ORIGINAL PAPER

Pentynyl dextran as a support matrix for immobilization of serineprotease subtilisin Carlsberg and its use for transesterificationof N-acetyl-L-phenylalanine ethyl ester in organic media

Muhammad Nazir Tahir • Eunae Cho •

Petra Mischnick • Jae Yung Lee • Jae-Hyuk Yu •

Seunho Jung

Received: 14 May 2013 / Accepted: 13 August 2013

� Springer-Verlag Berlin Heidelberg 2013

Abstract In this study, serine protease (subtilisin Carls-

berg) was immobilized on pentynyl dextran (PyD, O–

alkynyl ether of dextran, 1) and used for the transesterifi-

cation of N-acetyl-L-phenylalanine ethyl ester (2) with

different aliphatic (1-propanol, 1-butanol, 1-pentanol,

1-hexanol) and aromatic (benzyl alcohol, 2-phenyl ethanol,

4-phenyl-1-butanol) alcohols in tetrahydrofuran (THF).

The effect of carbon chain length in aliphatic and aromatic

alcohols on initial and average transesterification rate,

transesterification activity of immobilized enzyme and

yield of the reaction under selected reaction conditions was

investigated. The transesterification reactivity of the

enzyme and yield of the reaction increased as the chain

length of the alcohols decreased. Furthermore, almost no

change in yield was observed when the immobilized

enzyme was repeatedly used for selected alcohols over six

cycles. Intrinsic fluorescence analysis showed that the

catalytic activity of the immobilized enzyme in THF was

maintained due to retention of the tertiary structure of the

enzyme after immobilization on PyD (1).

Keywords Enzyme catalysis � Immobilization �N-acetyl-L-phenyl ethyl ester � Pentynyl dextran �Subtilisin Carlsberg � Transesterification

Abbreviation

APAEE N-acetyl-L-phenylalanine ethyl ester (2)

DS Degree of substitution

EA Elemental analysis

PyD Pentynyl dextran (1)

SC Subtilisin Carlsberg

Trp Tryptophan

Introduction

Polysaccharides after proper chemical modification have

been demonstrated to be candidates as immobilization sup-

ports for enzymes [1]. In addition to their low costs in some

cases, they are also non-toxic, biocompatible, inert in phys-

iological conditions and can provide an appropriate micro-

environment for enzymes under different reaction conditions.

Attachment to a hydrophobic support normally increases the

rigidity of the immobilized enzyme, making it more resistant

to small conformational changes induced by heat, organic

solvents and denaturing agents. Using chemical modifica-

tions, the hydrophilic–lipophilic balance in the polysaccha-

rides can be tuned to adapt it to the enzymes structure.

Different chemically modified polysaccharides e.g. agarose

and cross-linked dextran [2, 3] carboxymethyl cellulose,

alginate, chitosan [2, 4] and lignocelluloses [5] have been

used as immobilization supports for a variety of enzymes.

M. N. Tahir � E. Cho � S. Jung (&)

Department of Bioscience and Biotechnology, Center for

Biotechnology Research in UBITA(CBRU),

Konkuk University, Seoul 143-701, Republic of Korea

e-mail: shjung@konkuk.ac.kr

P. Mischnick

Institute of Food Chemistry, Technische Universitat

Braunschweig, Schleinitzstr. 20, 38106 Braunschweig, Germany

J. Y. Lee

Department of Biological Science, Mokpo National University,

Jeonnam 534-729, Republic of Korea

J.-H. Yu

Departments of Bacteriology and Genetics, and Molecular and

Environmental Toxicology Center, University of Wisconsin,

Madison, WI, USA

123

Bioprocess Biosyst Eng

DOI 10.1007/s00449-013-1038-8

Subtilisin Carlsberg (SC), a member of the protease class

of enzymes, has attracted much attention because of its use as

a biocatalyst in organic synthesis reactions [6, 7]; however, its

applications in synthetic chemistry have been limited due to

the very limited reactivity of the free (un-immobilized)

enzyme in organic media [8]. It has been hypothesized that

organic solvents extract enzyme-bound water, which is the

main factor contributing to altered enzyme activity in organic

media when these techniques are utilized [9]. The primary

method used to increase the activity of SC in organic media

includes addition of inorganic salts such as KCl before

lyophilization, co-lyophilization with cyclodextrin and

immobilization on a solid support [10–14]. There are some

examples where SC was immobilized on chitosan and used as

a biocatalyst in esterification or peptide synthesis reactions

[1, 15–17]. To the best of our knowledge, there have been no

examples in the literature where native or modified dextran

was used as an immobilization support to enhance SC activity

in organic media.

