Tyrosinase catalyzed production of 3,4-dihydroxyphenylacetic acid using immobilized mushroom (Agaricus bisporus) cells and in situ adsorption

Article abstract of DOI:10.1016/j.molcatb.2015.11.012

3,4-Dihydroxyphenylacetic acid (DHPAA), a catechol derivative with proposed beneficial human health applications, was synthesized in this work from 4-hydroxyphenylacetic acid (HPAA) using an unpurified, tyrosinase-containing, cell preparation from the fruiting body of the edible mushroom Agaricus bisporus which were immobilized in silica alginate matrix capsules. The formation of DHPAA was equimolar to the conversion of HPAA, as long as ascorbic acid was present in amounts sufficient for reduction of o-quinones generated by oxidation of DHPAA. With a concentration of 5 mM HPAA and 5, 10, or 25 mM ascorbic acid, the maximum yields of DHPAA were 26, 36, or 56%, respectively. When aluminum oxide, pretreated with ammonium acetate, was added for an in situ adsorption of DHPAA, the yield obtained with 5 or 10 mM ascorbic acid was increased to 42 or 52%, respectively, with a reaction time reduced by ～25%. In contrast to experiments without in situ adsorption, the yield remained almost constant after depletion of ascorbic acid. After desorption, the concentration of DHPAA in the eluent was up to 32 times higher than the concentration of HPAA. The results presented here will be useful for the design of production and purification processes for DHPAA.

Full text of DOI:10.1016/j.molcatb.2015.11.012

Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic
journal homepage: www.elsevier.com/locate/molcatb
Tyrosinase catalyzed production of 3,4-dihydroxyphenylacetic acid
using immobilized mushroom (Agaricus bisporus) cells and in situ
adsorption
Markus Kampmann, Natascha Riedel, Yee Li Mo, Laura Beckers, Rolf Wichmann∗
Department of Biochemical and Chemical Engineering, TU Dortmund University, Emil-Figge-Str. 66, 44227 Dortmund, Germany
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 12 July 2015
Received in revised form 6 November 2015
Accepted 9 November 2015
Available online 15 November 2015
3,4-Dihydroxyphenylacetic acid (DHPAA), a catechol derivative with proposed beneficial human health
applications, was synthesized in this work from 4-hydroxyphenylacetic acid (HPAA) using an unpurified,
tyrosinase-containing, cell preparation from the fruiting body of the edible mushroom Agaricus bisporus
which were immobilized in silica alginate matrix capsules. The formation of DHPAA was equimolar to
the conversion of HPAA, as long as ascorbic acid was present in amounts sufficient for reduction of o-
quinones generated by oxidation of DHPAA. With a concentration of 5 mM HPAA and 5, 10, or 25 mM
ascorbic acid, the maximum yields of DHPAA were 26, 36, or 56%, respectively. When aluminum oxide,
pretreated with ammonium acetate, was added for an in situ adsorption of DHPAA, the yield obtained with
5 or 10 mM ascorbic acid was increased to 42 or 52%, respectively, with a reaction time reduced by ∼25%.
In contrast to experiments without in situ adsorption, the yield remained almost constant after depletion
of ascorbic acid. After desorption, the concentration of DHPAA in the eluent was up to 32 times higher
than the concentration of HPAA. The results presented here will be useful for the design of production
and purification processes for DHPAA.
Keywords:
3,4-Dihydroxyphenylacetic acid
Tyrosinase
Mushroom cells
Immobilization
Adsorption
© 2015 Elsevier B.V. All rights reserved.
1. Introduction
4-hydroxyphenylacetic acid (HPAA) using tyrosinase as a cat-
alyst [16,17]. Tyrosinase catalyzes the orthohydroxylation of
3,4-Dihydroxyphenylacetic acid (DHPAA) is a catechol deriva-
tive which has recently received considerable attention since it
has been demonstrated to protect against pancreatic ␤-cell dys-
function induced by high cholesterol. DHPAA has, therefore, been
suggested as a promising drug for prevention or delay of the tran-
sition from pre-diabetes to diabetes, which is currently a major
health issue worldwide [1]. In addition, DHPAA exhibits antipro-
liferative activity in prostate [2] and colon cancer cells [2–4] and
is, therefore, of great interest for human health. DHPAA has been
examined as precursor for the synthesis of hydroxytyrosol [5–7],
a substance with antioxidative properties [8,9], commonly consid-
ered as beneficial for human health [10–12], though DHPAA itself
has antioxidant activity too [13–15]. These beneficial medicinal
characteristics could increase the demand for DHPAA and economic
processes for its production.
monophenols (HPAA) to o-diphenols (DHPAA) and the oxidation
of o-diphenols to o-quinones using molecular oxygen [18]. The
formed o-quinones can be reduced to the o-diphenol precursor
by addition of ascorbic acid (Fig. 1) [16,17,19–21]. This reaction
pattern has also been examined for the tyrosinse catalyzed produc-
tion of other o-diphenols [16,17], including hydroxytyrosol [22] as
well as 3,4-dihydroxy-l-phenylalanine (l-DOPA) [23–29] which is
a prescribed drug for treatment of Parkinson’s disease.
However, the economic feasibility of these methods suffers from
the high cost of purified enzyme preparation when pure tyrosinase
is used. To overcome this limitation, enzyme containing protein
mixtures, or whole cell biocatalysts are a promising alternative
which can achieve the enzymatic conversion while avoiding costly
protein purification steps. An example of this approach is the use
of tyrosinase containing cells from the fruiting body of the edible
mushroom Agaricus bisporus, immobilized in silica alginate matrix
capsules for degradation of bisphenol A [30,31].
