Conversion of pharmaceuticals and other drugs by fungal ... - IHI Zittau

Conversion of pharmaceuticals and other drugs by
fungal peroxygenases
Umsetzung von Pharmazeutika und psychoaktiven
Substanzen mit pilzlichen Peroxygenasen
Vom Institutsrat des Internationalen Hochschulinstitutes Zittau und der
Fakultät Mathematik und Naturwissenschaften der Technischen Universität
Dresden
genehmigte
DISSERTATION
Zur Erlangung des akademischen Grades
Doctor rerum naturalium
(Dr.rer.nat.)
von
geboren am
vorgelegt
Marzena Poraj-Kobielska, Dipl.-Ing. (Umwelttechnik)
25 Juli 1982 in Katowice (Polen)
Gutachter:
Herr Prof. Dr. Martin Hofrichter (TU Dresden-IHI Zittau)
Herr Prof. Dr. Kenneth E. Hammel (University of Madison)
Herr Prof. Dr.-Ing. Korneliusz Miksch (Politechnika Śląska,
Gliwice)
Tag der Verteidigung: 26 April 2013

Conversion of pharmaceuticals and other drugs by
fungal peroxygenases
Approved by the Council of International Institute Zittau and the Faculty of
Science of the TU Dresden
Academic Dissertation
Doctor rerum naturalium
(Dr.rer.nat.)
by
born on
Marzena Poraj-Kobielska, Dipl.-Ing. (Environmental Engineering)
July 25, 1982 in Katowice (Poland)
Reviewers: Prof. Dr. Martin Hofrichter (TU Dresden-IHI Zittau)
Prof. Dr. Kenneth E. Hammel (University of Madison)
Prof. Dr.-Ing. Korneliusz Miksch (Politechnika Śląska, Gliwice)
Day of defence: April 26, 2013

CONTENTS
Contents
Contents…………………………………………………………………………………….I
List of Abbreviations…………………………………………………………………….IV
Zusammenfassung……………………………………………………………………….VI
Summary……………………………………………………………………………..…VIII
Streszczenie………………………………………………………………………………..X
1 Introduction ................................................................................................................... 1
1.1 Physiological and environmental aspects of pharmaceuticals............................... 1
1.1.1 Pharmaceuticals............................................................................................. 1
1.1.2 Pharmacokinetics and Metabolism................................................................ 2
1.1.3 Pharmaceutical drugs development and studies ............................................ 5
1.1.4 Environmental aspects................................................................................... 6
1.2 Cytochrome P450 enzymes ................................................................................. 10
1.2.1 Classification of electron transport systems of P450s................................. 13
1.2.2 Bacterial P450s............................................................................................ 15
1.2.3 Plant P450s .................................................................................................. 16
1.2.4 Fungal P450s ............................................................................................... 17
1.2.5 Animal P450s .............................................................................................. 20
1.2.6 Engineered P450s ........................................................................................ 21
1.2.7 Catalyzed reactions...................................................................................... 22
1.2.8 Reaction mechanism of P450s..................................................................... 23
1.3 Peroxygenases ..................................................................................................... 27
1.3.1 Fungal unspecific peroxygenases ................................................................ 30
1.4 Aims and objectives............................................................................................. 31
2 Materials and Methods ................................................................................................ 33
2.1 Chemicals and reactants ...................................................................................... 33
2.1.1 Pharmaceuticals and their metabolites ........................................................ 33
2.1.2 Drugs and their metabolites......................................................................... 33
2.1.3 Steroids and anabolic agents........................................................................ 34
2.1.4 Benzodioxole derivatives ............................................................................ 34
2.2 Fungal cultivation and purification of peroxygenases......................................... 34
2.2.1 Agrocybe aegerita and its peroxygenase..................................................... 34
2.2.2 Coprinellus radians and its peroxygenase................................................... 38
2.2.3 Marasmius rotula and its peroxygenase...................................................... 41
2.3 UV-Vis spectroscopy........................................................................................... 42
2.3.1 Veratryl alcohol assay for peroxygenases ................................................... 42
2.3.2 ABTS assay for laccase ............................................................................... 43
2.3.3 Nitrobenzodioxole assay for peroxygenases ............................................... 43
2.3.4 Determination of protein concentration....................................................... 45
2.3.5 Difference spectra........................................................................................ 46
2.4 Standard reaction conditions for peroxygenase conversions............................... 46
2.5 Product identification .......................................................................................... 47
2.5.1 HPLC-Method I........................................................................................... 47
2.5.2 UHPLC-Method .......................................................................................... 47
2.5.3 HPLC-Method for chiral separation............................................................ 47
2.5.4 HPLC-MS Method I .................................................................................... 47
2.5.5 HPLC-MS Method II................................................................................... 48
2.5.6 HPLC-MS Method III ................................................................................. 48
2.5.7 Mass spectrometry for HPLC-MS methods ................................................ 48
I

2.5.8 IC-Method for organic acids ....................................................................... 48
2.6 Stoichiometrical analyses.................................................................................... 48
2.6.1 Sildenafil ..................................................................................................... 48
2.6.2 Safrole ......................................................................................................... 49
2.7 Enzyme kinetics .................................................................................................. 49
2.7.1 Pharmaceuticals........................................................................................... 49
2.7.2 Nitrobenzodioxole....................................................................................... 49
2.8 Experiments with 18 O-isotopes............................................................................ 50
2.8.1 Pharmaceuticals........................................................................................... 50
2.8.2 Nitrobenzodioxole....................................................................................... 50
2.9 Deuterium isotope effect experiment .................................................................. 50
2.10 Experiment with different peroxides and peroxide sources ................................ 51
2.11 In-vivo incubation experiment............................................................................. 51
2.12 Conversion of propranolol at environmentally relevant concentration............... 53
2.13 Long term experiments........................................................................................ 53
3 Results ......................................................................................................................... 55
3.1 Aromatic ring hydroxylation............................................................................... 55
3.1.1 Ring hydroxylation at pharmaceuticals....................................................... 55
3.1.2 Ring hydroxylation at drugs of abuse.......................................................... 58
3.2 Oxidation of alkyl side chains............................................................................. 59
3.3 O-Dealkylation .................................................................................................... 62
3.3.2 Ester cleavage.............................................................................................. 64
3.4 N-dealkylation ..................................................................................................... 64
3.5 O-Demethylenation ............................................................................................. 68
3.6 Oxidation of steroids ........................................................................................... 69
3.7 Stoichiometrical analyses.................................................................................... 71
3.7.1 Sildenafil ..................................................................................................... 71
3.7.2 Safrole ......................................................................................................... 72
3.8
18 O-Isotope incorporation.................................................................................... 72
3.9 Kinetics experiments ........................................................................................... 73
3.9.1 Michaelis-Menten kinetics .......................................................................... 73
3.9.2 Bisubstrate kinetics...................................................................................... 74
3.10 Deuterium isotope effect experiments................................................................. 75
3.11 Spectrophotometric nitrobenzodioxole assay...................................................... 75
3.12 Conversion of a set of human P450 substrates.................................................... 78
3.13 Difference spectroscopy...................................................................................... 81
3.14 Different peroxides and peroxide-generating systems ........................................ 82
3.15 In-vivo experiments ............................................................................................. 83
3.16 Tests with environmentally relevant concentrations of propranolol ................... 85
3.17 Long-term experiments ....................................................................................... 86
3.17.1 Pre-study with Vivaspin® (fed-batch method) ........................................... 86
3.17.2 Long-term experiment using a hollow fiber cartridge................................. 87
3.18 Scope of peroxygenase reactions concerning pharmaceuticals and drugs.......... 88
4 Discussion ................................................................................................................... 90
4.1 Preparation of human drug metabolites............................................................... 90
4.2 Mechanistic studies ............................................................................................. 99
4.2.1 Source of oxygen and stoichiometry........................................................... 99
4.2.2 Deuterium isotope effect ........................................................................... 100
4.2.3 Kinetics...................................................................................................... 100
4.2.4 Assays for human cytochrome P450 activities.......................................... 102
4.2.5 Peroxides and peroxides generation systems ............................................ 104
4.2.6 Difference binding spectra ........................................................................ 106
4.2.7 Phenoxyl radical scavenger....................................................................... 106
4.2.8 Stoichiometry and proposed mechanism of demethylenation................... 107
II

CONTENTS
4.3 NBD assay ......................................................................................................... 109
4.4 Long term and in-vivo experiments................................................................... 110
4.5 Homology model structures of APOs................................................................ 113
4.6 Key findings ...................................................................................................... 120
4.7 Outlook .............................................................................................................. 123
5 References ................................................................................................................. 125
6 Appendix ................................................................................................................... 143
6.1 Aromatic ring hydroxylation ............................................................................. 143
6.2 Oxidation of alkyl side chains ........................................................................... 144
6.3 O-Dealkylation .................................................................................................. 144
6.4 N-dealkylation ................................................................................................... 145
6.5 Oxidation of steroids ......................................................................................... 145
6.6 O-Demethylenation ........................................................................................... 146
6.7 Spectrophotometric nitrobenzodioxole assay.................................................... 146
6.8 Stoichiometrical analyses .................................................................................. 147
6.9 Experiment with propranolol in environmentally relevant concentrations ....... 148
6.10 List of publications ............................................................................................ 150
III