We have prepared and characterized a number of poly-

saccharide alkynyl ethers [18–20], which are interesting

compounds due to the physical and chemical properties of the

terminal alkynes. Pentynyl dextran (PyD) was prepared by

modification of high molecular weight (500 kDa) dextran

and was shown to have special properties due to its specific

structure [18]. Beside the hydrophilic backbone of the poly-

saccharide, which can bind residual water molecules, the

carbon rich �ðCH2Þ3 � C � CH residues introduce hydro-

phobic properties, which can be varied by the degree of

substitution (DS = average number of substituted OH/glu-

cosyl unit, max = 3 for dextran). In addition, terminal

acetylenic groups in PyD can act as both a hydrogen bonding

donor and acceptor. Thus, PyD enables several types of weak

cooperative interactions to support adsorption of the enzyme.

The effectiveness of synthetic as well as carbohydrate con-

taining block-copolymers as excipient for enzymes in organic

solvents has been demonstrated by Naka et al. [21]. There-

fore, PyD (1) is a promising candidate for use as an immo-

bilization support for SC.

Many techniques used for enzyme immobilization include

adsorption on polymer-based or inorganic materials, encap-

sulation, covalent attachment to carrier and cross-linking of

different materials for example using glutaraldehyde [22–

24]. Among these techniques, adsorption is most popular due

to its simple process and high activity yield [22], therefore, we

selected this technique for immobilization of SC on PyD (1).

In this study, we used PyD (1) to immobilize the protease

SC and examined the potential of using this system as a

biocatalyst in the transesterification reaction of N-acetyl-L-

phenylalanine ethyl ester (APAEE, 2) with different aliphatic

and aromatic alcohols at room temperature and in an organic

solvent (THF).

Experimental

Materials and methods

N-acetyl-L-phenylalanine ethyl ester (APAEE, 2), protease

(bacterial, SC 7.0–14.0 units/mg), 1-butanol (C99 %),

THF (C99.9 %, anhydrous), 1-propanol (C99.7 %),

1-pentanol (C99 %), and benzyl alcohol (C99.8 %) were

purchased from Sigma-Aldrich. 2-Phenyl ethanol (C99 %),

1-hexanol (C99.9 %) and 4-phenyl-1-butanol (C99 %)

were obtained from Fluka. All chemicals were used with-

out further purification or any other treatment unless stated

otherwise. Pentynyl dextran (PyD, 1) was synthesised from

dextran (Sigma, 500 kDa) and characterized as described

earlier [18]. Elemental analysis (%, w/w): found C 51.40,

H 6.64; theor. for DS(EA) 0.43: C 51.37, H 6.65. Monomer

composition [substituted position (mol %)]: un-68.6,

mono: 21.4 %, di: 7.9 %, tri: 2.3 %. DS values in position

2: 0.18, 3: 0.11, 4: 0.11, 6: 0.02; average DS(GC) 0.43;

about 13 % of PyD. Surface area: 3.3 ± 0.4 m2/g.

Enzyme immobilization

PyD (1) DS 0.43 was used as an immobilization support.

SC was dissolved in distilled water (2 ml) to form a 2 %

solution. The enzyme solution was mixed with 200 mg of

PyD for 60 min. Un-immobilized enzyme was removed by

centrifugation in water (three times) and the biocatalyst

(enzyme immobilized on PyD) was dried under vacuum at

low temperature.

The amount of enzyme protein immobilized on PyD (1)

was determined based on the nitrogen content from the ele-

mental analysis. The protein content was calculated accord-

ing to Mariotti et al. [25]: % of protein = % of N 9 6.25.

EA of PyD: Calculated: C 51.37, H 6.65, Found: C

51.40, H 6.64, N 0.00.

EA of PyD-protease: C 52.71, H 6.82, N 0.07

(0.07 9 6.25 = 0.44 % Protein).