The catechol structure of DHPAA contains two adjacent
hydroxyl groups which can be generated enzymatically from
To date, this catalyst system has not been examined for pro-
duction of o-diphenols such as DHPAA. In particular, so far no
information about side reactions catalyzed by other enzymes in
the cells has been published. Formation of byproducts or secondary
∗
Corresponding author. Fax: +49 231 755 5110.
E-mail address: rolf.wichmann@bci.tu-dortmund.de (R. Wichmann).
http://dx.doi.org/10.1016/j.molcatb.2015.11.012
1381-1177/© 2015 Elsevier B.V. All rights reserved.

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M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
immobilized tyrosinase may be a promising process alternative.
Separation of DHPAA and tyrosinase may be accomplished by a
selective adsorption of DHPAA during the reaction (Fig. 1). Adsorp-
tion and desorption of DHPAA has been examined using aluminum
oxide as an adsorbent [43,44]. Both catechol derivatives [45] and
carboxyl groups [46], which are present in the molecular structure
of DHPAA as well as HPAA, can adsorb onto aluminum oxide. There-
fore, it is desirable to limit the adsorption of HPAA while promoting
adsorption of DHPAA to enable conversion of HPAA by the immobi-
lized enzyme. The adsorption of catechol derivatives or carboxylic
acids onto aluminum oxide has been demonstrated to be affected
by ammonium acetate [43]. However, these issues have not yet
been extensively characterized with respect to a targeted reduc-
tion of the adsorption of HPAA and the applicability for an in situ
adsorption in the tyrosinase catalyzed production of DHPAA.
In this report, fundamental insights gained on the tyrosinase
catalyzed production of DHPAA with mushroom cells immobilized
in silica alginate matrix capsules are presented. Selectivity stud-
ies and the effect of the amount of cells, ascorbic acid, as well
as the size of the matrix capsules are presented. In addition, the
adsorption of DHPAA and HPAA onto aluminum oxide is examined,
wherein, the extent of adsorption of HPAA depending on modifica-
tion of the adsorption system was investigated. The results of these
analyses were used to conduct an in situ adsorption of DHPAA dur-
ing tyrosinase catalyzed production from HPAA and presented as
an alternative approach to economic generation of this potentially
valuable compound.
Fig. 1. Hypothetical reaction scheme of the tyrosinase catalyzed production of
DHPAA with in situ adsorption and possible role of ascorbic acid. (A) Enzymatic
steps (transfer of H+ and formation of H2O not shown; adapted from Ref. [33]). (B)
Non-enzymatic steps: 1. Regeneration of DHPAA [16,17,19–21]. 2. Adsorption of
DHPAA (representative equation simplified from Ref. [45,46]). Em, met-tyrosinase;
Ed, deoxy-tyrosinase; Eox, oxy-tyrosinase; AA, ascorbic acid; DAA, dehydroascor-
bic acid; Q, o-quinone; EmHPAA, complex of Em with HPAA; EmDHPAA, complex of
Em with DHPAA; EmAA, complex of Em with AA; EoxAA, complex of Eox with AA;
2. Materials and methods
E
oxHPAA, complex of Eox with HPAA; EoxDHPAA, complex of Eox with DHPAA; AO,
2.1. Materials
aluminum oxide; AO-DHPAA, DHPAA adsorbed on AO.
Mushrooms (A. bisporus) at developmental stages 2–3 [47],
were acquired from a local supermarket. Alginic acid sodium
salt from brown algae (suitable for immobilization of microor-
ganisms), ammonium acetate (≥98% purity), formic acid (98%
purity), HPAA (98% purity) and Ludox® HS-30 colloidal silica 30%
(w/w) were purchased from Sigma–Aldrich GmbH (Steinheim,
Germany). Acetonitrile (≥99.9% purity), aluminum oxide (neutral,
for column chromatography), l-(+)-ascorbic acid (≥99% purity),
CaCl2·2H2O (≥99% purity), ethanol (≥99.8% purity), HCl (37%) and
NaOH (≥99% purity) were purchased from Carl Roth GmbH &
Co KG (Karlsruhe, Germany). DHPAA (98+% purity) and l-DOPA
(98+% purity) were purchased from Alfa Aesar GmbH & Co KG
(Karlsruhe, Germany), deuterated dimethyl sulfoxide (DMSO-d6,
99.8atom%D) from Armar Chemicals AG (Döttingen, Switzerland)
and 2-morpholinoethanesulfonic acid (MES, molecular biology
grade) from AppliChem GmbH (Darmstadt, Germany). All solutions
were prepared with double distilled deionized water (ddH2O).
products is undesirable, as it would reduce the yield of DHPAA and
pollute the desired product. This would require extensive purifica-
tion processes and consequently increase the overall process costs.
Therefore, the selectivity of the catalyst is of great importance and
deserves further investigation.
