Generation of norisoprenoid flavors from carotenoids by fungal peroxidases

Article abstract of DOI:10.1021/jf901438m

To biotechnologlcally produce norisoprenoid flavor compounds, two extracellular peroxidases (MsP1 and MsP2) capable of degrading carotenoids were isolated from the culture supematants of the basidiomycete Marasmlus scorodonlus (garlic mushroom). The encod

Full text of DOI:10.1021/jf901438m

J. Agric. Food Chem. 2009, 57, 9951–9955 9951
DOI:10.1021/jf901438m
Generation of Norisoprenoid Flavors from Carotenoids by
Fungal Peroxidases
†
K
ATERYNA
Z
ELENA,† BJORN
H
ARDEBUSCH,† BARBEL ULSDAU,† RALF G. BERGER
H ,
,§
AND
H
OLGER
Z
ORN
*
†Leibniz Universitat Hannover, Institut fur Lebensmittelchemie, Callinstratsse 5, D-30167 Hannover,
Germany, and §Justus-Liebig-Universitat Giessen, Institut fur Lebensmittelchemie und Lebensmittelbio-
technologie, Heinrich-Buff-Ring 58, D-35392 Giessen, Germany
To biotechnologically produce norisoprenoid flavor compounds, two extracellular peroxidases (MsP1
and MsP2) capable of degrading carotenoids were isolated from the culture supernatants of the
basidiomycete Marasmius scorodonius (garlic mushroom). The encoding genes were cloned from
genomic DNA and cDNA libraries, and databank homology searches identified MsP1 and MsP2 as
members of the so-called “DyP-type” peroxidase family. Wild type enzymes and recombinant perox-
idases expressed in Escherichia coli were employed for the release of norisoprenoids from various
terpenoid precursor molecules. Carotenes, xanthophylls, and apocarotenals were subjected to the
enzymatic degradation. Released volatile products were characterized by GC-FID and GC-MS, whereas
nonvolatile breakdown products were analyzed by means of HPLC-DAD and HPLC-MS. C13 noriso-
prenoids together with C10 products proved to be the main volatile degradation products in each case.
KEYWORDS: Basidiomycete; peroxidase; carotenoids; norisoprenoid flavors
INTRODUCTION
investigated approaches comprise the so-called co-oxidation with
the lipoxygenase/linoleic acid system, the release of bound pre-
cursors by means of glycosidases, and the use of plant cell
cultures. In the co-oxidation, the carotenoids are oxidized by free
radical species generated from essential fatty acids by lipoxygen-
ase catalysis (6). As none of these former approaches could be
commercialized so far, a biotic carotenoid degradation by fungal
enzymes was considered. In their natural environment, fungi are
in a close contact with various carotenes and xanthophylls, and
they thus should dispose of the enzymatic tools to degrade them.
A first carotenoid cleaving fungalenzyme was purifiedand cloned
from Pleurotus eryngii, which was erroneously named Lepista
irina in the original publication (7). Only recently, two extra-
cellular enzymes (MsP1 and MsP2) capable of degrading carote-
noids have been purified from culture supernatants of the
basidiomycete Marasmius scorodonius (garlic mushroom). The
genes encoding MsP1 and MsP2 were cloned and sequenced from
genomic DNA and cDNA libraries, and databank homology
searches identified the enzymes as members of the so-called
“DyP-type” peroxidase family (8). The aim of the present
investigation was to open new biotechnological routes to natural
flavor compounds derived from carotenoids. To that end, various
carotenes, xanthophylls, and apocarotenals were isolated from
plant matrices and subjected to enzymatic degradation by wild
type and recombinant MsP1 and MsP2. Volatile and nonvolatile
cleavage products were characterized.
In plants and fruits, numerous apocarotenoids are derived
from an enzyme-catalyzed excentric cleavage of the polyene
chains of carotenes and xanthophylls. Many of these cleavage
products, so-called norisoprenoids, and especially the carbon
13 compounds, act as potent flavor compounds. β-Ionone, a
molecule first found in the Bulgarian rose, has an odor threshold
of 0.007 ppm (1), and β-damascenone, another important com-
ponent of rose scents, is one of the most potent flavor-active
organic molecules (2). The first carotenoid cleaving plant enzyme
characterized on a molecular level was AtCCD1 (Arabidopsis
thaliana carotenoid cleavage dioxygenase 1) from A. thaliana.