Number of enzyme units in biocatalyst was calculated as:

%Protein ¼ %N� 6:25 ¼ 0:07� 6:25 ¼ 0:44%

Enzyme in 50 mg Biocatalyst ¼ %Protein

100

� Biocatalyst ðmg) = 0:22 mg ðprotease)

Enzyme units in Biocatalyst ¼ units/mg

� wt: of enzyme in biocatalyst ¼ 1:54�3:08 u

¼ 4:4 mg enzyme/g of biocatalyst

Transesterification activity of the biocatalyst

The transesterification activities of the immobilized

enzyme was determined with reference to the initial

Bioprocess Biosyst Eng

123

transesterification rate between APAEE (2) and different

aliphatic and aromatic alcohols (Scheme 1) in THF.

APAEE (2, 235.3 mg, 1 mmol) and alcohol (10 mmol

each) were added to THF (10 mL) immediately followed

by the addition of the biocatalyst (protease SC immobi-

lized on PyD (1), 50 mg). The reaction was conducted at

room temperature with continuous stirring. A parallel

reaction with all reagents and identical conditions but

with the free (un-immobilized) enzyme was also con-

ducted. The progress of the reaction was monitored by

removing aliquots at different time intervals that were

then analyzed by (a) thin layer chromatography (EtOAc/

n-hexane, 3:1) and (b) high pressure liquid chromatog-

raphy (HPLC, water/acetonitrile 60:40). The transesteri-

fication activity was expressed as micromoles of alcohol

consumed per minute per gram of biocatalyst and was

calculated as follows:

Transesterification activity lmol min�1 g�1� �

¼Aver: consumption of subsrate ðlmolÞ

Time consumed ðminÞAmt: of biocatalyst ðgÞ

One protease unit was defined as the consumption of

1 lmol of alcohol per minute in the transesterification

reaction under conditions applied while the specific activity

was defined as the number of enzyme units per mg of solid

material.

High pressure liquid chromatography (HPLC)

A Jupiter C18 column (5 lM, 250 9 4.60 mm) was used

for HPLC (Shimadzu, Japan) analysis. The analysis was

carried out at 30 �C and a flow rate of 2 mL/min in the

mobile phase (water:acetonitrile = 60:40, v/v). Elution

was monitored at 220–270 nm.

Fluorescence measurement

The intrinsic fluorescence of SC (0.26 mg/mL) was

recorded using a spectrofluorophotometer (Shimadzu, RF-

5310PC), after being equilibrated (25 h) in the solvent at

room temperature. Trp fluorescence emission spectra were

collected in the range of 285–445 nm after excitation of the

samples at 280 nm. The excitation and emission slits had a

width of 1.5–3 nm, respectively.

Elemental analysis

Elemetal analysis was performed at Korea Basic Science

Institute (KBSI) Busan, Korea, using Vario-Micro Cube

elemental analyzer (Elementar Analysensystem GmbH,

Germany).

Results and discussion

Pentynyl dextran (PyD, 1) as immobilization support

PyD was obtained by the etherification of dextran, an a-

1,6-linked and partially branched glucan, with lithium

dimsyl and pentynyl chloride in DMSO [18]. The sub-

stitution pattern of PyD (1) in the glucosyl units, i.e. the

molar ratio of non-, mono- (2-,3- and 4-O), di- (2,3-, 2,4-

and 3,4-di-O), and 2,3,4-tri-O-pentynyl glucosyl units

present in the polymer derivative, showed a remarkable

deviation from a random distribution (analog to Spurlin

model for cellulose derivatives [26]) of substituents with

areas of high density of alkynyl residues and a high

portion of unsubstituted areas in the polymer chain. This

ensures very good protection, stablization and activation

of the enzymes, which was also observed by Naka et al.

[21], who found that block co-polymers with hydrophobic

and lipophilic sequences were very well suited for enzyme

stabilization. The surface area of PyD (1) was only

3.3 ± 0.4 m2/g and was relatively low compared to most

other commercial adsorbents but it can adsorb and protect

enzymes very well due to its specific structure, which was

visible at higher magnification (Fig. 1). PyD (1) is a soft

fluffy material with fibrous, sheet-like and spherical-

shaped nano-structures. Such small structures are consid-

ered more suitable for enzyme protection and stabilization

after immobilization as reported by Fuentes et al. [27].