The catalytic cycle of tyrosinase includes three intermediate
states of the active site (Fig. 1): deoxy-tyrosinase (Ed), oxy-
tyrosinase (Eox) and met-tyrosinase (Em). Ed is able to bind
reversibly with molecular oxygen, producing the oxygenated form
Eox. Eox can act on monophenols to form o-diphenols or o-quinones,
while being converted to Em or Ed, respectively. Eox can also oxi-
dize o-diphenols with release of o-quinones and Em. Em can react
with o-diphenols as well, producing o-quinones and Ed, while Em
and monophenols form an inactive complex [32–35]. That means
after hydroxylation of a HPAA molecule, a formed DHPAA molecule
is oxidized to the o-quinone, before another HPAA molecule is
hydroxylated, unless the transformation of Em to Ed is taken over
by external reductants [16,33,36]. DHPAA can be regenerated from
the o-quinone by reduction with ascorbic acid [16,17,19–21], how-
ever, competes with the HPAA to be re-oxidized by Eox [32–35],
suggesting that high yields of DHPAA only can be attained at high
concentrations of ascorbic acid. The effect of ascorbic acid on tyrosi-
nase is currently debated in scientific literature, as several studies
observe no effect on tyrosinase activity [29,37–39], other reports
suggest that tyrosinase is inactivated by ascorbic acid [40,41], or
may even oxidize it [33,42] with reduction of Eox to Em or Em to
Ed (Fig. 1) [33]. In any case, a high concentration of DHPAA in the
reaction medium leads to an inhibition of DHPAA formation (accu-
mulation) and to an acceleration of DHPAA oxidation, thus limiting
the overall productivity in batch conversion reactions [27,28].
These limitations could hinder economic DHPAA production
processes, therefore, separation of generated DHPAA from the
2.2. Adsorption experiments
In order to examine the adsorption of DHPAA and HPAA, dif-
ferent amounts of aluminum oxide (0.3–0.6 g) were prepared by
addition of 2 ml ddH2O or ammonium acetate solution (0.2–6 M)
and gentle shaking for 1 min. After this pretreatment the adsorbent
was allowed to settle (5 min) and the supernatant was removed
pipetting. Since phosphate buffer has been demonstrated to pre-
vent adsorption of substances with a similar molecular structure
like l-DOPA and dihydroxyphenylserine on aluminum oxide [43],
test solutions with either DHPAA (5 mM), HPAA (5 mM), or both
(each 2.5 mM) were prepared with 0.1 M MES (pH 5–8 (adjusted to
with NaOH)). The concentrations of DHPAA and HPAA were cho-
sen in accordance with the expected concentrations during the
synthesis experiments (Section 2.6). The test solution (3 ml) was

M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
115
added to the wet adsorbent and stirred for 30 min with a mag-
2.6. Production of DHPAA
2.6.1. Reactor configuration
netic stirrer (300 rpm) at 20 ◦C. Then the adsorbent was allowed
to settle (5 min) and the supernatant was removed. Samples of the
supernatant were centrifuged (11,500 × g, 10 min) and the residual
concentrations of DHPAA and HPAA were quantified by HPLC.
Experiments for production of DHPAA were carried out in a glass
vessel equipped with a cannula positioned at the bottom of the
vessel for aeration and a magnetic stirrer operating at 300 rpm.
Preliminary experiments demonstrated that a suspension of alu-
minum oxide in combination with the stirrer bar had an abrasive
effect on the matrix capsules when they were added directly to
the vessel. To ensure the integrity of the matrix capsules, a stain-
less steel mesh (mesh size 0.5 mm) was positioned above the stirrer
bar. The matrix capsules were placed on the mesh so that they were
separated from the stirrer bar, but liquid, air bubbles and aluminum
oxide could pass.
Since the solubility of oxygen in water in an ambient atmo-
sphere (approximately 0.23 mM at 30 ◦C) [48], is much lower than
the used HPAA concentration (5 mM), the experiments were car-
ried out using an air flow rate of 150 ml/min to avoid an oxygen
deficiency and to support agitation of the liquid. This configuration
was used for both experiments with and without aluminum oxide
(with and without in situ adsorption, respectively) to ensure similar
reaction conditions.
2.3. Desorption experiments
After adsorption, the loaded adsorbent was washed with ddH2O
(10 ml/(g adsorbent)) under gentle shaking for 2 min. Then the
adsorbent was allowed to settle (5 min) and the supernatant was
removed. Desorption experiments were carried out by incubat-
ing the washed adsorbent with different volumes (3.3–10 ml/(g
adsorbent)) of ethanol, HCl (0.1–0.8 M), or mixtures of the two for
30–90 min under stirring (300 rpm) at 20 ◦C. Then the adsorbent
was allowed to settle again (5 min) and samples of the supernatant
were withdrawn and centrifuged (11,500 × g, 10 min). Depending
on the solvent used for desorption, the supernatant obtained after
centrifugation was diluted with a certain amount of NaOH solution
prior to HPLC analysis to avoid a damage of the HPLC column due
to acidic pH. After removal of the solvent from the adsorbent, these
steps were repeated for a second desorption cycle. The recovery of
DHPAA was defined as the percentage mass ratio of desorbed and
adsorbed DHPAA.
2.6.2. Production of DHPAA without in situ adsorption
To examine the tyrosinase catalyzed production of DHPAA, dif-
ferent amounts (0.1–0.9 g) of various matrix capsules (17–83 mg
cdw/g, d = 0.9–1.3 mm) were incubated in 9 ml substrate solution
(5 mM HPAA, 0–25 mM ascorbic acid, 0.1 M MES with pH 6 adjusted
to with NaOH) under stirring and aeration at 30 ◦C. At certain time
intervals samples of the solution were withdrawn and the concen-
trations of HPAA and DHPAA were quantified by HPLC. The yield
of DHPAA in these experiments was determined as the percent-
age ratio of the amount of DHPAA in the reaction medium and the
amount of HPAA initially used.