When expressed in Escherichia coli, the recombinant enzyme
cleaved several carotenoids, including β-carotene, zeaxanthin,
lutein, and violaxanthin symmetrically at the 9,10 and 90,100
positions of the polyene backbone. A C14 dialdehyde and two
C13 norisoprenoid products were identified as reaction pro-
ducts (3). Convincing evidence for a dioxygenase mechanism of
AtCCD1 has been obtained from labeling experiments using
H218O and 18O2 (4). In the meantime, enzymes with properties
similar to those of AtCCD1 have been characterized from
numerousplants, for example, tomato(Lycopersiconesculentum),
crocus (Crocus sativus), petunia (Petunia hybrida), and wine (Vitis
vinifera L.) (5).
The occurrence of norisoprenoids in their producer plants is
typically restricted to trace amounts, and their extraction is often
expensive and laborious. Thus, biotechnological processes copying
natural carotenoid degradation have been envisaged. The best
MATERIALS AND METHODS
The M. scorodonius strain (CBS 137.86) was obtained from the
Dutch “Centraalbureau voor Schimmelcultures”, Baarn, The
Netherlands. Production and purification of wild type enzymes
*Corresponding author (telephone þ49 641-99 34 900; fax þ49 641-
99 34 909; e-mail holger.zorn@lcb.chemie.uni-giessen.de).
© 2009 American Chemical Society
Published on Web 10/09/2009
pubs.acs.org/JAFC

9952 J. Agric. Food Chem., Vol. 57, No. 21, 2009
Zelena et al.
Figure 1. Cleavage of β-carotene to flavor compounds (GC-MS chromatogram; CW 20): 1=2,6,6-trimethylcyclohexanone; 2=β-cyclocitral.
were performed as described in ref8, and recombinant MsP2 was
produced in cultures of E. coli according to the method of Zelena
et al. (9).
of the substrate concentration was performed by means of UV-vis
spectrophotometry using the parameters (solvents and wavelengths)
suggested by Britton et al. (10).
Substrates. β-Apo-80-carotenal and β-apo-120-carotenal were
obtained from BASF (Ludwigshafen, Germany). Lycopene and zea-
xanthin were donated from DSM(Delft, The Netherlands), and β-carotene
was purchased from Fluka (Seelze, Germany). Violaxanthin and neo-
xanthin were extracted from spinach and purified by preparative HPLC.
Therefore, 250 g of spinach was extracted with 3ꢀ250 mL of an acetone/
saturated NaHCO3 solution (4:1, v/v) in a kitchen blender. The combined
extracts were filtered through a Buchner funnel, and the solvent was
removed at 40 °C and 500 mbar in a rotary evaporator. After saturation
of the aqueous phase with NaCl, the xanthophylls were extracted with
3ꢀ100 mL of diethyl ether. The combined organic extracts were dried over
anhydrous sodium sulfate and concentrated, and ether residues were
removed under a stream of nitrogen. The resulting extract was redissolved
in 5 mL of acetone, filtered through a 0.45 μm membrane filter, and
subjected to semipreparative HPLC.
For semipreparative HPLC separation, a Nucleosil 120-5 column
(250 ꢀ 16 mm; Macherey & Nagel, Duren, Germany) with 5 μm C18
reversed phase material was used. The mobile phase consisted of mixtures
of methanol/water (80:20 v/v) (A) and ethyl acetate (B), starting with 80%
A, followed by a gradient to obtain 23% B after 5 min, 50% B after 40 min,
and 100% B after 50 min at a flow rate of 5.0 mL/min. The injection
volume was 500 μL. Detection was performed with a UV-1570 detector
(Jasco, Gross-Umstadt, Germany) at 450 nm. The respective violaxanthin-
and neoxanthin-containing fractions of several runs were combined and
concentrated in a rotary evaporator at 200 mbar and 40 °C. The aqueous
residue was extracted with ethyl acetate, and the organic extract was filled
to a final volume of 20 mL. The yieldsof neoxanthinand violaxanthin were
18.5 and 38.8 mg/kg, respectively.