Tran et al. [28] also reported that immobilization of

enzymes in or on flat sheets or fibrous structures (same as

shown for PyD in Fig. 1) show better results regarding the

stability and activity of immobilized enzymes. Along with

the other advantages of using such materials (like PyD)

for enzyme immobilization, it also cannot be broken down

into small pieces during magnetic stirring, which can

result in a loss of immobilized enzyme activity [29, 30].

The spherical and fibrous structures of PyD (Fig. 1a) were

not visible after immobilization of enzyme (Fig. 1b). This

was likely the case because the small (spherical) struc-

tures were destroyed and converted into sheet-like struc-

tures or just combined together to form aggregates, which

helps not only in the immobilization of the enzyme but

also in protecting the enzyme from denaturing in organic

media.

Evaluation of free and immobilized protease

for transesterification of APAEE (2)

Protease enzymes (bacterial SC, 7-14 u/mg) were dissolved

in water and immobilized on PyD (1) by the adsorption

method. The immobilized enzyme (biocatalyst) was used

for transesterification of APAEE (2) with different ali-

phatic (1-propanol, 1-butanol, 1-pentanol, 1-hexanol) and

Bioprocess Biosyst Eng

123

aromatic (benzyl alcohol, 2-phenyl ethanol, 4-phenyl-1-

butanol) alcohols in THF. The same amount of biocatalyst

(50 mg per 10 mL reaction mixture) was used in all reac-

tions. Free protease (without immobilization) was also used

in parallel under the same reaction conditions as described

in the experimental section but it did not catalyze the

reaction under the conditions applied.

The enzyme used in this study contained 7–14 units/mg

of solid material (data from supplier). The binding capacity

of PyD (1) for SC was 0.44 % as determined from the

Scheme 1 Transesterification

reaction of N-acetyl-L-

phenylalanine ethyl ester

(APAEE, 2) with different

aliphatic and aromatic alcohols

catalyzed by subtilisin

Carlsberg (SC) immobilized on

pentynyl dextran (PyD, 1)

Fig. 1 SEM image of pentynyl

dextran (PyD, 1) a before

immobilization of subtilsin

Carlsberg (Hitachi Field-

Emission Scanning Electron

Microscop S-4800, KTH

Stockholm), b after

immobilization of subtilsin

Carlsberg (JEOL JSM-6380,

Konkuk University Seoul)

Bioprocess Biosyst Eng

123

nitrogen content of the biocatalyst. Thus, the enzyme units

in 50 mg of biocatalyst can vary from 1.5 to 3.1 units/mg

of the biocatalyst, which is based on the equations

described in the experimental section. Thus on average,

there were about 2 units of enzyme per 50 mg (amount

used in all reactions) of biocatalyst (calculated on the base

of 10 units/mg of solid material as an average). Although,

there was only a small amount of enzyme in the biocatalyst

used in each reaction, it was very active in catalyzing the

transesterification reaction as indicated from their yields

(Table 1).

The specific activity of the enzyme (number of enzyme

units per mg of the solid material/biocatalyst), as defined

by Chang et al. [31], was enhanced after immobilization. In

all reactions, it was higher than 2. In some cases, e.g.

propanol (3), butanol (4) and pentanol (5), it was above 5.

This hyperactivation of enzymes after immobilization is

already reported in the literature for other enzymes [32]

and is most probably related to the structure and confor-

mation of the enzyme on the immobilization support.

Normally, a specific hydrophobic–hydrophilic balance is

required in the immobilization support to protect the

enzyme in organic media, while maintaining or enhancing

its activity. A change in the chemical structure of the

immobilization support will not only influence its binding

capacity but also the conformation of the enzyme. A

change in the hydrophilicity–hydrophobicity of the

immobilization support leads to a different spatial

arrangement and as a consequence, its ability to accom-

modate the enzyme will also be changed. Chen et al. [33]

showed that a small change in the hydrophobic–hydro-

philic balance of the immobilization support not only

changes the binding ability but also specific activity of the

bound enzyme.