2.4. Preparation and immobilization of mushroom cells
Preparation and immobilization of mushroom cells were con-
ducted as previously described [30]. After lyophilization and
grinding, the mushroom cell preparation exhibited a tyrosinase
activity of 80 3 U/(g cell dry weight (cdw)) using l-DOPA as sub-
strate following previously described protocols [30]. Mushroom
powder (0.1 or 0.5 g) was added to 0.2 g sodium alginate and 10 ml
ddH2O containing 2.5% (w/w) Ludox® HS-30 (pH 6.8) and stirred
with an agitator. After the alginate was dissolved, the polymer solu-
tion was dropped into 2% (w/w) CaCl2 solution for gelation using a
diameter-controllable droplet generator [30]. The matrix capsules
were stored in ddH2O until use to avoid drying and shrinkage from
exposure to air. All capsule masses reported below refer to their wet
weight immediately after removal of external water by filtration.
2.6.3. Production of DHPAA with in situ adsorption
For production of DHPAA with in situ adsorption, 9 ml substrate
solution (adopted from Section 2.6.2) was added to 1.8 g aluminum
oxide prepared according to procedures in Section 2.2. The reac-
tions were started by addition of 0.9 g matrix capsules (17 mg
cdw/g, d = 1.3 mm) and carried out under stirring and aeration
at 30 ◦C. At designated time intervals the reactions were stopped
by removal of the matrix capsules. The adsorbent was allowed
to settle (5 min) and samples of the reaction medium were cen-
trifuged (11,500 × g, 10 min) before they were analyzed by HPLC.
After removal of the supernatant, the adsorbent was washed and
subjected to the desorption procedure described in Section 2.3. The
yield of DHPAA in these experiments was determined as the per-
centage ratio of the amount of DHPAA in the eluent after desorption
and the amount of HPAA initially used.
2.5. Characterization of matrix capsules
To estimate the cell content of the matrix capsules, it was
assumed that both the volume of the polymer solution and the
density of the matrix capsules remained unchanged when the cells
were suspended or immobilized. As demonstrated in the previ-
ous study [30], the cells and tyrosinase from fractured cells were
completely immobilized. The cell content of the matrix capsules
was calculated by dividing the cell concentration in the polymer
solution by the yield of the matrix capsules (0.6 g/(ml polymer
solution)) [30].
2.7. HPLC measurements
The diameter of the matrix capsules was determined utilizing
a Traveler SU 1071 USB microscope with Ulead Video Studio 7
SE VCD software (Supra Foto-Elektronik-Vertriebs-GmbH, Kaiser-
slautern, Germany) and graph paper as a reference. Photographs of
matrix capsules were processed by image analysis software, ImageJ
1.46p. Reported diameters (d) are the arithmetic mean of 50 ana-
lyzed matrix capsules, taking into account their smallest diameter
(dmin) and largest diameter (dmax) orthogonal to it. The aspect ratio
AR = dmin/dmax was at least 0.94.
Prior to HPLC analysis the samples were filtered through
0.2 ␮m PTFE filter. HPLC was conducted with a Knauer
a
Smartline series with detection at 227 nm, Eurospher II 100-
3C18 A (150 × 3 mm) column (Knauer GmbH, Berlin, Germany),
mobile phase water/acetonitrile/formic acid (volumetric ratio
87.3/12.5/0.2), flow rate 0.7 ml/min at 40 ◦C. The concentrations of
HPAA and DHPAA were quantified by comparing the retention time
and area of the peaks in the chromatograms with those of authentic
samples.

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M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
1.5
A
1.2
0.9
0.6
0.3
0
0
50
100
150
200
time [min]
5
4
3
2
1
0
B
0
50
100
150
200
time [min]
0.1 g (83 mg cdw/g)
0.9 g (83 mg cdw/g)
0.3 g (83 mg cdw/g)
0.9 g (17 mg cdw/g)
Fig. 2. Formation of DHPAA (A) and decrease in the concentration of HPAA (B) as a function of time, as well as the amount and the cell content of the matrix capsules (d = 1.3
mm) in 9 ml substrate solution (5 mM HPAA, 5 mM ascorbic acid, pH 6) at 30 ◦C. Dashed and solid lines were interpolated to illustrate the reaction progress in a qualitative
manner.
Bremen, Germany) with a SynergiTM Polar-RP (4 ␮m, 80 Å) column
(Phenomenex Ltd., Aschaffenburg, Germany). The mobile phase
consisted of solvent A (ddH2O + 0.2% formic acid) and solvent B
(acetonitrile) and the flow rate was 0.4 ml/min. The initial chro-
matographic conditions were 90% A and 10% B. After 1 min, B was
increased to 80% with a gradient time of 8 min and kept constant
2.8. Identification of DHPAA
Formation of DHPAA in the synthesis experiments (Section 2.6)
was confirmed by LC–MS and 1H NMR. For LC–MS, samples of the
reaction medium were analyzed with a TSQ Quantum UltraTM AM
Triple Quadrupole mass spectrometer (Thermo Finnigan GmbH,
1.4
1.2
1
d = 1.3 mm
d = 0.9 mm
0.8
0.6
0.4
0.2
0
0
20
40
60
80
100
120
time [min]
Fig. 3. Production of DHPAA as a function of time and the size of the matrix capsules (0.9 g, 17 mg cdw/g) in 9 ml substrate solution (5 mM HPAA, 5 mM ascorbic acid, pH 6)
at 30 ◦C.