LIST OF ABBREVIATIONS
List of Abbreviations
Aae Agrocybe aegerita
Ala Alanine
APOs aromatic/unspecific peroxygenases
Arg Arginine
BPs By-products
Cfu Caldariomyces (Leptoxyphium) fumago
CPO Chloroperoxidase
Cra Coprinellus radians
Cve Coprinopsis verticillata
DAD Diod array detector
DNPH 2,4-Dinitrophenylhydrazine (Brady's reagent)
EC Enzyme Commission
ESI Electrospray Ionization
FPLC Fast Protein Liquid Chromatography
Glu Glutamic acid
Gly Glycine
HDMs Human drug metabolites
HPLC High Performance Liquid Chromatography
IC
Ion exchange Chromatography
IEF Isoelectric Focussing
Ile
Isoleucine
k cat
k cat /K m
kDa
((k H /k D ) obs )
Leu
LRET
mCPBA
MEA
MPO
Mro
IV
turnover number
catalytic efficiency
kilo Dalton
observed deuterium isotope effect
(k H /k D ) intrinsic deuterium isotope effect
K m
Michaelis-Menten-(Henri) constant
Leucine
Long-Range Electron Transfer
meta-Chloroperbenzoic acid
Malt-extract agar
Myeloperoxidase
Marasmius rotula

LIST OF ABBREVIATIONS
MS
NADH
NADPH
NBD
NSAIDs
P450 BM3
P450 BSβ
P450 cam
P450 foxy
P450 lun
P450 nor
P450s
P450 SPα
Phe
pI
PPs
Pro
SDS-PAGE
SEC
Ser
Thr
Tyr
UHPLC
UV
VA
Val
Vis
WWTPs
Mass Spectrometry
Nicotinamide Adenine Dinucleotide
Nicotinamide Adenine Dinucleotide Phosphate
Nitrobenzodioxole
Non-steroidal anti-inflammatory drugs
P450-reductase fusion enzyme from Bacillus megaterium
Fatty acid peroxygenase from Bacillus subtilis
Camphor 5-Monooxygenase from Pseudomonas
P450 from Fusarium oxysporum
P450 from Curvularia lunata
Nitric oxide reductase
Cytochrome P450 Monooxygenases
Fatty acid peroxygenase from Sphinomonas paucinobilis
Phenylalanine
Isoelectric point
Pharmaceutical products
Proline
Sodium dodecyl sulfate polyacrylamide
Size Exclusion Chromatography
Serine
Threonine
Tyrosine
Ultra High Performance Liquid Chromatography
Ultraviolet
Veratryl alcohol
Valine
Visible
Waste Water Treatment Plants
V

SUMMARY
Zusammenfassung
In den letzten Jahren sind Pharmazeutika und deren Metabolite mehr und mehr in den
Fokus der Wissenschaft gerückt. Diese Substanzen sind aufgrund ihrer physiologischen
und toxikologischen sowie ökotoxikologischen Wirkungen im menschlichen Körper bzw.
in der Umwelt von besonderem Interesse. Cytochrom-P450-Enzyme (P450s) spielen eine
Schlüsselrolle bei der Umsetzung und Detoxifizierung bioaktiver Substanzen, darunter
vieler Pharmazeutika und Drogen. Es handelt sich bei diesen Enzymen in erster Linie um
Monooxygenasen, die intrazellulär lokalisiert und relativ instabil sind; sie benötigen
komplexe, teure Kofaktoren und sind nur unter hohem Aufwand zu reinigen, was ihre
Anwendung in isolierter Form insgesamt erschwert. Die hier durchgeführten
Untersuchungen zu pilzlichen Peroxygenasen haben gezeigt, dass diese Enzymsubklasse
(EC 1.11.2.x) ein hohes Oxyfunktionalisierungspotenzial besitzt und eine Vielzahl P450-
typischer Reaktionen zu katalysieren vermag. Peroxygenasen sind extrazelluläre, d.h.
sekretierte Pilzenzyme, die eine hohe Stabilität aufweisen und lediglich ein Peroxid als
Kosubstrat benötigen. Die unter Verwendung der unspezifischen/aromatischen
Peroxygenasen (APOs) von Agrocybe aegerita, Coprinellus radians und Marasmius rotula
gewonnenen Ergebnisse belegen, dass APOs verschiedene H 2 O 2 -abhängige
Monooxygenierungen von Pharmazeutika und psychoaktiven Substanzen realisieren. Dazu
gehören i) die Monooxygenierung von Aromaten, ii) die benzylische Hydroxylierung von
Toluolderivaten, iii) die O-Dealkylierung verschiedener Etherstrukturen einschließlich der
Spaltung von Benzodioxolen (O-Demethylenierung) und Estern sowie iv) die N-
Dealkylierung von sekundären und tertiären Aminen. Die untersuchten Peroxygenasen
wiesen teilweise deutliche Unterschiede im Substratspektrum und den präferierten
Oxidationspositionen auf. Dieser Befund eröffnet die Möglichkeit, zukünftig einen
„enzymatischen Werkzeugkasten“ auf Basis pilzlicher Peroxygenasen für die
Oxyfunktionalisierung von pharmazeutisch relevanten Wirkstoffen zu entwickeln.
Mechanistische Experimente zeigten, dass (1) die Monooxygenierungen stets unter Einbau
eines aus dem Peroxid stammenden Sauerstoffatoms erfolgen, (2) die Deethylierung von
Phenacetin-d1 einen Deuteriumisotopeneffekt ähnlich dem der P450s aufweist, (3) die
katalytischen Effizienzen für die untersuchten Oxidationen im gleichen Bereich wie die der
P450s liegen (wobei die k cat - und K m -Werte deutlich höher ausfallen), (4) die kinetischen
Untersuchungen zur Oxidation von Nitro-1,3-Benzodioxol parallele Verläufe der
ermittelten Ausgleichsgeraden in der doppelt reziproken Darstellung ergaben, was für
einen “Ping-Pong-Mechanismus“ spricht, (5) sich das Substratspektrum und die
Aktivitätsmuster der APOs in einem weiten Bereich mit denen der wichtigsten
VI

SUMMARY
menschlichen P450s decken sowie dass (6) die in Bindungsstudien gewonnenen
Differenzspektren denen des Phenoltyps der P450s entsprechen. Desweiteren erwiesen sich
APOs in Langzeitexperimenten über zwei Wochen als stabil und aktiv und sie waren in der
Lage, Pharmazeutika in umweltrelevanten Konzentrationen (ppb-Bereich) zu oxidieren.
All die genannten Eigenschaften legen nahe, dass APOs eine interessante Alternative zur
enzymatischen Umsetzung von Pharmazeutika sowie zur Herstellung von humanen
Pharmazeutika-Metaboliten darstellen, die z.B. Einsatz in der medizinischpharmakologischen
Forschung oder im Umweltbereich (Entfernung von Pharmazeutika
aus Umweltmedien) finden könnten.
VII

SUMMARY
Summary
Over the recent years, increasing scientific attention has been paid to pharmaceuticals,
other drugs and their metabolites. These substances are of particular interest because of
their physiological, toxicological and ecotoxicological effects in the human body and
respectively in the environment. Cytochrome P450 enzymes (P450s) play a key role in the
conversion and detoxification of bioactive compounds including many pharmaceuticals
and drugs. Most of these enzymes belong to the monooxygenases; they are intracellular
and rather unstable biocatalysts that are difficult to purify and require expensive, complex
cofactors, which alltogether hampers their use in isolated form. The investigations carried
out here with fungal peroxygenases have shown that this enzyme sub-subclass (EC
1.11.2.x) has a promising potential for oxyfunctionalizations and can catalyze a variety of
reactions typical for P450s. Peroxygenases are extracellular, i.e. secreted fungal enzymes
with high stability, which merely need peroxide for function. Results obtained with the
unspecific/aromatic peroxygenases (APOs) of Agrocybe aegerita, Coprinellus radians and
Marasmius rotula have demonstrated that APOs catalyze numerous H 2 O 2 -dependent
monooxygenations of pharmaceuticals and psychoactive drugs. Among them are i) the
monooxygenation of aromatic compounds, ii) the benzylic hydroxylation of toluene
derivatives, iii) the O-dealkylation of different ether structures including the scission of
benzodioxoles (O-demethylenation) and esters as well as iv) the N-dealkylation of
secondary and tertiary amines. The peroxygenases studied considerably differ in their
substrate spectrum and the preferred positions of oxidation. This finding opens the
possibility to develop in the future an “enzymatic toolbox“ on the basis of fungal
peroxygenases for the oxyfunctionalization of pharmaceutically relevant compounds.
Mechanistic studies showed that (1) the monooxygenations always proceed via
incorporation of one oxygen atom from the peroxide, (2) the demethylation of phenacetind1
established a deuterium isotope effect similar to P450s, (3) the catalytic efficiencies for
the studied oxidations are in the same range as those of P450s (though the k cat - and K m -
values are noticeably higher), (4) the kinetic studies with nitro-1,3-benzodioxole gave
parallel double reciprocal plots suggestive of a “ping pong” mechanism, (5) the substrate
spectrum and the activity pattern of APOs follows in a wide range those of the human key
P450s as well as that (6) the difference spectra obtained in bindings studies are of the
phenol type of P450s. Furthermore, APOs were found to be stable and active in long term
experiments over two weeks and they oxidized pharmaceuticals at low, environmentally
relevant concentration (ppb range). All the above properties strongly indicate that APOs
respresent an interesting alternative for the enzymatic conversion of pharmaceuticals as
VIII

