums compounded by wheat bran, soybean rind, rice bran and Jatropha curcas
seed cake.
Fig. 1. Optima (A) and stability (B) temperature value for crude cutinase produced
by culture mediums compounded by wheat bran, soybean rind, rice bran and Jat-
ropha curcas seed cake.
3
.1.2. Effects of pH and pH stability
As shown in Fig. 2A, the pH optima of crude cutinases produced
For the analysis, H2 was used as the carrier gas and the detector
and injector were set at 250 C and 220 C, respectively.
◦
◦
on mediums composed of wheat bran (28.0 U/mL), soybean rind
25.0 U/mL) and rice bran (11.0 U/mL) were found to be 8.0, whereas
(
the crude cutinase produced on medium with J. curcas seed cake
was found to be 9.0 (11.0 U/mL). Similar results were reported ear-
lier for pH optima of cutinases [10,11]. The results confirm the
literature values, which indicate that most fungal and bacterial
cutinases presented optimum pH in the range of 9.0 [12].
3
. Results and discussion
The results of biochemical characterization of crude cutinases
produced in different culture mediums are presented below. It
was possible to confirm the catalytic specificities of these enzymes
according to the medium used.
Crude cutinase produced in medium composed of rice bran was
stable in the pH range of 6.0–7.0 after 24 h, retaining more than 80%
of the initial activity. This enzyme was also more stable at acid pH,
with over 60% of residual activity after 24 h in pH ranging from 3.5
to 5.5. The crude cutinase of soybean rind retained more than 70% of
the initial activity from pH 6.0 to 8.5 after 24 h. The crude cutinase
of J. curcas seed cake showed pH-stability ranging from pH 6.0 to
6.6 after 24 h. The enzyme produced in medium composed of wheat
bran, was stable at pH 6.0 and relatively stable at pH 6.6 (Fig. 2B).
The stability of enzymes in acid, neutral and basic pH is important
in terms of industrial application. These results are in accordance
with earlier reports in which the pH-stability of cutinase from F.
solani pisi was between 6.0 and 8.5 [8].
3
3
.1. Biochemical characteristics of the crude enzymes
.1.1. Effects of temperature and thermal stability
Optimum temperature of crude cutinases produced on medi-
ums composed of rice bran (8.5 U/mL) and J. curcas seed cake
◦
(
9.0 U/mL) was 30 C, while enzymes produced on mediums with
wheat bran (24.0 U/mL) and soybean rind (21.0 U/mL), showed
optimal temperature at 37 C. The other peaks found in the enzyme
produced in J. curcas seed cake may indicate the presence of iso-
forms of cutinase or other enzymes present on crude extract that
also hydrolyze p-NPB (Fig. 1A). These results are in accordance
with earlier reports in which the temperature optimums of two
◦
3.1.3. Effect of some metal ions
◦
cutinases from Fusarium solani are between 35 and 45 C [8].
To determine whether the cutinase requires a metal cofactor for
The enzymes produced on mediums composed of wheat bran
activity and if the need of a cofactor changes according to the cul-
ture medium of the enzyme, they were incubated with the metal
ions and then assayed for cutinase activity against p-NPB. The val-
ues of enzymatic activity of the controls were: 32.0 U/mL (wheat
bran), 24.0 U/mL (soybean rind), 15.0 U/mL (rice bran) and 9.5 U/mL
◦
and rice bran were completely stable at 30 C when incubated for
1
remained completely stable at temperatures 30 and 40 C, the same
stability was observed in the enzyme produced in the medium
composed of J. curcas seed cake. This enzyme exhibited superior
h. The enzyme produced on medium composed of soybean rind
◦
(J. curcas seed cake). The results in Fig. 3 indicates that the enzyme
◦
thermostability, with residual activity of over 60% after 1 h at 50 C
produced on mediums composed with soybean rind and rice bran
were inhibited or slightly activated in the presence of the metal
ions tested. The enzyme produced in medium with wheat bran was
(
Fig. 1B). Earlier results reported indicated that cutinase from F.
◦
solani pisi was stable from 40 to 60 C [9].

1
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P. Speranza et al. / Journal of Molecular Catalysis B: Enzymatic 72 (2011) 181–186
Fig. 3. Effect of metal ions (1 and 10 mM) on the activity of cutinases produced by culture mediums compounded by wheat bran (A), soybean rind (B), rice bran (C) and
Jatropha curcas seed cake (D).
mostly activated in the presence of ZnSO4 and CoCl2 in concentra-
tion of 1 mM. The enzyme produced in medium with J. curcas seed
cake, different of the other enzymes, was activated in the pres-
ence of most metal ions, highlighting MnSO4, CaCl2, ZnSO4, FeSO4,
the enzyme produced in wheat bran, 21.0 U/mL for the enzyme pro-
duced in soybean rind, 11.0 U/mL for the enzyme produced in rice
bran and 9 U/mL for the enzyme produced in J. curcas seed cake. The
results in Fig. 5 indicate that the enzymes produced in mediums
composed with wheat bran, soybean rind and rice bran were not
very stable in organic solvents. The enzyme of soybean rind and rice
bran were relatively more stable in the presence of hexane, keeping
71% and 66% respectively of their activities compared with control.