M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
117
3
2.5
2
60
50
40
30
20
10
0
2.5 mM
5 mM
1.5
1
10 mM
25 mM
0.5
0
0
1
2
3
4
5
6
time [h]
Fig. 4. Effect of ascorbic acid on the production of DHPAA using 0.9 g matrix capsules (d = 1.3 mm, 17 mg cdw/g) in 9 ml substrate solution (5 mM HPAA, pH 6) at 30 ◦C.
for 2 min. After 11 min, B was returned to 10% and this composi-
tion was maintained for 4 min before the next injection. Detection
was carried out using an electrospray interface in negative or posi-
tive ionization mode. A negative fragment ion was observed at m/z
167.07 which suggested the chemical structure [DHPAA-H]−.
1H NMR was performed after lyophilization of the product that
has been isolated from the reaction medium using the adsorp-
tion and desorption procedure (Section 2.3). 1H NMR spectra
were recorded in DMSO-d6 using a Bruker DRX-400 spectrometer
(Bruker BioSpin GmbH, Rheinstetten, Germany).
decrease in the concentration of HPAA (Fig. 2B). After 3 h of reac-
tion, the concentration of DHPAA increased to almost 1.4 mM, while
the concentration of HPAA decreased from 5 mM to approximately
3.6 mM, indicating near equimolar conversion of HPAA to DHPAA.
Therefore, the selectivity of the catalyst of this reaction system may
be formally described as almost 100% under these conditions.
After removal of the immobilized cells from the reaction
medium, the concentrations of either HPAA or DHPAA did not
change, suggesting that the tyrosinase was completely retained in
the matrix capsules. Therefore, the use of the immobilized mush-
room cells instead of purified tyrosinase could be beneficial to
reduce the cost of the tyrosinase catalyzed production of DHPAA.
1H NMR (400 MHz, DMSO-d6): ı (ppm) 3.33 (s, 2H, CH2 ),
6.46–6.49 (dd, J = 7.91, 1.88 Hz, 1H, Ph H), 6.63–6.64 (d, J = 5.52 Hz,
1H, Ph H), 6.65 (s, 1H, Ph H), 8.80 (br s, 2H, Ph OH), 12.12 (br s,
1H, COOH).
3.1.2. Effect of the catalyst concentration
3. Results and discussion
For production of DHPAA, HPAA must diffuse into the matrix
capsules before it can be converted by tyrosinase in the immobi-
lized cells. Conversion of HPAA is dependent upon the diffusion
rate, enzymatic activity, and the cell content of the matrix capsules.
The three dimensional shape of the capsules may cause a concentra-
tion gradient of HPAA penetration, and therefore, have an influence
on its conversion efficiency to DHPAA. For this reason the effect of
the amount of cells was examined in two ways: by variation of the
amount of matrix capsules and by variation of the immobilized cell
content. Fig. 2A depicts that with greater amounts of matrix cap-
sules (83 mg cdw/g) the production of DHPAA was considerably
accelerated.
3.1. Production of DHPAA
3.1.1. Selectivity of the catalyst
Using 0.1 g matrix capsules (d = 1.3 0.08 mm) with a cell con-
tent of 83 mg cdw/g and a substrate solution with an equimolar
ratio of HPAA and ascorbic acid, almost 2.1 mg DHPAA was pro-
duced (Fig. 2A). Although the catalyst is contained within an
unpurified cell preparation, peaks of other substances were not
detected in the HPLC chromatograms of this experiment. Addition-
ally, the increase in the concentration of DHPAA correlates with the
100
80
60
40
20
0
DHPAA
HPAA
0 M
0.2 M 0.5 M 0.9 M 1.8 M 3.8 M
ammonium acetate concentration
6 M
Fig. 5. Effect of the pretreatment of aluminum oxide (0.6 g) with different concentrations of ammonium acetate on the subsequent adsorption of DHPAA or HPAA (5 mM in
3 ml 0.1 M MES (pH 6)).

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M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
However, in experiments with 0.3 g and 0.9 g matrix capsules,
respectively, when the smaller matrix capsules were used com-
pared to 0.33 mM and 0.85 mM obtained with the larger ones.
The smaller capsules have a higher surface area to volume ratio,
therefore, the diffusion distance to reach the immobilized tyrosi-
nase is shorter in the smaller matrix capsules resulting in the more
rapid production of DHPAA observed here. It is likely that some
degree of investigation is required to optimize the cell content or
the size of the matrix capsules to optimally balance cell concen-
tration and capsule size in order to adapt this method to industrial
application. However, the obtained results may serve as a basis for
reaction kinetic studies.
an apparent maximum concentration of DHPAA was obtained. The
time elapsed to reach comparable conversion amounts (60 min or
20 min) was proportional to the amount of matrix capsules, which
suggests a proportional conversion rate and that air flow rate and,
thereby oxygen consumption, was not limiting for the amount of
catalyst used. After the apparent maximum was reached, the con-
centration of DHPAA decreased and remained below 0.1 mM up to
3 h, although substrate was still available in considerable concen-
trations (Fig. 2B). It was also observed that the initially colorless
solution turned yellowish or brownish. When these samples were
analyzed by HPLC, additional peaks, which were not previously
observed, were detected in the chromatograms indicating the pres-
ence of other substances.