SUMMARY
well as for the preparation of human drug metabolites, for example, in medicinal and
pharmacological research or the bioremediation sector (removal of pharmaceuticals from
environmental media).
IX

SUMMARY
Streszczenie
W ostatnich latach, w świecie nauki, coraz więcej uwagi poświęca się lekom i ich
metabolitom. Substancje te cieszą się szczególnym zainteresowaniem z powodu ich
fizjologicznego oraz toksykologicznego działania w ludzkim organiźmie, a także na
środowisko naturalne. Cytochromy P450 (P450s) odgrywają kluczową rolę w konwersji i
detoksyfikacji bioaktywnych substancji, między innymi wielu leków i substancji
psychoaktywnych. W pierwszym rzędzie należą do tej grupy enzymów monooksygenazy,
które są wewnątrzkomórkowe i relatywnie niestabilne. Potrzebują kompleksowych,
drogich kofaktorów i ich oczyszczanie wymaga dużego nakładu pracy, co utrudnia ich
zastosowanie w wyizolowanej formie. Przeprowadzone badania z grzybowymi
peroksygenazami pokazały, że ta podklasa enzymów (EC 1.11.2.x) posiada wysoki
potencjał oxofunkcjonalizacyjny i katalizuje wiele typowych reakcji dla cytochromów
P450. Peroksygenazy są zewnątrzkomórkowe, to znaczy należą do wydzielanych
enzymów grzybowych. Charakteryzują się wysoką stabilnością i wymagają jedynie
obecności nadtlenku wodoru jako kosubstratu. Wyniki otrzymane przy użyciu
aromatycznych/niespecyficznych peroksygenaz (APOs) z Agrocybe aegerita, Coprinellus
radians i Marasmius rotula pokazują, że APOs katalizują wiele rożnych, zależnych od
H 2 O 2 , monooksygenacji leków i substancji psychoaktywnych. Do tego należą i)
monooksygenacja aromatów, ii) hydroksylacja grupy benzylowej pochodnych fenolu, iii)
O-dealkilacja różnych struktur eterycznych włączając rozszczepianie benzodioksoli (Odemetylenowanie)
oraz estrów, jak rownież iv) N-dealkilacja. Badane peroksygenazy
wykazują częściowo znaczne różnice w spektrum substratów i preferowanej pozycji
oksydacji. Powyższe wyniki otwierają możliwość opracowania w przyszłości tak zwanej
„enzymatycznej skrzynki narzędziowej“ na bazie grzybowych peroksygenaz do
oxofunkcjonalizowania farmaceutycznie istotnych substancji czynnych. Mechanistyczne
eksperymenty pokazały, że (1) monooksygenacje nastepują przez wbudowanie atomu tlenu
pochodzącego z nadtlenku wodoru, (2) demetylowanie fenacetyny-d1 wykazuje podobny
efekt izotopowy deuteru jak cytochromy P450, (3) wydajność katalityczna dla badanych
oksydacji znajduje się w podobnym zakresie jak dla P450s, przy czym wartości k cat i K m są
znacznie wyższe, (4) kinetyczne badania w zakresie oksydacji nitro-1,3-benzodioxolu
wykazały równoległe przebiegi prostych w podwójnym wykresie Lineweavera-Burka, co
odpowiada tak zwanemu mechanizmowi “Ping-Pong“, (5) spektrum substratów i rodzaj
aktywności w przypadku APOs pokrywa się w dużym zakresie z najważniejszmi ludzkimi
P450s, oraz że (6) badania w zakresie wiązań wykazały spektra różnicowe w dużym
stopniu przypominające spektra fenolowe cytochromów P450. Ponadto w eksperymentach
X

SUMMARY
długoterminowych (dwa tygodnie) grzybowe peroksygenazy okazały się być stabilne
i aktywne. Były również w stanie oksydować leki, w koncentracjach wystepujących
w środowisku naturalnym (stężenie w zakresie ppb).Wszystkie wymienione wyżej
właściwości wskazują na to, że grzybowe peroksygenazy stanowią interesującą
alternatywę do enzymatycznej konwersji leków, a także do produkcji ludzkich
metabolitów. Mogą być na przykład stosowane w badaniach medyczno-farmakologicznych
oraz w obrębie ochrony środowiska (usuwanie leków ze środowiska).
XI

INTRODUCTION
1 Introduction
1.1 Physiological and environmental aspects of pharmaceuticals
1.1.1 Pharmaceuticals
On 31 March 2004, the European Parliament and the Council of the European Union
defined a medicinal product: "(a) Any substance or combination of substances presented as
having properties for treating or preventing disease in human beings; or (b) Any substance
or combination of substances which may be used in or administered to human beings either
with a view to restoring, correcting or modifying physiological functions by exerting a
pharmacological, immunological or metabolic action, or to making a medical diagnosis
"(Parliment and Council 2004). Pharmaceuticals from a wide spectrum of therapeutic
classes are used worldwide in human and animal medicine. Pharmaceuticals include more
then 4,000 molecules with different physico-chemical and biological properties and
distinct modes of biochemical action (Beausse 2004, Mompelat et al. 2009, Whitacre et al.
2010). Pharmaceuticals can be classified by chemical properties, mode or route of
administration, biological system affected or therapeutic effects. An elaborate and widely
used classification system is the Anatomical Therapeutic Chemical Classification System
(ATC system) (Fricke et al. 2012, WHO Collaborating Centre for Drug Statistics
Methodology 2012). Important classes of pharmaceuticals include:
⎯ Analgesics (painkillers),
⎯ Antibiotics (inhibiting microbial growth),
⎯ Antihypertensives/α-/β-blockers (α-/β-adrenoreceptor antagonists; used to treat
hypertension = high blood pressure),
⎯ Antiepileptic drugs (enhance the effect of the neurotransmitter γ-aminobutyric
acid),
⎯ Nonsteroidal anti-inflammatory drugs, NSAIDs (with analgesic and antipyretic
(fever-reducing) effects; in higher doses with anti-inflammatory effects),
⎯ Antitussives (cough medicines),
⎯ Bronchodilators/antiasthmatic drugs (β2-agonists, anticholinergics, and
theophylline),
⎯ Psychoactive drugs (antidepressants, stimulants, antipsychotics, mood stabilizers,
anxiolytics, depressants, hallucinogens),
⎯ Chemotherapeutic agents (anticancers), and
1

INTRODUCTION
⎯ Hormones (contraceptives, anabolics).
The World Health Organization keeps a list of essential medicines. Essential medicines are
defined as: "those drugs that satisfy the health care needs of the majority of the population;
they should therefore be available at all times in adequate amounts and in appropriate
dosage forms, at a price the community can afford." (World Health Organization 2012).
The core list presents a number of minimum medicine needs for a basic health-care system,
listing the most efficacious, safe and cost-effective medicines for priority conditions.
Priority conditions are selected on the basis of current and estimated future public health
relevance, and potential for safe and cost-effective treatment. The complementary list
presents essential medicines for priority diseases, for which specialized diagnostic or
monitoring facilities, and/or specialist medical care, and/or specialist training are needed.
In case of doubt, medicines may also be listed as complementary on the basis of consistent
higher costs or less attractive cost-effectiveness in a variety of settings (World Health
Organization 2011). As on 17 June 2011, the BfArM (Bundesamt für Arzneimittel und
Medizinprodukte; the Federal Institute for Drugs and Medical Devices) reported marketing
authorizations or registrations for 60,237 pharmaceutical drugs in all indications. The
"Rote Liste®" for 2011, the comprehensive drug registry for Germany, includes only 8,280
preparations, with a total of 33,737 prices (pharmaceuticals are almost always marketed at
different prices for different package sizes). This difference between nearly 60,000
authorized pharmaceuticals on the one hand and only roughly 10,000 drug entries in the
"Rote Liste®" can be explained by the different tallying methods and the only partial
listing of self-medication drugs in the "Rote Liste®" (German Association for
Pharmaceutical Industry 2011).
1.1.2 Pharmacokinetics and Metabolism
Pharmacokinetics, "the movement of drugs" is the study of a drug and/or its metabolite
kinetics in the body. The human body is a very complex system and a drug undergoes
many steps as it is being absorbed (A), distributed through the body (D), metabolized (M),
and/or excreted (E) (figure 1). When the studies are focused solely on one specific
pharmacokinetic aspect (A,D,M or E) by in vitro, in situ, in vivo or in silico techniques
they are usually referred to as ADME studies, whereas the term pharmacokinetics is
normally reserved for in vivo studies where all four aspects are considered (Ruiz-Garcia et
al. 2008). Biotransformation/metabolism is an integral component of the processes that
govern the pharmacokinetic profile of therapeutically used drugs. Biotransformation
generally converts nonpolar, lipophilic pharmacologically active drug molecules into polar,
2