On the other hand, the enzyme produced in medium with J. curcas
seed cake was fairly stable in the presence of organic solvents, its
activity remained high in all solvents evaluated, highlighting the
hexane, butanol and propanol. The results suggest that the enzyme
of J. curcas seed cake is stable in the presence of organic solvents,
which is a very interesting property for industrial applications. The
use of organic solvents in biocatalysis reactions has some advan-
tages such as increased solubility of organic substrates insoluble
in water and the ability to shift the thermodynamic equilibrium of
many processes for the synthesis path [18]. As an example, we can
mention the reactions of esterification and transesterification for
the production of flavorings [19,20], production of biodiesel [21]
and the resolution of racemic mixtures [3].
MnSO , CoCl and KCl (1 mM) and FeSO4 (10 mM). The enzymes
4
2
were quite inhibited in the presence of HgCl2 in both concen-
trations, indicating that –SH groups are essential for the enzyme
activity [13].
The distinct behavior regarding the need for minerals in the
enzyme produced in J. curcas seed cake can be attributed to changes
in the protein structure of this enzyme and consequent change in
the active site. Other possible explanations can be attributed to the
binding of minerals to some inhibitor present in the crude extract or
even due to its own medium, which may be deficient in these min-
erals. Previous research has shown that esterase and cutinases were
similar to the enzymes produced in wheat bran, soybean rind and
rice bran, the enzymes are inhibited or lightly activated by minerals
tested [14–16]. Cutinase activated by the presence of minerals were
not found in previous studies, nor were they found with the enzyme
produced in J. curcas seed cake. This feature can be exploited in
industrial processes in which these minerals are necessary.
3.1.4. Substrate specificity
The chain length specificity of esters was investigated using
p-nitrophenyl-fatty acyl esters (Fig. 4). The enzymes produced in
wheat bran (54.0 U/mL), soybean rind (34.0 U/mL) and J. curcas seed
cake (25.0 U/mL) had higher specificity in medium-chain fatty acids
(
(
p-NPC). The enzyme produced in medium composed of rice bran
10 U/mL) showed higher specificity in short-chain fatty acids (p-
NPB). You can see that the four enzymes had higher specificity for
short and medium chain fatty acids and the activities toward long
chain p-NP-fatty acids were low. Other studies have indicated that
cutinases showed specificity for short and medium carbon chain of
fatty acids [8,14,17].
3.1.5. Stability in organic solvent
In order to evaluate the stability in organic solvents the samples
were compared with controls, with activity values of 24.0 U/mL for
Fig. 4. Substrate specificities of crude cutinase produced in different culture medi-
ums toward p-nitrophenyl esters.

P. Speranza et al. / Journal of Molecular Catalysis B: Enzymatic 72 (2011) 181–186
185
Table 1
Esterification of pure alcohols (R) and (S)-2-octanol with octanoic and hexanoic acids catalyzed by crude cutinases produced in the culture mediums compounded by wheat
bran and soybean rind.
Substrate for production of cutinase
% Esterification
Octanoic acid
Hexanoic acid
(
R)-2-octanol
(S)-2-octanol
(R)-2-octanol
(S)-2-octanol
Wheat bran
Soybean bran
15.4
26.3
45.4
42.5
8.5
4.8
7.8
6.9
Table 2
Conversion (c) and enantiomeric excess of S-ester product (eep) after 168 h of reaction of racemic alcohol (R,S)-2-octanol with octanoic and hexanoic acids catalyzed by crude
cutinase produced in the culture mediums compounded by wheat bran and soybean rind.
Substrate for production of cutinase
Octanoic acid
Hexanoic acid
c (%)
eep (%)
E
c (%)
eep (%)
E
Wheat bran
Soybean bran
38.3
40.4
45.6
59.8
3.5
5.9
6.8
7.8
25.7
23.8
1.7
1.7
c: conversion given as the percentage of initial alcohol esterified after the reaction time; eep enantiomeric excess of the ester produced; E: enantiomeric ratio.