Tyrosinase catalyzes the orthohydroxylation of HPAA to DHPAA
and the oxidation of DHPAA to an o-quinone, which is reduced by
ascorbic acid to form DHPAA again (Fig. 1). However, since the
ascorbic acid is consumed in this reaction, the reaction rate of
the o-quinone reduction becomes lower after a certain number of
reduction cycles. This finally results in a lack of o-quinone reduc-
tion and subsequent loss of DHPAA. In LC–MS analysis, a positive
fragment ion was observed at m/z 167.06 which likely originated
from [o-quinone + H]+. The o-quinones can undergo polymeriza-
tion reactions resulting in further secondary products, this likely
accounts for the additional peaks observed in the chromatograms.
It was also observed that when the last samples were centrifuged, a
brown precipitate was obtained, suggesting the formation of insol-
uble polymeric products.
Using 0.9 g matrix capsules, approximately 0.39 mM DHPAA
was produced within the first 4 min of reaction. Simultaneously,
a rapid decrease in the initial concentration of HPAA from 5 mM
to approximately 4.2 mM was observed (Fig. 2B). The difference
in concentration change observed in this short time span is a
consequence of the high water content of the matrix capsules
(approximately 0.8 ml/g) [31], which results in a dilution of the
reaction medium depending on the amount of matrix capsules
applied. When 0.9 g matrix capsules without immobilized cells
were added to 9 ml substrate solution, the concentration of HPAA
decreased from 5 mM to 4.6 mM (data not shown). Therefore, the
HPAA conversion in the first 4 min was only 0.4 mM which is
consistent with the observed DHPAA formation (0.39 mM). The
minor differences in the apparent maximum DHPAA concentra-
tions observed with 0.3 g (1.38 mM) and 0.9 g matrix capsules
(1.3 mM) can be attributed to dilution effects as well in accordance
with previous observations of high selectivity of the catalyst.
In experiments with 0.3 g matrix capsules (83 mg cdw/g), the
apparent maximum DHPAA concentration was obtained after
60 min, however, in experiments with 0.9 g matrix capsules with a
cell content of 17 mg cdw/g, the DHPAA concentration was begin-
ning to decrease after the same reaction time (Fig. 2A), despite the
lower total mass of cells (15 mg cdw compared to 25 mg cdw). Due
to the large DHPAA concentration difference (1.38 mM compared
to 1 mM), this observation cannot be explained by dilution effects
alone, suggesting that the reaction rate was higher in the latter
experiment.
3.1.4. Effect of ascorbic acid
Since the decrease in concentration of DHPAA after a certain
reaction time was correlated with the depletion of ascorbic acid,
the molar ratio of ascorbic acid to HPAA was varied to examine the
effect on the formation of DHPAA.
In reactions without ascorbic acid, only low concentrations of
DHPAA (< 0.02 mM) were detected in the reaction medium over the
whole examined reaction time of 3 h. Given that a loss of DHPAA
was observed over long reaction times, Fig. 2, it was presumed that
DHPAA, which may have been formed from HPAA even without
ascorbic acid, was mainly oxidized to the o-quinone derivative so
that only traces of DHPAA were found in the solution. This would
confirm the need for a reducing agent to allow stable formation of
DHPAA.
Using 2.5 mM ascorbic acid the concentration of DHPAA
increased to approximately 0.7 mM within 14 min (Fig. 4), indicat-
ing the positive effect of a certain amount of ascorbic acid in this
reaction system. The higher the concentration of ascorbic acid used,
the longer and higher the concentration of DHPAA increased (Fig. 4).
When 5 mM ascorbic acid was used, a concentration of approxi-
mately 1.3 mM DHPAA was obtained after 52 min, which is a yield
of 26%. Using 10 mM or 25 mM ascorbic acid, the concentration of
DHPAA was increased to approximately 1.8 mM (36% yield) after
2.7 h or 2.8 mM (56% yield) after 4.4 h, respectively, likely resulting
from the higher capacity for reduction of o-quinones due to ascorbic
acid in solution.
Using ascorbic acid at concentrations from 2.5 up to 25 mM, the
initial increase in the concentration of DHPAA was 0.33 mM 8%
within the first 6 min in all experiments. Therefore, the reaction is
comparably effected by the concentration of ascorbic acid within
the examined range. Even the use of 50 mM ascorbic acid did not
change the initial reaction rate (data not shown). Although long-
term inactivation of tyrosinase by ascorbic acid [40,41] cannot be
excluded, no inhibitory effects were observed here, which is con-
sistent with other studies [29,37–39]. It is indeed possible that no
inhibition of tyrosinase activity occurred, although here, a slight
inhibition may not have been detected due to the diffusion limita-
tion of the reaction.
Although relatively high yields of DHPAA can be attained with
the immobilized mushroom cells, relatively large amounts of
ascorbic acid are required to sustain DHPAA in solution after its for-
mation (Fig. 4). After a certain reaction time, DHPAA formed in this
reaction is lost, likely due to its competitive oxidation, therefore
long reaction times are required to obtain high yields. When the
ascorbic acid is consumed the concentration of DHPAA decreases
rapidly.
It is likely that the production rate was limited by the diffusion
of reactants within the matrix capsules, suggesting that relatively
high production rates can be achieved with relatively low amounts
of immobilized mushroom cells. This was further examined by vari-
ation of the size of the matrix capsules as discussed below.