INTRODUCTION
inactive or nontoxic metabolites that are readily eliminated by kidneys or other organs. The
liver is the major site of drug metabolism, with intestinal drug extraction playing a
secondary yet potentially important role in the presystemic elimination of orally
administered agents. Traditionally, biotransformation pathways are classified into two
groups:
⎯ Phase I reactions that involve (oxy)functionalization of drug substrates to yield
more polar derivatives such as alcohols, phenols or carboxylic acids, and
⎯ Phase II reactions involving bimolecular conjugation with endogenous hydrophilic
moieties to yield products such as glucuronides and sulfates that are readily
excreted by kidneys (Venkatakrishnan et al. 2001).
Figure 1 Processes involved in the determination of pharmacokinetic parameters. Adapted from
Gibson and Skett (Gibson and Skett 2008).
Cytochrome P450s (P450s) are the enzymes principally responsible for Phase I oxidations
in animals. Because the chemicals that must be metabolized are structurally very diverse,
animals express multiple P450s that have evolved via gene duplications and conversions,
thus giving rise to genes encoding new P450 forms that metabolize not only xenobiotics,
but also endogenous substances. One consequence of human P450 gene evolution is the
polymorphism of drug metabolism, leading to marked differences in the response of
individuals to the toxic and carcinogenic effects of drugs and other environmental
3

INTRODUCTION
chemicals. Differences in P450 expression between human individuals, which are
correlated with ethnicity in some cases, may reflect the presence or absence of a particular
P450 gene, but may also involve transcriptional regulation. These differences may be
clinically manifested in quantitative differences in the metabolism of certain drugs,
resulting in "poor" or "extensive" metabolizers (Souček and Gut 1992). P450-mediated
drug metabolism can also lead to altered drug efficacies through inactivation of an active
drug or activation of a prodrug (Ding and Kaminsky 2003). Three P450 gene families
(CYP1, CYP2, and CYP3) and to some degree a fourth one (CYP4) are currently thought
to be responsible for the majority of hepatic drug metabolism in humans (Wrighton and
Stevens 1992).
Figure 2 A summary of major human P450s involved in drug metabolism, including major
substrates. The circle sizes approximate relative levels of expression in human liver. The overlap of
the circles conveys the point that an overlap of catalytic action is often observed (Gibson and Skett
2008, Guengerich and Montellano 2005).
All three members of the CYP1 family (CYP1A2, CYP1B1, CYP1A1) are upregulated by
halogenated and polycyclic aromatic hydrocarbons, such as those found in cigarette smoke
and charred food (e.g. benzo(a)pyrene). CYP1A2 is responsible for melatonin oxidation
and uroporphyrin formation from uroporphyrinogen: it metabolizes about 24 drugs and all
three members are metabolically active and detoxify many environmental carcinogens
(Nelson and Nebert 2011). CYP2 is the largest P450 family in mammals with 13
subfamilies and 16 genes in humans. CYP2C8, -9, -18, and -19 together metabolize more
than 50 drugs, as well as arachidonic acid and steroids. CYP2D6 metabolizes more than 70
drugs. CYP2A6, -A13, -B6, -E1 and -F1 are also major contributors to drug metabolism
(Guengerich et al. 1998, Meyer and Zanger 1997, Nelson and Nebert 2011). P450
subfamily 3A is the most abundantly expressed P450 gene in the human liver and
gastrointestinal tract and known to metabolize more than 120 commonly used
4

INTRODUCTION
pharmaceutical agents (Nelson and Nebert 2011). The human fetus is capable of
metabolizing a large number of compounds, and its major P450 is a member of the 3A
subfamily, specifically 3A7. This single P450 comprises 30 to 50% of the total fetal P450s.
However, 3A7 has not been detected in the liver of adults (Wrighton and Stevens 1992).
The CYP4 family metabolizes some drugs, but is involved mainly in the metabolism of
fatty acids (Nelson and Nebert 2011).
1.1.3 Pharmaceutical drugs development and studies
In the course of modern drug development, there are four areas in which issues pertaining
to drug metabolites need to be addressed, namely (i) drug metabolites as components of
(plasma) drug assays, (ii) drug metabolites in bioavailability studies, (iii) drug metabolites
in bioequivalence studies, and (iv) drug metabolites in safety testing (Baillie et al. 2002).
Pharmaceutical companies and regulatory agencies are becoming increasingly interested in
the role of metabolites as potential mediators of the toxicity of new drug products (Baillie
et al. 2002). In 2005, the U.S. Food and Drug Administration issued a draft guidance
document "Safety testing of drug metabolites" which was followed in 2008 by the final
version (Baillie 2009, Center for Drug Evaluation and Research (HFD-130) and Food and
Drug Administration 2008). Drugs entering the body undergo biotransformation via Phase
I and Phase II metabolism. Based on the nature of the chemical reactions involved,
metabolites formed from Phase I reactions can be chemically more reactive or even
represent the actually bioactive form of the drug. An active metabolite may bind to the
therapeutic target receptors or other receptors, interact with other targets (e.g. enzymes,
proteins), and cause unintended effects (Center for Drug Evaluation and Research (HFD-
130) and Food and Drug Administration 2008). It is recommended that - while there is a
necessity to maintain a flexible, case-by-case evaluation of the need to perform additional
safety studies on human drug metabolites - those metabolites present at disproportionately
higher levels in human plasma than in any of the animal test species should be considered
for safety assessment (Baillie 2009).
By the early 1990s, much of the interest in P450 research had shifted to the human P450
enzymes because of the availability of systems for handling them. In particular, studies in
the areas of drug metabolism and chemical toxicology/ carcinogenesis were facilitated by
the knowledge that a relatively small number of the P450s account for a large fraction of
the metabolism of the drugs. In the pharmaceutical industry, the roles of the major hepatic
P450s are extensively studied to develop predictions about bioavailability, drug-drug
interactions, and toxicity (Guengerich and Montellano 2005). The metabolism of foreign
5

INTRODUCTION
compounds by P450s generally leads to products that are less toxic and more easily
excreted. However, in some cases oxidative metabolism produces substances that are
reactive toxins and mutagens (Williams et al. 2000). Toxic compounds may be detoxified
following P450-catalyzed biotransformation while inert xenobiotics, including drugs, may
be activated and thus become toxicants. Most xenobiotic compounds require metabolic
activation by P450s to form ultimate carcinogens or toxicants. Most hepatic P450s are also
present in extrahepatic tissues, and often at higher levels. Furthermore, some P450s are
expressed preferentially in extrahepatic tissues, which may lead to unique extrahepatic
metabolites and tissue-specific consequences in cellular toxicology and organ pathology.
Risk assessment of potential human toxicants is currently based primarily on data obtained
from animal bioassays and the knowledge of mechanism of toxicity derived from animal
experiments (Ding and Kaminsky 2003). Metabolic studies using biological models are
important for determining the significance of specific enzymes or pathways for toxicity
(Ding and Kaminsky 2003).
Pharmaceutical products (PPs) and by-products (BPs) are spread into 24 therapeutic
classes, among which four are dominant in terms of references. About 40% of studies
concern non-steroidal anti-inflammatory drugs (NSAIDs), the three others being
anticonvulsants, antibiotics, and lipid regulators. NSAIDs are essentially represented by
ibuprofen and diclofenac. The most studied lipid regulator is gemfibrozil. Carbamazepine
is the most tested anticonvulsant (Mompelat et al. 2009).
1.1.4 Environmental aspects
In the European Union today, about 3,000 different substances are being used in medicines
such as painkillers, antibiotics, contraceptives, β-blockers, lipid regulators, tranquilizers,
and impotence drugs (Ternes et al. 2004, Wiegel et al. 2004). In Germany, in the year
2000, about 7,000 t of synthetic medicines for human consumption were sold. (report from
the Federal Environmental Authority; Umweltbundesamt). The figure for veterinary drugs
was 2,320 t in 2002 (Wiegel et al. 2004). Table 1 shows some examples of pharmaceutical
drugs chosen as environmentally relevant together with their annual consumption and
quantities remaining as unconverted substances. Human-use medicines are designed to
have a biological effect and to be bioavailable. However, it is only in recent years that
there has been increasing concern over the trace amounts of pharmaceuticals that are
appearing in the environment and the effects they may cause (Daughton and Ternes 1999,
Whitacre et al. 2010).
6