3.2. Esterification of (R) and (S)-2-octanol and enantioselective
of conversion, enantiomeric excess and enantiomeric rate, its use
esterification of (R,S)-2-octanol
as a catalyst in the resolution of racemic alcohols such as (R,S)-
2
-octanol, appears to be quite interesting. The results obtained
3
.2.1. Esterification of (R) and (S)-2-octanol
Table 1 presents the values of esterification of enantiomerically
using hexanoic acid in relation to the values of conversion, enan-
tiomeric excess and enantiomeric rate were lower than the reaction
with octanoic acid. The results obtained using the octanoic acid
showed to be promising. The optimization of enantioselective reac-
tions may facilitate the achievement of the most interesting results,
allowing the use of these enzymes especially that produced in soy-
bean rind, for the enantioselective process.
pure alcohols (R) and (S)-2-octanol with octanoic and hexanoic
acids catalyzed by crude cutinases produced in mediums composed
with wheat bran and soybean rind. The results obtained using the
octanoic acid show good rate of esterification for both enzymes,
indicating enantiopreference for the (S)-enantiomer. The results
obtained using hexanoic acid show low yields for both enzymes
after 168 h of reaction, and it is also difficult to ascertain any pref-
erence for one of the enantiomers. Although octanoic and hexanoic
acids are similar in number of carbons, the results show a greater
presence in the esterification of octanoic acid. A possible expla-
nation may be due to the fact that the active site of each enzyme
recognizes more efficiently the carbon chain of octanoic acid.
4. Conclusions
The results confirm that although the enzyme has been pro-
duced by the same microorganism, its catalytic characteristics
differ depending on the medium used. One possible explanation
for this phenomenon is that the use of different mediums for
the growth of microorganism forces it to adapt to these differ-
ent sources of nutrients, thus producing isoforms. Another possible
explanation is that possible components of the medium stabiles in
a different way the enzyme.
3.2.2. Enantioselective esterification of (R,S)–2-octanol
In Table 2 the results of conversion (c), enantiomeric excess of S-
ester product (eep) and enantiomeric rate (E) after 168 h of reaction
of racemic alcohol (R,S)-2-octanol with octanoic and hexanoic acids
catalyzed by crude cutinase produced in mediums composed with
wheat bran and soybean rind are presented. The results obtained
using the octanoic acid show that the enzyme produced in soybean
rind was more enantioselective. Since this enzyme has good values
Acknowledgement
The authors wish to thank the Coordena c¸ ão de Aperfei c¸ oamento
de Pessoal de Nível Superior (CAPES) for the scholarship.
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Fig. 5. Effect of different organic solvents on the activity of crude cutinase.

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2007) 588–594.
17] M.B. Rubio, R.E. Cardoza, A.R. Hermosa, S. Gutiérrez, E. Monte, Curr. Genet. 54
2008) 301–312.
Her research is in improvement of catalytic performance of enzymes and industrial
applications of enzymes.
(
Patricia de Oliveira Carvalho Pharmaceutical at Federal University of Alfenas
1
(
1989), Master (1994) and Ph.D. (1998) in Food Science from State University of
Campinas. She is currently an associate professor at the University San Francisco
working in undergraduate and graduate, with the following lines of research: 1,
composition and functional properties of food; 2, enzymology; 3, biocatalysis and
biotransformation. It also develops projects in the Pharmaceutical Care.
(
(
[
[
18] A.M. Klibanov, Trends Biotechnol. 15 (1997) 97–101.
19] D.P.C. Barros, F. Lemos, L.P. Fonseca, J.M.S. Cabral, Mol. Catal. B: Enzym. 66
Gabriela Alves Macedo Food engineer at the State University of Campinas (1993),
Master (1995) and Ph.D. (1997) in Food Science from State University of Camp-
inas. She worked at the Research Center Rhodia Food, a researcher in the area of
food ingredient (1999–2002). She is currently Associate Professor in the Depart-
ment of Food Science in Biochemistry and Food Biotechnology at the Faculty of
Food Engineering at State University of Campinas-UNICAMP. Focuses her research
on the production of microbial enzymes and development of microbial and enzy-
matic processes for producing compounds of interest and increase the bioavailability
of bioactive compounds in foods and feeds.
(
2010) 285–293.
[
[
20] K. Horri, T. Adachi, T. Tanino, T. Tanaka, A. Kotaka, H. Sahara, T. Hashimoto, N.
Kuratani, S. Shibasaki, C. Ogino, H. Noda, Y. Hata, M. Ueda, A. Kondo, Enzyme
Microb. Technol. 46 (2010) 194–199.
21] S.M. Badenes, F. Lemos, J.M.S. Cabral, J. Chem. Technol. Biotechnol. 85 (2010)
9
93–998.
Paula Speranza Food Engineer at the State University of Campinas (2001), Mas-
ter (2010) and Ph.D. student in Food Science from State University of Campinas.