Biotransformation. Substrate emulsions (0.01%) were prepared as
described in ref 11. The biotransformation was performed with 300 μg of
substrate in a total volume of 15 mL of sodium acetate buffer (50 mM, pH
5.0) for 60 min at 27 °C (150 rpm) and initiated by the addition of wild type
or recombinant enzyme preparation (12 mU) and 5 μL of 20 mM H2O2
solution. For the blanks, the enzyme samples were heat inactivated
(100 °C, 20 min) prior to the reaction.
Product Purification and Identification. The degradation products
were purified by SPE (Chromabond C18, Macherey & Nagel) and
identified by GC-MS by comparing their Kovats indices and mass spectra
with published data. Quantification was performed by GC-FID using (þ)-
R-terpineol (1.42 mM) as an internal standard.
After further purification of the extract by flash chromatography on
silica 60 (60-200 mesh, Merck, Darmstadt, Germany; elution with
pentane/diethyl ether 80:20, v/v), nonvolatile breakdown products were
analyzed by HPLC-DAD and HPLC-MS.
Gas Chromatography-Flame Ionization Detector (GC-FID).
High-resolution GC-FID using a polar phase was performed on a Trace
GC equipped with a DB-Wax column (30 mꢀ0.32 mm i.d., film thickness=
0.25 μm, SGE, Griesheim, Germany). A Fisons GC 8000, equipped with a
split-splitless injector (230 °C) and a flame ionization detector (300 °C),
was applied for quantitative analyses on a nonpolar phase. Separation of
volatiles was achieved on a DB-5 column (30 m ꢀ 0.25 mm i.d., film
thickness=0.25 μm, J&W, Folsom, CA). Hydrogen was used as carrier gas
at a flow rate of 2.5 mL/min, and the same temperature program was used
as for GC-MS.
Gas Chromatography-Mass Spectrometry (GC-MS). GC-MS
analysis using a polar phase was conducted on a Fisons GC 8000 equipped
with a (polyethylene glycol) ZB-Wax (30 mꢀ0.32 mm i.d., film thickness=
0.25 μm, Phenomenex, Torrance, CA) column connected to a Fisons
MD800 mass selective detector. GC-MS analysis using a nonpolar phase
was performed on an HP5890 series II GC equipped with a ZB-5MS
(30 mꢀ0.32 mm i.d., film thickness=0.25 μm, Varian, Palo Alto, CA)
column connected to an HP quadrupole mass spectrometer 5989. Both
GC-MS instruments were operated at 70 eV in the EI mode over the range
of 33-500 amu. Helium was used as the carrier gas at a flow rate of 3.1
mL/min (polar phase) or 3.3 mL/min (nonpolar phase), respectively. The
injection volume was 1 μL cool on-column (40 °C). The oven temperature
was held at 40 °C for 3 min, raised at 5 °C/min to a final temperature of
Lutein was released by saponification of marigold (Tagestes erecta)
oleoresin. To this end, 10 g of oleoresin was mixed with 50 mL of diethyl
ether and saponified at room temperature overnight with methanolic
potassium hydroxide (10% w/v). For complete removal of alkali, the
solution was washed with 3ꢀ50 mL of water; the organic layer was dried
over anhydrous sodium sulfate, filtered through a folded filter, and
evaporated to dryness. The extract was stored at -18 °C under nitrogen
until use.