3.2. Adsorption of DHPAA and HPAA
3.1.3. Effect of the size of the matrix capsules
Fig. 3 depicts that the reaction rate was significantly increased
when the diameter of the matrix capsules (17 mg cdw/g) was
reduced from 1.3 0.08 mm to 0.9 0.06 mm. After 6 min and
22 min the concentration of DHPAA reached 0.46 mM and 1.06 mM,
In order to circumvent these disadvantages, a new production
concept with in situ adsorption of DHPAA was examined. To find a
suitable adsorption system, several adsorption experiments were
first carried out separately.

M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
119
100
80
60
40
20
0
2. desorption
1. desorption
0.1 M
0.4 M
0.4 M
0.8 M
0.8 M
10 ml/g 3.3 ml/g 10 ml/g 3.3 ml/g 10 ml/g
conditions of desorption with HCl
Fig. 6. Recovery of adsorbed DHPAA (load 4.2 mg/(g aluminum oxide)) after desorption with different concentrations and volumes of HCl. The volumes are given in relation
to the mass of the aluminum oxide.
First, 0.1 g aluminum oxide/ml at pH 6 resulted in adsorption of
the dissolved DHPAA (5 mM) up to 98%, whereas 0.2 g aluminum
oxide/ml resulted in adsorption of 100%, at 4.2 mg/(g adsorbent),
suggesting that aluminum oxide could be a suitable adsorbent for
the proposed reaction system. To maintain a quantitative adsorp-
tion of DHPAA, the concentration of aluminum oxide was kept at
0.2 g/ml.
However, as can be seen from Fig. 5 also a considerable adsorp-
tion of HPAA occurred with this system. Although the adsorbed
fraction, 68%, was lower than that of DHPAA (100%), HPAA adsorp-
tion was considered as undesirable, as this would mean loss
of substrate for catalysis. Since it has been reported that the
adsorption of l-DOPA, dihydroxyphenylserine, and other related
compounds on aluminum oxide was decreased in presence of
ammonium acetate [43], the adsorbent was treated with this sub-
stance prior to adsorption experiments.
As shown in Fig. 5 the extent of the adsorption of HPAA
decreased with higher ammonium acetate concentrations. By use
of 3.8 M ammonium acetate, only 7% of the HPAA was removed
from the solution, while the adsorbed fraction of DHPAA remained
almost unaffected (99%). In comparison, pretreatment of the adsor-
bent with 6 M ammonium acetate had only a slight effect on the
45
14
12
10
8
A
40
yield
molar ratio
35
30
25
20
15
10
5
6
4
2
0
0
0.33
0.66
1
1.5
2
3
4
time [h]
60
50
40
30
20
10
0
40
30
20
10
0
B
yield
molar ratio
0.5
1
1.5
2
2.5
3
4
time [h]
Fig. 7. Production of DHPAA with in situ adsorption using 0.9 g matrix capsules (d = 1.3 mm, 17 mg cdw/g), 9 ml substrate solution (5 mM HPAA, 5 mM (A) or 10 mM (B)
ascorbic acid, pH 6), 1.8 g aluminum oxide at 30 ◦C. The diagrams show the yield of DHPAA and the molar ratio of DHPAA and HPAA in the eluent after different reaction
times.

120
M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
reduction of the adsorption of HPAA (5%), but resulted in a slight
decrease in the adsorption of DHPAA (97%).
of DHPAA (<0.1 mM) were detected in the reaction medium. How-
ever, after the desorption step, considerable amounts of DHPAA
were obtained (Fig. 7).
The effect of ammonium acetate was confirmed with equimolar
mixtures (2.5 mM) of HPAA and DHPAA (data not shown) suggest-
ing a higher selectivity of the adsorbent prepared with ammonium
acetate for DHPAA. These results could be a consequence of pre-
viously described differences in the adsorption mechanisms of
carboxylic acids [46] and catechols [45]. Due to the favorable ratio
of adsorbed DHPAA and HPAA the aluminum oxide was conse-
quently prepared with 3.8 M ammonium acetate for all subsequent
experiments.
Using 5 mM ascorbic acid in the substrate solution, the maxi-
mum yield of DHPAA in the eluent was 42% (Fig. 7A), significantly
higher than the maximum yield obtained without in situ adsorp-
tion, 26% (Fig. 4), an increase of 61%. The yield was even higher
than the highest yield of DHPAA without in situ adsorption using
10 mM ascorbic acid (36%, Fig. 4). This strategy used considerably
less ascorbic acid, at least 50%, compared to strategies without
in situ adsorption.
The solution pH was also considered as an important parameter
for reaction optimization. Contradictory results have been reported
for the adsorption of catechol derivatives on aluminum oxide at dif-
ferent pH values [43,45,49,50]. However, in this study no significant
effect of the pH on the adsorption of these substances was observed
between pH 5 and pH 8 (data not shown). It is worth noting that the
adsorption can be accompanied by a shift of the reaction medium
pH [45]. Therefore, the use of higher concentrations of reactants
could require further investigations to determine the effect of pH
changes, or ion concentrations, however, this is beyond the scope
of this study. In accordance with the pH optimum of the catalyst
preparation [31] and the effective pH range of MES buffer, the pH
of the substrate solution was, therefore, maintained at pH 6 in all
subsequent analyses.
The maximum yield with in situ adsorption was achieved faster
than in the experiments with 10 mM ascorbic acid without adsorp-
tion. Here, maximum yield was observed at 2 h compared to 2.7 h,
representing a time saving of 25% provided that only the reaction
time is considered.