INTRODUCTION
Table 1 Consumption of pharmaceutical drugs in Germany in 2001 and percentages remaining
unconverted (Sachverständigen Rat für Umweltfragen 2007).
Pharmaceutical class/drug
Consumption per year Unconverted substrate
Analgesics
(kg)
(% of dose)
Metamizole 64,400 0
Phenazone 24,850 5
Propyphenazone 24,180 2
Codeine 9,700 70
Morphine 880 10
NSAIDs (see 1.1)
Ibuprofen 344,880 1
Diclofenac 85,800 70
Indometacin 3,700 30
Ketoprofen 1,613 10
Piroxicam 724 5
Meclofenamic acid not determined not determined
Antitussives/cough medicines
Ambroxol 14,470 6
Codeine 9,700 70
Dihydrocodeine 1,245 40
Hydrocodone 8 40
Bronchodilators/antiasthmatic
drugs
Salbutamol 414 80
Terbutaline 118 60
Fenoterol 72 50
Clenbuterol 1 90
Antibiotics
Sulfamethoxazole 53,600 33
Doxycycline 24,180 2
Ciprofloxacin 17,973 45
Clindamycin 16,100 30
Trimethoprim 11,426 80
7

INTRODUCTION
Roxithromycin 9,550 60
Clarithromycin 7,159 35
Norfloxacin 4,724 70
Ofloxacin 2,280 95
Oxytetracycline 2,020 not determined
Tetracycline 1,530 not determined
Spiramycin 300 25
Chloramphenicol 202 10
Chlortetracycline 99 not determined
Antihypertensives/α/β-blockers
Metoprolol 93,000 10
Sotalol 26,600 90
Atenolol 13,500 90
Propranolol 3,900 5
Bisoprolol 2,914 50
Antiepileptics
Carbamazepine 87,600 30
Psychoactive drugs
Diazepam 1,107 30
Chemotherapy agents
Cyclophosphamide 385 7
Ifosfamide 170 50
Hormones/contraceptives
17α-Ethinylestradiol 50 85
Data origin: SRU/Stellungnahme Nr.12-2007; Huschek and Krengel 2003
The occurrence of anthropogenic organic contaminants in aquatic ecosystems has drawn
the attention of both the public and scientists in recent years, a fact confirmed by the
numerous studies published in this field. With respect to pharmaceuticals, many
investigations have focused on high consumption substances currently being introduced
into the environment (Hass et al. 2012). Accurate assessment of drugs impact on the
environment is difficult, since there is a multitude of input sources in the environment with
no evident quantitative data available concerning the relative distribution of
pharmaceuticals from all emission sources (shown in figure 3).
8

INTRODUCTION
Figure 3 Origin and routes of pharmaceutical products (adapted from Petrović et al. (Petrović et al.
2003) and from Mompelat et al. (Mompelat et al. 2009). DWT: drinking water treatments; WWTP:
waste water treatment plant.
However, humans and animals account for the main contamination source of potable water
resources (Mompelat et al. 2009). Once ingested and metabolized, pharmaceuticals (native
and/or metabolites) are excreted via urine and feces, or unused or expired drugs are
improperly disposed of in toilets, and travel in urban wastewater to the wastewater
treatment plant (WWTP) or, for countryside households, they are directly released into
septic tanks (Carrara et al. 2008, Mompelat et al. 2009). To a minor but relevant extent,
hospital wastewater also contributes to the total loads. Pharmaceuticals can also enter
agricultural land when digested sludge is applied as fertilizer. Veterinary pharmaceuticals
are excreted directly into the environment, and thus contaminate soil while the resulting
runoff gets into rivers. Therefore, the efficiency with which WWTPs can remove
pharmaceuticals is crucial (Ternes et al. 2004).
In recent decades, more than 100 different drugs have been detected in aquatic
environments at concentrations from ng L -1 to µg L -1 (Daughton and Ternes 1999). Recent
studies have demonstrated that, despite relatively low concentrations of pharmaceuticals in
the environment (typically at sub-parts-per-billion level; < ppb), pharmaceuticals are of
ecological concern due to their potential long-term adverse effects on humans and wildlife
(Ankley et al. 2007, Celiz et al. 2009). In most of the investigations reported to date, the
efficiency of drugs removal during water treatment is determined by measuring the
disappearance of the parent compound, but not the formation of by-products. Similarly,
9

INTRODUCTION
there is a lack of available information on the environmental fate and occurrence of
excreted human and animal drug metabolites. The European Medicines Agency has set
guidelines for reporting concentrations of drugs and their metabolites. A systematic
approach has been developed to predict the total risk associated with pharmaceuticals
entering the environment. The guidelines say that any metabolite formed at a concentration
greater than 10 percent of the parent compound needs further investigation (Boxall et al.
2004, Celiz et al. 2009).
Ecotoxicity studies in the laboratory have demonstrated effects of pharmaceuticals on end
points such as reproduction, growth, behavior and feeding for fish and invertebrates
(Martinović et al. 2007, Quinn et al. 2008, Whitacre et al. 2010). Drugs have already been
detected in fish tissues (Brooks et al. 2005). In the terrestrial environment, the catastrophic
decline of Indian vulture populations was found to be the result of exposure to the human
anti-inflammatory drug diclofenac (Oaks et al. 2004, Whitacre et al. 2010). In the river
Elbe, in 1998, diclofenac, ibuprofen and carbamazepine were found as well as various
antibiotics and lipid regulators in a concentration range from

INTRODUCTION
terminology P450 is uncommon for enzymes because it is not based on function, but
originally describes the spectral properties of this b-type heme containing red pigments,
which display a typical absorption band around 450 nm in their reduced carbon-monoxide
bound form (Omura and Sato 1964). All P450s contain a cysteine-ligated heme (iron
protoporphyrine IX) that can bind dioxygen (O 2 ) in the active site (Dawson and Sono
1987).
The official nomenclature in enzymology, which has been in use since the early 1960s,
follows the EC 1 classification system that is strictly based on the reaction catalyzed.
Enzymes catalyzing a particular reaction get so-called EC-numbers that comprise four
digits (Nomenclature Committee of the International Union of Biochemistry and Molecular
Biology (NC-IUBMB) 2012).
Most P450s belong to the main class of oxidoreductases (EC 1.x), more precisely to those
oxidoreductases acting on paired donors, with incorporation or reduction of molecular
oxygen (EC 1.14.x). There are some exceptions, where P450s are found, for example, in
the main class of isomerases and lyases (intramolecular oxidoreductases, EC 5.3.x and
carbon-oxygen lyases, EC 4.2.x) (Degtyarenko 2004). Members of the P450 enzyme
family are involved in the biotransformation of drugs, the bioconversion of xenobiotics, the
metabolism of chemical carcinogens, the biosynthesis of physiologically important
compounds (such as steroids, fatty acids, eicosanoids, fat-soluble vitamins, and bile acids),
the conversion of alkanes, terpenes, and aromatic compounds as well as in the degradation
of herbicides and insecticides (Bernhardt 2006). The reactions catalyzed can be extremely
diverse and include e.g. hydroxylations, N-, O- and S-dealkylations, sulphoxidations,
epoxidations, deaminations, desulphurations, dehalogenations, peroxidations, N-oxide
reductions, dehydrations and isomerizations(Bernhardt 2006, Guengerich 2001, Mansuy
1998). Many P450s convert a broad range of substrates and catalyze multiple reactions.
For example, unspecific monooxygenases (EC 1.14.14.1) act on xenobiotics, steroids, fatty
acids, vitamins and prostaglandins and they catalyze amongst others hydroxylations,
epoxidations, N-oxidations, sulfooxidations, N-, S- and O-dealkylations, desulfations,
deaminations, and the reduction of azo-, nitro- and N-oxide groups.
Since many of the individual P450s catalyze multiple reactions, the usual method of
naming enzymes is inadequate for this group of proteins, so a systematic nomenclature has
been developed on the basis of structural and sequence homology (Nelson et al., Nelson et
1 EC - Enzyme Commission: Nomenclature Committee of the International Union of Biochemistry and
Molecular Biology (NC-IUBMB), in consultation with the IUPAC-IUBMB Joint Commission on
Biochemical Nomenclature (JCBN), http://www.chem.qmul.ac.uk/iubmb/enzyme/
11

INTRODUCTION
al. 1996). P450 genes are identified by the abbreviation CYP followed by a number
denoting the family; members of a family show more than 40% sequence identity.
Following is a letter designating a subfamily whose members share more than 55%
sequence identity. Finally, the second number indicates the individual gene within the
subfamily (e.g. CYP2A1) (Hannemann et al. 2007). At present, there is a total of over
1,000 P450 families and over 2,500 subfamilies (Nelson 2011). Altogether, more than
12,000 P450 genes have been reported across all kingdoms of life. Mammals have usually
between 50 and 100 encoded P450 enzymes. There are even more CYP genes found in
plant genomes than in animal genomes, because plants are sessile and carry out their
biochemistry de novo from carbon dioxide, ammonia, water and minerals.
Table 2 Named cytochrome P450s in March 2010, adapted from Nelson et al., 2011 (Nelson 2011)
Kingdom/ organisms P450 number CYP families a Sequenced
genomes b
Animals (Metazoa) 4088
CYPs/genome
Insects 2137 67 11 37-178 c
Vertebrates 1461 18 8 57-102 d
Invertebrates (without
490 71 4 76-89 e
insects)
Plants (Plantae) 4267 126 14 10-332 f
Fungi (Eumycota) 2784 399 53 1-153 g
Protists 247 62 22 1-55 h
Eubacteria 1042 333 212 1-52 i
Archaea 26 13 17 1-4
Viruses (Mimivirus) 2 2 1 2
Total 12,456
a includes all named CYP families; not
just those from sequenced genomes.
b
only genomes included in the
nomenclature in 2010
c
Aedes aegypti
d
Mouse (Mus
musculus)
e
Tetranychus
urticae
f Rice (Oryza
sativa)
g
Aspergillus
oryzae
h Dictyostelium
discoideum
i
Frankia sp.
Angiosperm plants may have between 250 (Arabidopsis spp.) and over 400 P450s (potato,
Solanum tuberosum) that are involved in stress responses, reaction to wounding, seedling
development, taste, colour and aroma of fruits and flowers, structural support (lignin),
water barriers (cutins and suberins), hormone signaling and synthesis of allelochemicals
12