UV-Vis Spectroscopy. Absorption spectra were recorded using a
::
Lambda 12 (Perkin-Elmer, Uberlingen, Germany) spectral photometer
equipped with thermostatable cell holder and magnetic stirrer. Calculation

Article
J. Agric. Food Chem., Vol. 57, No. 21, 2009 9953
Table 1. Tentatively Identified Volatile Degradation Products of Carotenes and Xanthophylls
yield
(mol %)
Kovats
index
substrate
degradation product
β-iononec
β-ionone-5,6-epoxide
dihydroactinidiolide
β-cyclocitralc
(mg L-1
)
GC-MS (m/z)
β-carotene
7.9 (0.6)
1.3 (0.1)
7.0 (0.5)
1.5 (<0.1)
2.5 (0.1)
1911a
1964a
2321a
1595a
1583a
192 (M•þ), 177 (100), 135, 107, 105
208 (M•þ), 135, 124, 123 (100)
180 (M•þ), 137, 111 (100), 110, 109
152 (M•þ), 137, 123, 109, 67 (100)
156 (M•þ), 128, 110, 95, 71 (100)
2-hydroxy-2,6,6-
trimethylcyclohexanone
lutein
3-hydroxy-R-ionone
3-hydroxy-β-ionone
11.0 (0.8)
6.3 (0.5)
1627b
1677b
208 (M•þ), 147, 125, 124, 109 (100)
208 (M•þ), 193 (100), 175, 147, 131
zeaxanthin
3-hydroxy-β-ionone
3-hydroxy-β-cyclocitral
5.7 (0.4)
traces
1677b
2346a
208 (M•þ), 193 (100), 175, 147, 131
168 (M•þ), 135 (100), 121, 107, 91
violaxanthin,
neoxanthin
3-hydroxy-β-ionone-5,6-epoxide
6.9 (0.5)
6.8 (0.5)
1688b
224 (M•þ), 125, 124, 123 (100), 109
lycopene
geranial
6-methyl-5-heptene-2-one
1.3 (<0.1)
7.2 (0.3)
1269b
1324a
152 (M•þ), 136, 121, 107, 69 (100)
126 (M•þ), 111, 108 (100), 93, 71
a M&N CW 20 M, 30 m ꢀ 0.32 mm i.d., 0.25 μm film thickness. b J&W DB 5, 30 m ꢀ 0.32 mm i.d., 0.25 μm. c Compound identified via reference standard.
Table 2. Tentatively Identified Volatile Degradation Products of Apocarote-
nals
substrate (mol %) (mg L-1
β-apo-80-carotenal β-apo-120-carotenal
)
product
β-iononea
β-ionone-5,6-epoxide
dihydroactinidiolide
5.4 (0.5)
1.9 (0.2)
2.1 (0.2)
1.4 (0.1)
1.6 (0.1)
3.2 (0.4)
1.0 (0.1)
2.2 (0.2)
traces
2-hydroxy-2,2,6-trimethylcyclohexanone
β-cycolcitrala
2.4 (0.2)
a Compound identified via reference standard.
phase Nautilus material including a precolumn (C18, 10ꢀ4 mm, 5 μm)
was used. The mobile phase consisted of mixtures of water/acetonitrile
(0-4 min 50:50 v/v; 4-20 min 0:100 v/v; 20-60 min isocratic 0:100 v/v) at
a flow rate of 1.0 mL/min. The injection volume was 20 μL. Detection
was performed with a SPD-M6A UV-vis photodiode array detector
(Shimadzu, Duisburg, Germany).
HPLC-Mass Spectrometry. HPLC-MS spectra were recorded on a
LCMS-QP8000R system (Shimadzu) in the APcIþ mode, using the mobile
phase and flow rate described above.
RESULTS AND DISCUSSION
Various carotenes, xanthophylls, and apocarotenals were sub-
jected to enzymatic degradation by the M. scorodonius perox-
idase. The biotransformation was performed at pH 5.0, where
MsP1 exhibited maximal catalytic activity (11). Although the
dimeric enzyme MsP1 turned out to be rather thermostable up to
temperatures of about 65 °C (12), a reaction temperature of 27 °C
was chosen to avoid losses of the released volatiles. Under these
conditions, all of the substrates were readily degraded within 60 min,
and C13 norisoprenoids together with C10 products proved to be
the main volatile degradation products in each case (Figure 1 and
Table 1). None of the degradation products was detected when
heat-inactivated enzyme samples were employed as blanks.
A potential mechanism for the carotenoid cleavage between C9
and C10 could start with the abstraction of a hydrogen atom from
the allylic methyl group, resulting in a resonance-stabilized
carbon radical. Hydroperoxides may be formed intermediately
through reaction with oxygen, and subsequent Hock cleavage
would yield two carbonyl compounds (7).
Figure 2. Norisoprenoid compounds released from carotenes and xantho-
phylls by M. scorodonius peroxidase catalysis.