To compare the yield of the reaction (Scheme 1) and

transesterification activity of the immobilized enzyme for

different alcohols, the reaction time was fixed for 24 h. To

calculate the initial rate of the transesterification reaction,

samples were withdrawn after 0, 15, 30, 45 and 60 min and

the initial rate (Table 1) was calculated from the slope of

the straight line (R2 = 0.992) between the yield and

reaction time (Slope = Y2-Y1/X2-X1). The initial rate in

first 15 min (0–15 min) was little bit lower than in the

second interval (15–30 min). This likely occurred because

the biocatalyst required some time to adapt to the reaction

conditions. Therefore, the interval from 15 to 30 min was

used to calculate the initial rate of the reaction. The aver-

age rate of the reaction was calculated for the first 60 min

of the reaction. The yield of the transesterification reaction

was calculated from the HPLC chromatogram as shown in

Fig. 2 for 1-hexanol. For quantitative analysis, an APAEE

(2) solution of different concentrations was subjected to the

HPLC and standard curve was established using the con-

centration and peak area. The yield, initial rate, average

rate and transesterification activity (Fig. 3, Table 1)

decreased as the length of the aliphatic alcohol chain was

increased. The same trend was observed for aromatic

alcohols. The rate of the reaction or transesterification

activity decreased with an increase in aliphatic chain length

for both aliphatic and aromatic alcohols, but the aliphatic

alcohols were in general more reactive than aromatic

alcohols under the same reaction conditions. A good yield

was observed for alcohols with C3–C5 (propanol–penta-

nol) but it was the lowest for Ar–C4 (4-phenyl-1-butanol,

9). The initial and average transesterification rate of this

relatively low yield reaction suggests that its final yield

could be increased by increasing the reaction time. To test

this hypothesis, the transesterification reaction of 4-phenyl-

1-butanol (9) was repeated at a prolonged reaction time.

The results of these experiments showed that under the

same reaction conditions, its yield increased from 42 to

56 % when the reaction time was increased from 24 to

60 h, after that almost no change in yield was observed.

Macquarrie et al. [1] immobilized SC on chitosan after

converting it into different forms (film, gel, fibers) or after

cross-linking with glutaraldehyde. The enzyme load ranged

from 0.34 to 65 mg/g of the biocatalyst but the activity of the

immobilized enzyme, which was calculated for peptide

synthesis in non-aqueous media, was 0.039–12.7 lmol m-

in-1 g-1 while for PyD (1), it was much higher

(38–81 lmol min-1 g-1) for the same enzyme but under

different reaction conditions and for a different reaction.

Table 1 Initial, average and transesterification activity of N-acetyl-L-phenylalanine ethyl ester (APAEE, 2) with different aliphatic and aromatic

alcohols after 24 h reaction at room temperature catalyzed by subtilisin Carlsberg (SC) immobilized on pentynyl dextran (PyD, 1)

Initial rate (lmol min-1) Average rate (lmol min-1) Transesterification activity (lmol min-1 g-1) Yield %

1-Propanol (3) 5.63 4.30 81 91

1-Butanol (4) 5.50 3.70 79 87

1-Pentanol (5) 5.12 4.29 71 81

1-Hexanol (6) 4.38 4.11 68 71

Benzyl alcohol (7) 4.20 3.26 63 62

2-Phenyl ethanol (8) 3.99 3.15 59 57

4-Phenyl-1-butanol (9) 2.55 2.01 38 42

Bioprocess Biosyst Eng

123

Recycling of biocatalyst

To implement the transesterification reaction at industrial

scale, where many cycles of high yield are required, the

stability and transesterification activity of the immobilized

enzyme should allow for repeated use [34, 35]. SC

immobilized on PyD (1) was repeatedly applied to the

transesterifications of APAEE (2) with 1-propanol (3) and

Fig. 2 RP-HPLC (C18 column)

chromatogram of the

transesterification reaction of N-

acetyl-L-phenylalanine ethyl

ester (APAEE, 2) with

1-hexanol at different reaction

times. Peaks at 3.06 and

17.28 min corresponds to N-

acetyl-L-phenylalanine ethyl

ester and N-acetyl-L-

phenylalanine propyl ester

respectively. Water:acetonitrile

(60:40, v/v), 2 mL/min, 30 �C,

UV 220 and 270 nm

Fig. 3 Final yield of the

transesterification of N-acetyl-L-

phenylalanine ethyl ester

(APAEE, 2) with different

aliphatic and aromatic alcohols

after 24 h in the reaction

catalyzed by protease

immobilized on pentynyl

dextran (PyD, 1)

Bioprocess Biosyst Eng

123

benzyl alcohol (7), which was selected because these

alcohols produced the highest yield in a reasonable time

(Table 1).