INTRODUCTION
for defence against predators (Nelson and Nebert 2011). Fungi can also possess numerous
P450s; thus in the genome of the basidiomycetous white-rot fungus Phanerochaete
chrysosporium, more than 150 P450 genes (most of them monooxygenases) were found
(Subramanian et al. 2010), and 153 genes are present in the ascomycetous mold
Aspergillus oryzae (Nelson 2011). Table 2 gives an overview of the number and
distribution of P450s in different groups of organisms.
1.2.1 Classification of electron transport systems of P450s
After the discovery of cytochrome P450s, two main classes reflecting the redox partners
involved were described: the adrenal mitochondrial P450 systems obtaining electrons from
NAD(P)H via adrenodoxin reductase and adrenodoxin (Hannemann et al. 2007, Omura et
al. 1966), and the liver microsomal P450s obtaining electrons from NAD(P)H via a flavincontaining
(FAD, FMN) P450 reductase (Hannemann et al. 2007, Lu and Coon 1968). It
was not until the 1980s, when CYP102 (P450BM3) was discovered, that an example came
to light showing an unusual property, the fusion of the P450 and reductase components
into one polypeptide chain (Narhi and Fulco 1987). Since then, several more of such
“unusual” electron transfer chains have been described for different P450s (table 3). Class
II cytochrome P450s are the most common type found in all eukaryotic organisms (Črešnar
and Petrič).
Table 3: Classes of P450 systems grouped according to the topology of protein components
involved in the electron transfer towards the actual P450 component, including schematic
organization of the proteins (Hannemann et al. 2007). Protein components contain the following
redox centers: Fdx (iron-sulfur-cluster); FdR, Ferredoxin reductase (FAD); CPR, cytochrome P450
reductase (FAD, FMN); Fldx, Flavodoxin (FMN); OFOR, 2-oxoacid: ferredoxin oxidoreductase
(thiamin pyrophosphate, [4Fe-4S] cluster); PFOR, phthalate-family oxygenase reductase (FMN,
[2Fe-2S] cluster).
Class/ source
Class I
Bacterial
Mitochondrial
Electron transport
chain
NAD(P)H►[FdR] ►[Fdx]
►[P450]
NADPH►[FdR] ►[Fdx]
►[P450]
Scheme
Localization/
remarks
Cytosolic, soluble
P450s; inner
mitochondrial
membrane; FdRmembrane
associated; Fdxmitochondrial
matrix, soluble
13

INTRODUCTION
Class II
Bacterial,
fungal,
Microsomal A
Microsomal B,
fungal
Microsomal C
NADPH►[CPR] ►[P450]
NADPH►[CPR] ►[P450]
NADPH►[CPR] ►[cytb5]
► [P450]
NADPH►[cytb5Red] ►
[cytb5] ► [P450]
Cytosolic, soluble
Membrane
anchored,
endoplasmatic
reticulum
(A,B,C)
Class III
Bacterial
NAD(P)H►[FdR] ►[Fldx]
►[P450]
Cytosolic,
soluble,
Citrobacter
braakii
Class IV
Bacterial
Pyruvate, CoA►[OFOR]
►[Fdx] ►[P450]
Cytosolic,
soluble,
Sulfolobus
tokadaii
Class V
Bacterial
NADPH►[FdR] ►[Fdx-
P450]
Cytosolic,
soluble,
Methylococcus
capsulatus
Class VI
Bacterial
NAD(P)H►[FdR] ►[Fldx-
P450]
Cytosolic soluble,
Rhodococcus
rhodochrous
strain 11Y
Class VII
Bacterial
NADPH►[PFOR-P450]
Cytosolic,
soluble,
Rhodococcus sp
strain NCIMB
9784,
Burkholderia sp.,
Ralstonia
metallidurans
14

INTRODUCTION
Class VIII
Bacterial,
fungal
Class IX
Only NADH
dependent,
fungal
Class X
Independent in
plants/
mammals
NADPH►[CPR-P450]
NADH►[P450]
[P450]
Cytosolic,
soluble, Bacillus
megaterium,
Fusarium
oxysporum
Cytosolic,
soluble, Fusarium
oxysporum
Membrane bound,
ER, example:
Thromboxane
synthase
1.2.2 Bacterial P450s
Bacterial P450s form a large proportion of the superfamily. They are soluble enzymes
whereas their eukaryotic counterparts are usually membrane-bound. This fact simplifies
the overexpression and purification of bacterial enzymes (Munro and Lindsay 1996). Most
eubacterial species contain at least one P450 enzyme while some archaea seemingly do not
possess any P450 enzyme. It is possible that P450s may have evolved during the advent of
atmospheric oxygen in the atmosphere (about 1.5 to 1 billion years ago), where their
original role was in the detoxification of molecular oxygen (O 2 ) in anaerobic bacteria
(Lewis and Wiseman 2005, Wickramasinghe and Villee 1975).
A bacterial P450 was first found in nitrogen-fixing Rhizobium bacteroides (Appleby 1967).
In contrast to the microsomal and mitochondrial P450s reported before, the Rhizobium
P450 was soluble (i.e. dissolved in the cytosol); however, the function was not elucidated.
Then P450s in Pseudomonas putida and Coryebacterium sp. were reported (Cardini and
Jurtshuk 1968, Katagiri et al. 1968) . P450 cam (CYP101) from Pseudomonas was found to
catalyze the NADH-dependent oxidation of camphor and required soluble NADH,
putidaredoxin and NADH-putidaredoxin reductase for the reaction. Corynebacterium P450
catalyzes the NADH-dependent terminal oxidation of n-octane to 1-octanol. More soluble
P450s were found in various prokaryotic organisms in the following years (Omura 2005).
The first self-sufficient P450-reductase fusion enzyme was found in Bacillus megaterium
and named P450 BM-3 (CYP102A2). Similar P450-reductase fusion proteins were later
found in other bacterial species (Narhi and Fulco 1987, Omura 2005). P450 eryF from
15

INTRODUCTION
Saccharopolyspora erythrea is the only bacterial P450 to which a biosynthetic function has
been assigned. The bacterium produces erythromycin A, and P450 eryF catalyzes the
hydroxylation of 6-deoxyerythronolide B to generate erythronolide B, which is a precursor
of this macrolide antibiotic (Munro and Lindsay 1996). Among the P450s characterized
from bacteria of the order Actinomycetales (Rhodococcus spp. strains) are enzymes
catalyzing alkane hydroxylation, pentachlorophenol dehalogenation and alkoxyphenol O-
dealkylation (Karlson et al. 1993, Munro and Lindsay 1996). The most known function of
bacterial P450s is the oxidation of recalcitrant organic compounds so that the hosts can
utilize them as sole carbon source for growth. This is also the generally recognized
difference between prokaryotic and eukaryotic P450s (involvement in carbon metabolism
vs. detoxification/biosynthesis) (Fulco 1991). However, the role of bacterial P450s in the
utilization of unusual organic compounds is non-essential for the viability of the
organisms. The ability of bacterial P450s to metabolize compounds such as terpenes and
alkanes has attracted interest, not only regarding the decomposition of organic wastes and
pollutants, but also for biotransformations in the synthesis of pharmaceuticals and fine
chemicals (Munro and Lindsay 1996). Aside from the inherent interest in the physiological
roles and biotechnological potential of bacterial P450s, these enzymes have become
attractive and productive targets for spectroscopic and structural analyses (Munro and
Lindsay 1996).
1.2.3 Plant P450s
Plants utilize a diverse array of P450s in their biosynthetic and detoxification pathways.
Biosynthetic P450s play paramount roles in the synthesis of lignin intermediates, sterols,
terpenes, flavonoids/isoflavonoids, furanocoumarins, cyanogenic glycosides, lipids, plant
growth regulators (phytohormones) such as gibberellins, jasminoic acid, and
brassinosteroids, and a variety of other secondary plant metabolites (Chapple 1998,
Nielsen et al. 2005, Schuler 1996a).
Many scientific studies have shown the important role of P450s in the biosynthesis and
degradation of compounds important in plant insect interactions (Schuler 1996b), and the
importance of plant P450s in the metabolism of herbicides (Frear 1995). Plant P450
monooxygenases are membrane-bound heme proteins which consist of a protoporphyrin
IX and an apoprotein that confers the substrate specificity. Plant P450s have been shown to
carry out hydroxylations, epoxidations, heteroatom dealkylations and oxygenations
(Bolwell et al. 1994). Cinnamic acid 4-hydroxylase from Helianthus tuberosus (Jerusalem
artichoke) was the first plant P450 to be functionally characterized. This enzyme catalyzes
16