240 °C (polar phase) or 280 °C (nonpolar phase), respectively, and held
constant at the final temperature for 10 min. Linear retention indices (RIs)
were calculated according to the Kovats method using n-alkanes (C7-C28
)
as external references. Mass spectral identification was completed by
comparing spectra with commercial mass spectral databases Wiley, NIST,
and LIBTX and by comparison with authentic reference standards.
β-Ionone (95%) and R-terpineol (puriss.) were obtained from Fluka,
and β-cyclocitral (>90%) was purchased from Sigma-Aldrich.
High-Performance Liquid Chromatography with Diode Array
Detection (HPLC-DAD). For HPLC separation, a Nucleosil CC 100-5
analytical column (250ꢀ4 mm; Macherey & Nagel) with 5 μm C18 reversed

9954 J. Agric. Food Chem., Vol. 57, No. 21, 2009
Zelena et al.
parts per million concentrations only, all of the carotenoids were
readily degraded under the experimental conditions chosen.
Whereas the cleavage of zeaxanthin, lycopene, and neoxanthin
amounted to about 30%, almost 60% of the β-carotene was
degraded within 60 min under the same conditions. Considering
the obtained yields of volatiles and the presence of only trace
amounts of nonvolatile degradation products, there is still a
significant gap in the mass balance. Further investigations will
be necessary to close this gap and to fully understand the reaction
pathway.
Data on carotenoid degradation by peroxidases are rather
scarce. Kanner and Mendel (18) identified a carotenoid
bleaching enzyme in aqueous paprika extracts, which showed
typical characteristics of plant peroxidases. A carotenoid
degrading peroxidative activity was found in solubilized
thylakoid membranes of olives (19), and at least partial
degradation of β-carotene was observed with soybean and
horseradish peroxidase as well as with lactoperoxidase (20).
Different from the highly selectively acting plant carotenoid
cleavage enzymes, the peroxidase-catalyzed reaction resulted
in a broader spectrum of cleavage products. When an enzyme
model based on a ruthenium tetramesitylporphyrin catalyst
was used to degrade β-carotene, similar product spectra were
obtained (21, 22).
In conclusion, the extracellular peroxidases of M. scorodonius
efficiently degraded carotenoids to norisoprenoid flavor com-
pounds. The H2O2 required for the catalytic activity of the
peroxidases may be supplemented or generated in situ by the
addition of glucose and glucose oxidase. Apart from the produc-
tion of “bioflavors”, the novel enzymes could become interesting
tools in detergents and food-bleaching applications (23).
Figure 3. Enzymatic cleavage of β-carotene by extracellular enzymes of
M. scorodonius; HPLC-MS chromatogram (1 = β-apo-140-carotenal; 2 =
β-apo-120-carotenal; 3=β-apo-100-carotenal; 4=β-carotene-monoepoxide;
5=β-carotene-5,6-epoxide).
Table 3. Nonvolatile Degradation Products of Carotenes and Xanthophylls
Tentatively Identified by HPLC-DAD and HPLC-MS Analyses
product retention
time (min)
substrate
[M þ Hþ]
tentative identifiaction
β-carotene
21.3
23.5
24.4
33.0
40.1
311
351
377
553
553
β-apo-140-carotenal
β-apo-120-carotenal
β-apo-100-carotenal
β-carotene-monoepoxide
β-carotene-5,6-epoxide
lutein/zeaxanthin
15.5
19.5
20.8
23.5
40.9
327
367
393
433
585
3-hydroxy-apo-140-carotenal
3-hydroxy-apo-120-carotenal
3-hydroxy-apo-100-carotenal
3-hydroxy-apo-80-carotenal
lutein/zeaxanthin epoxide
violaxanthin/
neoxanthin
12.4
16.5
17.7
343
383
409
3-hydroxy-β-apo-140-carotenal-
5,6-epoxide
3-hydroxy-β-apo-120-carotenal-
5,6-epoxide
3-hydroxy-β-apo-100-carotenal-
5,6-epoxide
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Received for review April 30, 2009. Revised manuscript received August
6, 2009. Accepted September 10, 2009. Support of the work by the
Deutsche Forschungsgemeinschaft” (ZO 122/1-2) is gratefully
acknowledged.