After completion of a cycle, the biocatalyst was filtered,

washed several times with THF, dried in air at room

temperature, and reused. The products were analyzed by

HPLC as described above. The biocatalyst was repeatedly

used for the transesterification of APAEE (2) with propanol

and benzyl alcohol over six cycles. In these experiments,

there was, in general, no loss in activity for both alcohols

(Fig. 4). For 1-propanol, the yield decreased from 92 to

81 % in third cycle (Fig. 4) but its recovery in the next

cycle indicates that this likely occured because of experi-

mental or instrumental error. For benzyl alcohol, a con-

tinuous increase in yield was observed and it increased

from 61 to 68 % in the first four cycles and remained

almost constant for the next two cycles (Fig. 4). This might

be due to slow adaptation of the most suitable changes and

configuration for the enzyme, resulting in higher activation

and higher yield than previous cycles in the same reaction

time.

Fluorescence spectroscopy

Protein fluorescence has been used as a measure of the

conformational state of a protein [36]. Most of the intrinsic

fluorescence is due to excitation of tryptophan (Trp) resi-

dues, which is strongly influenced by the proximity of other

residues. Therefore, the fluorescence emission spectra of

the Trp residues were analyzed to compare the tertiary

structure of native SC and SC immobilized on PyD (1). The

emission spectrum of the original SC in aqueous buffer was

obtained and the emission maximum of SC in the organic

solvent (THF) was found to be red-shifted (Fig. 5). The

fluorescence maximum wavelength was shifted from 305 to

321 nm, suggesting greater exposure of the Trp residues to

the solvent or partial denaturation of the tertiary structure

[37–39]. On the other hand, the fluorescence maximum of

the PyD immobilized SC was not significantly altered

although the absolute intensity was reduced because of its

solubility in THF [38]. This may have resulted because the

tertiary structure of SC was maintained after immobiliza-

tion on the PyD. These fluorescence spectroscopy experi-

ments demonstrated that the activity of the PyD

immobilized SC for the catalytic transesterification in THF

was different from the inactive free SC in THF.

Conclusions

Protease (Subtilisin Carlsberg, 7.0–14.0 units/mg) can be

immobilized on pentynyl dextran (PyD, 1, a derivative of

dextran with 500,000 average Mw, DSPy = 0.43) and can be

used for the transesterification of N-acetyl-L-phenylalanine

ethyl ester (2) with different aliphatic and aromatic alcohols

in THF. All alcohols used in this study show good to excellent

yield (42–92 %). The biocatalyst (protease immobilized on

PyD, 1) remained stable and active and could be applied

Fig. 4 Transesterification

reaction of N-acetyl-L-

phenylalanine ethyl ester

(APAEE, 2) with 1-propanol (3)

and benzyl alcohol (7) after

repeated use of the protease

immobilized on pentynyl

dextran (PyD, 1)

Bioprocess Biosyst Eng

123

repeatedly over several cycles while producing the same or

enhanced yield. Intrinsic fluorescence analysis also demon-

strated that the tertiary structure of the enzyme immobilized

on PyD (1) was retained in THF.

Acknowledgments Financial support of Konkuk University (KU

Brain pool) is greatfully acknowledged. This work is also supported

by the National Research Foundation of Korea Grant funded by the

Korean Government (NRF-2011-0024008 and NRF-2011-619-

E0002).

References

1. Macquarrie DJ, Bacheva A (2008) Efficient subtilisin immobili-

zation in chitosan, and peptide synthesis using chitosan-subtilisin

biocatalytic films. Green Chem 10:692–695

2. Alloue W, Destain J, El Medjoub T, Ghalfi H, Kabran P, Thonart

P (2008) Comparison of Yarrowia lipolytica lipase immobiliza-

tion yield of entrapment, adsorption, and covalent bond tech-

niques. App Biochem Biotechnol 150:51–63

3. Cabrera Z, Fernandez-Lorente G, Fernandez-Lafuente R, Palomo

JM, Guisan JM (2009) Novozym 435 displays very different

selectivity compared to lipase from Candida antarctica B

adsorbed on other hydrophobic supports. J Mol Catal B Enzym

57:171–176

4. Liu X, Guan Y, Shen R, Liu H (2005) Immobilization of lipase

onto micron-size magnetic beads. J Chromatog B 822:91–97

5. Gomes F, Silva G, Pinatti D, Conte R, de Castro H (2005) Wood

cellulignin as an alternative matrix for enzyme immobilization.