INTRODUCTION
an essential step in the phenylpropanoid pathway and is considered to be ubiquitous in
plants (Nielsen et al. 2005). Like vertebrates and fungi, certain plants produce
polyhydroxylated steroidal hormones, called brassinosteroids, to regulate and control tissue
morphology. They are nonessential phytohormones with impacts on morphological
characteristics, for example, leaf shape and dwarfism (Clouse and Sasse 1998, Nielsen et
al. 2005).
1.2.4 Fungal P450s
Cytochrome P450 monooxygenases of fungi are involved in many essential cellular
processes and play diverse roles. They catalyze the conversion of hydrophobic
intermediates of primary and secondary metabolic pathways, detoxify natural toxins and
environmental pollutants and allow fungi to grow under different conditions. Fungal P450s
represent three electron transport systems, Class II, Class VIII and Class IX (table 1;
1.2.1). Class II P450s are involved in metabolite synthesis as well as in detoxification
and/or degradation of xenobiotic compounds. Class VIII is represented by P450 foxy (from
Fusarium oxysporum). This enzyme is responsible for the hydroxylation of fatty acids.
P450s belonging to Class IX of electron transfer systems are involved in the process of
fungal denitrification (Črešnar and Petrič, Shoun et al.). The key enzyme, nitric oxide
reductase (EC 1.7.1.14), is also designated as P450 nor and shows structural similarity to
"classic" P450 monooxygenases. But on the other hand, P450 nor functionally differs from
the rest of P450s as it catalyzes the reduction of two molecules of nitric oxide (NO) to
nitrous oxide (N 2 O) whereby the electrons are directly transferred from NAD(P)H to the
enzyme without the need of any additional redox partner. P450 nor is the only eukaryotic
P450 that is soluble, i.e. not membrane-associated (Črešnar and Petrič, Shoun et al.).
Another fungal P450 family (CYP53) uses e.g. benzoic acid as substrate to produce 4-
hydroxybenzoic acid or catalyzes the demethylation of 3-methoxybenzoic to 3-
hydroxybenzoic acid, and has thus an essential role in the detoxification of aromatic
compounds (Lah et al.). One well-studied class of bioconversions carried out by fungal
P450 enzymes is the (stereo)-specific hydroxylation of pharmaceutically interesting
steroids, especially progesterone (Mahato and Majumdar 1993, van den Brink et al. 1998).
Another P450-catalyzed process is the 14α-demethylation of lanosterol in yeasts and of
eburicol in filamentous fungi. Ergosterol (the product of this reaction), has a similar
structure as mammalian cholesterol and performs an important role in membrane
stability/fluidity and in completion of the cell cycle (van den Brink et al. 1998).
17

INTRODUCTION
The evolutionarily conserved (CYP51) and fungi-specific (CYP61 and CYP 56) P450s are
essential for primary metabolism, enabling these microorganisms to preserve cell wall
integrity and to form the spore outer wall, respectively, regardless of the fungus’
morphology or ecological niche. Furthermore, several other fungal P450s with other
activities and functions have been characterized. They were found to catalyze e.g. the
detoxification of xenobiotic compounds, the initial step in n-alkane assimilation, fatty acid
hydroxylation, nitrite reduction and mycotoxin production (Črešnar and Petrič). A wellstudied
P450-catalyzed reaction in fungi is the alkane assimilation/degradation by some
Candida species (Scheller et al. 1996, van den Brink et al. 1998). Table 4 below lists
selected fungal P450s and their functions (those known so far).
Table 4 Compilation of most known fungal P450s and their functions.
Fungus
(fungal group)
Aspergillus
parasiticus
(ascomycete)
Candida albicans
(ascomycetous
yeast)
Candida maltosa
(ascomycetous
yeast)
Cunninghamella
bainieri
(zygomycete)
Curvularia lunata
(ascomycete)
Fusarium
oxysporum
(ascomycete)
Fusarium
sporotrichioides
(ascomycete)
Giberella fujikuroi
(ascomycete)
Nectria
haematococca
(ascomycete)
Penicillium paxilli
(ascomycete)
Phanerochaete
chrysosporium
(basidiomycete)
P450 /known
information
Catalyzed
reaction
Known
substrate(s)
Ref.
CYP64 family oxygenation O-methylsterigmatocystin,
(Yu et al. 1998)
dihydrosterigmatocystin
P450 14αdm (CYP51 O-demethylation steroids (Joseph-Horne
family)
and Hollomon
CYP52 family
P450
oxygenation/
hydroxylation
hydroxylation
N-demethylation
n-alkane
hydrocarbons
codeine
1997)
(Ohkuma et al.
1998)
(Ferris et al.
1976, Gibson et
al. 1984)
P450 lun 11β-hydroxylation steroids (Suzuki et al.
1993)
P450 foxy (CYP505) hydroxylation fatty acids (Nakayama et
al. 1996)
CYP65A1 hydroxylation mycotoxin
biosynthesis
CYP68 family hydroxylation kaurenoic acid
(giberellin
biosynthesis)
(Alexander et al.
1998)
(Malonek et al.
2005, Tudzynski
et al. 2001)
CYP57 family O-demethylation pisatin (Desjardins and
VanEtten 1986,
Maloney and
VanEtten 1994)
P450 paxP,
P450 paxQ
oxygenation paspalin (paxilline
biosynthesis)
149 P450s estimated,
12 families,
23 subfamilies.
Functions like plant
CYP73 and CYP98
Functions like bacterial
CYP102 (P450 BM3 ).
Seven sequences in the
hydroxylation
cinnamic acid
(Saikia et al.
2007)
(Doddapaneni et
al. 2005,
Matsuzaki and
Wariishi 2004,
Syed and Yadav
2012)
hydroxylation cumene (Matsuzaki and
Wariishi 2004)
18

INTRODUCTION
Pleurotus ostreatus
(basidiomycete)
Pleurotus
pulmonarius
(basidiomycete)
Rhisopus nigricans
(zygomycete)
Saccharomyces
cerevisiae
(ascomycetous
yeast). Only three
P450 sequences:
same phylogenic cluster
as CYP102
Functions like yeast
CYP52, seven sequences
were categorized into the
CYP52 family
Functions like bacterial
CYP105A1 and
CYP105B1, mammalian
CYP2B, plant CYP76B1
and fungal
CYP510A1,but similarity
above 35% only with
CYP510A1
Functions like
mammalian CYP2A, but
no sequence similarity
above 35%
Functions like fungal
CYP53, one sequence of
P450 from
P.chrysosporium fell into
the CYP53 familiy
Functions like bacterial
CYP101(P450 cam ), no
similarity above 35%
Functions like bacterial
CYP176A1(P450 cin ) and
mammalian CYP3A, but
no sequential similarity
above 35%
PcCYP65a2, and
PcCYP11a3 clones,
functions like human and
hydroxylation 1,12-dodecanediol (Matsuzaki and
Wariishi 2004)
deethylation
Hydroxylation
7-position, by
P.chrysosporium
Four different
positions
hydroxylation
4-position, by
P.chrysosporium
three different
positions
Hydroxylation
5- and 6- position,
by
P.chrysosporium
three different
positions
4-ethoxybenzoic
acid and 7-
ethoxycoumarin
coumarin
benzoic acid
camphor
(Matsuzaki and
Wariishi 2004)
(Matsuzaki and
Wariishi 2004)
(Matsuzaki and
Wariishi 2004)
(Matsuzaki and
Wariishi 2004)
hydroxylation 1,8-cineol (Matsuzaki and
Wariishi 2004)
hydroxylation
mono- and
dichloro-p-dioxins
(Kasai et al.)
rat CYP1A2
CYP5136A3 hydroxylation alkylphenols, PAH (Syed et al.
2011)
P450 dNIR , P450 nor denitrification nitrite (Shoun et al.,
Shoun and
Tanimoto 1991)
CYP53 (cloned
PcCYP1f)
P450 hydroxylation pyrene,
anthracene,
fluorene,
hydroxylation benzoate (Matsuzaki and
Wariishi 2005)
(Bezalel et al.
1996)
dibenzothiophene
P450 hydroxylation benzo[a]pyrene (Masaphy et al.
1995, Maspahy
et al. 1999)
P450 11α hydroxylation steroids (Bossche and
Koymans 1998,
Breskvar et al.
1995)
CYP61 desaturation steroids (Kelly et al.
2005, Nelson
2009)
19