Using in situ adsorption, the produced DHPAA was not lost as
was observed in longer non in situ reactions. Here, 40% yield was
observed after 3 h and 38% after 4 h, in contrast to experiments
without in situ adsorption where a rapid decrease of the yield was
observed after the depletion of ascorbic acid (Fig. 4).
The positive effect of the in situ adsorption was confirmed using
10 mM ascorbic acid. Here, a yield of 52% was attained after 2.5 h
(Fig. 7B). In comparison to experiments without in situ adsorption,
where the yield was 36% (Fig. 4), this represents an increase of
44%. To achieve this yield without in situ adsorption, an ascorbic
acid concentration of almost 25 mM was required (Fig. 4). Also in
this setup, the observed product yield with in situ adsorption was
achieved faster than without (approximately 3.4 h) and the product
remained stable at 48% yield after 4 h.
3.3. Desorption of DHPAA
To achieve a high recovery of DHPAA after adsorption (load
4.2 mg/(g aluminum oxide)) different solvents were tested for their
desorption capacity.
It is likely that the separation of the produced DHPAA from
the immobilized tyrosinase restricts the oxidation of DHPAA and
consequent formation of o-quinones. Therefore, the number of
reduction cycles and the amount of ascorbic acid required to pro-
duce and to maintain a certain amount of DHPAA is reduced. This
results in higher yields of DHPAA which are maintained for a longer
time, since the adsorbed DHPAA is less prone to oxidation by the
immobilized enzyme. It may be the case that sequestration of the
DHPAA prevents competition of the enzyme for HPAA conversion,
however, this will require kinetic analysis of a purified enzyme.
Although it is uncertain if the adsorbent could wrest the DHPAA
from the tyrosinase or promote the release of DHPAA from the
enzyme before DHPAA is oxidized, it should be noted that in
absence of DHPAA or other reductants the catalytic cycle of tyrosi-
nase (Fig. 1) cannot be closed [16,32–36], leading to a standstill
of the reaction. However, as presented here, the in situ adsorption
concept may offer great potential for process intensification.
Taking into account the produced amounts of DHPAA and the
residual concentrations of HPAA in the reaction medium (data not
shown), the adsorbed fraction of the initial amount of HPAA was
determined to be less than 6%, which is in accordance with the
HPAA adsorption studies (Fig. 5). Therefore, only low amounts of
HPAA were found in the solvent after desorption (Fig. 7). The con-
centration of DHPAA in the eluent was up to 12 times (Fig. 7A)
or even 32 times (Fig. 7B) higher than the concentration of HPAA,
whereas in experiments without in situ adsorption, the concentra-
tion of DHPAA was always lower than the concentration of HPAA
until a yield of 50–56% was attained, when the molar ratio of DHPAA
and HPAA was 1–1.5. Although it is still unclear how much ammo-
nium acetate might have entered this system, the results reveal, at
least in part, a sequestering effect on the produced DHPAA. There-
fore, the results presented here may be also useful to facilitate the
separation of DHPAA and HPAA in subsequent downstream pro-
cesses.
It was found that ethanol was not able to elute DHPAA from
aluminum oxide. Mixtures of ethanol and 0.4 M HCl could recover
DHPAA in different quantities (data not shown), however, were
less effective than pure 0.4 M HCl. No difference in desorption
was observed for either 30 min or 90 min reaction time (data
not shown), suggesting that 30 min was sufficient for the high-
est possible elution under the examined conditions. Therefore,
the desorption step was set to 30 min and further experiments
were carried out with different concentrations and volumes of HCl
(Fig. 6).
When 0.1 M HCl was used in amounts of 10 ml/(g adsorbent) 55%
of the adsorbed DHPAA was eluted. Although 0.1 M HCl has been
used for elution in a previous study [43], the recovery was consid-
erably lower than the recovery obtained with 0.4 M (84%) or 0.8 M
HCl (90%) in the same proportion, suggesting a strong adsorption
of DHPAA onto aluminum oxide.
When the adsorbent was subjected to another desorption cycle,
the recovery obtained with 0.1 M HCl was increased by 20% to 75%
in total. Using 0.4 M HCl or 0.8 M HCl the recovery was increased
to 91% or 93%, respectively, demonstrating the feasibility of a high
recovery. The loss of approximately 7–9% could be explained by
irreversible binding of DHPAA or loss of loaded adsorbent during
the separation of liquid and adsorbent after the adsorption or sub-
sequent washing step. Similar total recoveries were also attained
when the volume of the 0.4 M or 0.8 M HCl was reduced to 3.3 ml/(g
adsorbent). Therefore, this proportion of 0.4 M HCl was adopted for
desorption after the DHPAA production with in situ adsorption.
3.4. Production of DHPAA with in situ adsorption
The findings from Sections 3.1–3.3 were applied to investigate
the tyrosinase catalyzed production of DHPAA with in situ prod-
uct separation. During the experiments, only low concentrations

M. Kampmann et al. / Journal of Molecular Catalysis B: Enzymatic 123 (2016) 113–121
121
4. Conclusion
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The authors are grateful to Stefan Konieczny for the assistance
with NMR analysis and Dr. Marc Lamshöft as well as Patrick Feike
from the Institute for Environmental Research (TU Dortmund Uni-
versity) for LC–MS analysis. We are grateful to Dr. Kyle J. Lauersen
for critical reading of the manuscript. The research leading to these
results has received funding from the Ministry of Innovation, Sci-
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CLIB-Graduate Cluster Industrial Biotechnology, contract no. 314-
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