Appl Biochem Biotechnol 121:255–268

6. James J, Simpson BK, Marshall MR (1996) Application of

enzymes in food processing. Crit Rev Food Sci Nutr 36:437–463

7. Yoshimaru T, Matsumoto K, Kuramoto Y, Yamada K, Sugano M

(1997) Preparation of microcapsulated enzymes for lowering the

allergenic activity of foods. J Agric Food Chem 45:4178–4182

8. Santos AM, Montanez Clemente I, Barletta G, Griebenow K

(1999) Activation of serine protease subtilisin Carlsberg in

organic solvents: combined effect of methyl-b-cyclodextrin and

water. Biotechnol Lett 21:1113–1118

9. Zaks A, Klibanov AM (1988) The effect of water on enzyme

action in organic media. J Biol Chem 263:8017–8021

10. Griebenow K, Laureano YD, Santos AM, Clemente IM, Rodrı-

guez L, Vidal MW, Barletta G (1999) Improved enzyme activity

and enantioselectivity in organic solvents by methyl-b-cyclo-

dextrin. J Am Chem Soc 121:8157–8163

11. Khmelnitsky YL, Welch SH, Clark DS, Dordick JS (1994) Salts

dramatically enhance activity of enzymes suspended in organic

solvents. J Am Chem Soc 116:2647–2648

12. Lindsay JP, Clark DS, Dordick JS (2004) Combinatorial formu-

lation of biocatalyst preparations for increased activity in organic

solvents: salt activation of penicillin amidase. Biotechnol Bioeng

85:553–560

13. Ooe Y, Yamamoto S, Kobayashi M, Kise H (1999) Increase of

catalytic activity of a-chymotrypsin in organic solvent by co-

lyophilization with cyclodextrins. Biotechnol Lett 21:385–389

14. Ru MT, Hirokane SY, Lo AS, Dordick JS, Reimer JA, Clark DS

(2000) On the salt-induced activation of lyophilized enzymes in

organic solvents: effect of salt kosmotropicity on enzyme activ-

ity. J Am Chem Soc 122:1565–1571

15. Bacheva A, Isakov M, Lysogorskaya E, Macquarrie D, Philipp-

ova I (2008) Biocomposite of subtilisin Carlsberg with chitosan

as an effective biocatalyst for hydrolysis and synthesis of pep-

tides. Russ J Bioorg Chem 34:334–338

16. Hedstrom M, Plieva F, Galaev IY, Mattiasson B (2008) Mono-

lithic macroporous albumin/chitosan cryogel structure: a new

matrix for enzyme immobilization. Anal Bioanal Chem

390:907–912

Fig. 5 Fluorescence emission

spectra of free subtilisin

Carlsberg (SC) and SC

immobilized on pentynyl

dextran (PyD, 1). The emission

spectrum of SC solution in

50 mM Na phosphate buffer

(pH 7.7) was denoted by the

dot-dash line, SC in THF by the

dash line, and SC immobilized

on the PyD in THF by solid line

Bioprocess Biosyst Eng

123

17. Kise H, Hayakawa A (1991) Immobilization of proteases to porous

chitosan beads and their catalysis for ester and peptide synthesis in

organic solvents. Enzyme Microb Technol 13:584–588

18. Tahir MN, Bork C, Risberg A, Horst JC, Komoß C, Vollmer A,

Mischnick P (2010) Alkynyl ethers of glucans: substituent dis-

tribution in propargyl-, pentynyl- and hexynyldextrans and -am-

yloses and support for silver nanoparticle formation. Macromol

Chem Phys 211:1648–1662

19. Tankam PF, Mischnick P, Hopf H, Jones PG (2007) Modification

of methyl O–propargyl-d-glucosides: model studies for the syn-

thesis of alkynyl based functional polysaccharides. Carbohydr

Res 342:2031–2048

20. Tankam PF, Muller R, Mischnick P, Hopf H (2007) Alkynyl

polysaccharides: synthesis of propargyl potato starch followed by

subsequent derivatizations. Carbohydr Res 342:2049–2060

21. Naka K, Yamashita R, Nakamura T, Ohki A, Shigeru M, Aoi K,

Takasu A, Okada M (1998) Chitin-graft-poly(2-methyl-2-oxaz-

oline) enhanced solubility and activity of catalase in organic

solvent. Int J Biol Macromol 23:259–262

22. Yang G, Wu J, Xu G, Yang L (2009) Enhancement of the activity

and enantioselectivity of lipase in organic systems by immobili-

zation onto low-cost support. J Mol Catal B Enzym 57:96–103

23. Jiang Y, Guo C, Xia H, Mahmood I, Liu C, Liu H (2009)

Magnetic nanoparticles supported ionic liquids for lipase immo-

bilization: enzyme activity in catalyzing esterification. J Mol

Catal B Enzym 58:103–109

24. Ozmen EY, Sezgin M, Yilmaz M (2009) Synthesis and charac-

terization of cyclodextrin-based polymers as a support for

immobilization of Candida rugosa lipase. J Mol Catal B Enzym

57:109–114

25. Mariotti F, Tome D, Mirand PP (2008) Converting nitrogen into

protein—beyond 6.25 and Jones’ factors. Crit Rev Food Sci Nutr

48:177–184

26. Spurlin HM (1939) Arrangement of substituents in cellulose

derivatives. J Am Chem Soc 61:2222–2227

27. de Fuentes IE, Viseras CA, Ubiali D, Terreni M, Alcantara AR

(2001) Different phyllosilicates as supports for lipase immobili-

sation. J Mol Catal B Enzym 11:657–663

28. Tran DN, Balkus KJ (2011) Perspective of recent progress in

immobilization of enzymes. ACS Catalysis 1:956–968

29. Deng L, Tan T, Wang F, Xu X (2003) Enzymatic production of

fatty acid alkyl esters with a lipase preparation from Candida sp.

99-125. Eur J Lipid Sci Technol 105:727–734

30. Tahir MN, Adnan A, Stromberg E, Mischnick P (2012) Stability

of lipase immobilized on O–pentynyl dextran. Bioprocess Biosyst

Eng 35:535–544

31. Chang SW, Shaw JF, Yang KH, Chang SF, Shieh CJ (2008)

Studies of optimum conditions for covalent immobilization of

Candida rugosa lipase on poly(c- glutamic acid) by RSM. Bi-

oresour Technol 99:2800–2805

32. Tahir MN, Adnan A, Mischnick P (2009) Lipase immobilization

on O-propargyl and O-pentynyl dextrans and its application for

the synthesis of click beetle pheromones. Process Biochem

44:1276–1283

33. Chen B, Pernodet N, Rafailovich MH, Bakhtina A, Gross RA

(2008) Protein immobilization on epoxy-activated thin polymer

films: effect of surface wettability and enzyme loading. Langmuir

24:13457–13464

34. Chatterjee S, Barbora L, Cameotra S, Mahanta P, Goswami P

(2009) Silk-fiber immobilized lipase-catalyzed hydrolysis of

emulsified sunflower oil. Appl Biochem Biotechnol 157:593–600

35. Mateo C, Palomo JM, Fernandez-Lorente G, Guisan JM, Fer-

nandez-Lafuente R (2007) Improvement of enzyme activity,

stability and selectivity via immobilization techniques. Enzyme

Microb Technol 40:1451–1463

36. Vivian JT, Callis PR (2001) Mechanisms of tryptophan fluores-

cence shifts in proteins. Biophys J 80:2093–2109

37. Farivar F, Moosavi-Movahedi AA, Sefidbakht Y, Nazari K, Hong

J, Sheibani N (2010) Cytochrome c in sodium dodecyl sulfate

reverse micelle nanocage: from a classic electron carrier protein

to an artificial peroxidase enzyme. Biochem Eng J 49:89–94

38. Ganesan A, Moore BD, Kelly SM, Price NC, Rolinski OJ, Birch

DJS, Dunkin IR, Halling PJ (2009) Optical spectroscopic meth-

ods for probing the conformational stability of immobilised

enzymes. Chem Phys Chem 10:1492–1499

39. Pasta P, Riva S, Carrea G (1988) Circular dichroism and fluo-

rescence of polyethylene glycol-subtilisin in organic solvents.

FEBS Lett 236:329–332

Bioprocess Biosyst Eng

123