INTRODUCTION
CYP51F1,CYP56A1
and CYP61A1
Trametes (Coriolus)
versicolor
(basidiomycete)
Trichosporon
moniliiforme
and T.cutaneum
(basidiomycetous
yeast)
Yarrowia lipolytica
(ascomycetous
yeast)
Cloned gene, similarity
with CYP512A1, but less
then 40% identity with
any P450
sulfur-oxidizing
reactions
DBT derivatives
(Ichinose et al.
2002)
P450 decarboxylation salicylate, phenol (Stundl et al.
2000)
CYP52 family
oxygenation/
hydroxylation
n-alkane (Iida et al. 2000)
1.2.5 Animal P450s
1.2.5.1 Insect P450s
In insects, the number of cytochrome P450s in sequenced genomes ranges from 37 in the
body louse, Pediculus humanus (Lee et al. 2010, Sztal et al. 2012), up to 160 (178,
corrected in 2011 (Nelson 2011)) in the dengue mosquito, Aedes aegypti (Strode et al.
2008, Sztal et al. 2012). There are four large clades of insect P450 genes that existed
before the divergence of the class of Insecta and that are also present in vertebrates: the
CYP2 clade, the CYP3 clade, the CYP4 clade and the mitochondrial P450 clade.
Mitochondrial P450s are not found in plants or fungi, but seem to be restricted to animals
(Feyereisen 2006). P450s from each of these clades have been linked to insecticide
resistance or to the metabolism of natural products and xenobiotics. In particular, insects
appear to maintain a repertoire of mitochondrial P450 paralogues devoted to the response
to environmental stress (Feyereisen 2006). The metabolism of xenobiotics, insecticides in
particular, to less toxic metabolites is probably the best-known feature of P450s
(Feyereisen 1999). Cytochrome P450 monooxygenases are involved in many cases of
resistance of insects to insecticides. Resistance has been proposed to be associated with
increase in monooxygenase activities and the cytochrome P450 content (Bergé et al. 1998).
Insect P450s can metabolize a wide range of plant allelochemicals, including
furanocoumarins, terpenoids, indoles, glucosinolates, flavonoids, cardenolides,
phenylpropenes, ketohydrocarbons, alkaloids, lignans, pyrethrins, and the isoflavonoid
rotenone (Li et al. 2007). However, there is an increasing number of studies indicating that
P450s have a much broader role in insects than just detoxification. For example,
biosynthesis of juvenile and molting hormones are processes in which the role of insect
P450s has been well documented (Scott 2008).
20

INTRODUCTION
1.2.5.2 Mammalian/ Human P450s
There are eight P450 families in mammalians, which are involved in the metabolism of
pharmaceuticals and drugs, products of incomplete combustion (e.g. polycyclic aromatic
hydrocarbons, PAHs) as well as of plant ingredients (e.g. phenolics, phytoalexins). It has
been proposed that four of these P450 gene families (I, II, III, and IV) evolved and
diverged in animals due to their exposure to plant ingredients and decayed plant products
during the last one billion years. The overlapping substrate specificities of these P450
enzymes are remarkable (Nebert and Gonzalez 1987). There are at least 43 gene families
recognized in animals. In humans, 57 P450s were identified and this number seems to be
complete because of our excellent knowledge of the human genome (Guengerich and
Montellano 2005, Nelson 2009).
Most eukaryotic P450s are intrinsic membrane proteins that are expressed in the
endoplasmic reticula of plant, fungal, and animal cells. Animal cells also express P450s
that are located in the inner membrane of mitochondria. The human enzymes contribute to
the synthesis of steroid hormones, prostacyclin, thromboxane, cholesterol, and bile acids,
as well as to the degradation of endogenous compounds, such as fatty acids, retinoic acid,
and steroids, and exogenous compounds that include drugs and carcinogens (Williams et
al. 2000). In humans, of the P450s with known catalytic activities, 14 are involved in
steroidogenesis, 4 are involved in certains aspects of vitamin metabolism, 5 are involved in
the metabolism of eicosanoids, 4 have fatty acids as substrates, and 15 catalyze the
transformation of xenobiotics. Many of the xenobiotic-converting P450s can also oxidize
steroids and fatty acids. Most of the steroid-oxidizing enzymes are constitutive and their
levels are relatively invariable, in contrast to the xenobiotic-metabolizing P450s, the levels
of which can vary considerably (Guengerich and Montellano 2005).
The organ that expresses the highest levels of P450s is the liver, which plays the dominant
role in the first-pass clearance of ingested xenobiotic compounds and controls the systemic
level of drugs and other chemicals. Extrahepatic tissues, especially those that are the
portals of entry for xenobiotic compounds, such as the respiratory and the gastrointestinal
tracts, also express P450s. In these tissues, 450 enzymes not only contribute to the firstpass
clearance, but may also influence the tissue burden of xenobiotic compounds or
bioavailability of therapeutic agents (Ding and Kaminsky 2003).
1.2.6 Engineered P450s
The catalytic versatility, substrate diversity, and sheer number of P450s have led to
considerable interest over the last 25 years in the engineering of P450 enzymes for a
21

INTRODUCTION
variety of purposes (Gillam 2008). Mammalian P450 enzymes, as membrane-bound twoenzyme
systems, are notoriously difficult to express and hard to maintain in an active form.
Parallel heterologous expression in addition must provide exact ratios of components to
ensure catalytic activity (Vail et al. 2005). P450s can be engineered using rational and
evolutionary approaches. Rational approaches are characterized by the deliberate mutation
of one or more amino acids based on mechanistic or structural information and are
appropriate for altering P450 selectivities (Jung et al., Urlacher and Girhard). By directed
evolution, mutations are introduced in a random or semi-random manner, for example by
site-saturation mutagenesis at residues thought to be important for the desired property,
and the resulting mutant P450s are screened for enhancement of that property or set of
properties. A recent area of interest is the design and development of novel P450s as
biocatalysts of reactions of commercial interest, such as in pharmaceutical or fine chemical
synthesis (Gillam 2008).
For example, over the past decade, P450 BM3 was engineered for the oxidation of alkanes,
terpenes, heteroaromatics, alkaloids, steroids and other chemical substances, which are
either hydroxylated, epoxidized or demethylated (Jung et al., Urlacher and Girhard). P450
BM3 from Bacillus megaterium (discovered in the 1980s (Warman et al. 2005)) possesses
favorable properties that makes it an attractive target for engieneering (Munro et al. 2002).
It is a soluble bacterial, highly active enzyme, which is naturally fused to its reductase and
routinely expressed in E. coli. The hydroxylation of arachidonic acid catalyzed by P450
BM3 is the most rapid P450-catalyzed hydroxylation known so far (k cat = 17,000 min -1 )
(Jung et al., Warman et al. 2005).
In the 1990s the technique of DNA shuffling (Crameri et al. 1998, Stemmer 1994),
involving homology-dependent recombination of fragments of orthologous P450 genes in a
primerless PCR reassembly, was applied for P450-engineering with great success (Gillam
2008). A novel approach for DNA recombination (the so-called SCHEMA algorithm) was
used to obtain chimeras based on P450 BM3 and its homologs CYP102A2 and CYP102A3
from Bacillus subtilis. A survey of the activities of new chimeras demonstrated that this
approach created completely new functions absent in the wild-type enzymes, including the
ability to oxidize drugs (Urlacher and Girhard).
1.2.7 Catalyzed reactions
P450 enzymes play two main roles in living organisms. Some cytochrome P450s catalyze
oxidation steps involved in the biosynthesis or biodegradation of endogenous compounds
like steroids, fatty acids or prostaglandins. The second group is responsible for oxidative
22

INTRODUCTION
biotransformation of exogenous substances from the environment, facilitating their
elimination from living organisms (Mansuy 1998). Cytochrome P450s catalyze more than
20 different reaction types (Sono et al. 1996). Some of the "atypical" reactions are
oxidations involving C-C or C=N bond cleavage, reductions, dehydrations,
dehydrogenations or isomerizations. The classical reactions performed by P450s involve
the transfer of one oxygen atom from dioxygen (O 2 ) to the substrate.
R
+
+
− H + O2 + NAD( P)
H + H → R − OH + H
2O
+ NAD(
P)
1.2.8 The most important P450 reactions are summarized in Table 5.
Reaction mechanism of P450s
The catalytic cycle of P450 has been studied over decades. The active center for catalysis
is the iron(III)-protoporphyrin IX (heme) with the thiolate of the conserved cysteine
residue as fifth ligand (Werck-Reichhart and Feyereisen 2000) (Figure 4). Four pyrrole
rings are bonded by methine bridges (-CH 2 =), giving a planar and highly conjugated
porphyrin system (Nagababu and Rifkind 2004). Six
heteroatoms coordinate the heme iron octahedrally. The
four porphyrin nitrogens are the equatorial ligands. The
remaining two axial ligands are located below (fifth
coordination site, proximal) and above (sixth coordination
site, distal) the plane of the heme (Cirino and Arnold
2003).
Figure 4 Iron (III) protoporphyrin-IX linked with a proximal cysteine ligand. Modified according to
Meunier et al. (Meunier et al. 2004).
The P450 reaction cycle has been proposed by several authors(Guengerich 2001, Munro et
al. 2007, Ogliaro et al. 2002) and can be summarized as follows (Figure 5). It is initiated
by the binding of a substrate, usually with concomitant displacement of the distal water
ligand [(a), H 2 O…heme(Fe III )] (Ortiz de Montellano et al. 2005, Sligar and Gunsalus
1976). The substrate binds to the species (a) near the distal region of the heme, which
causes a dehydration of the active site resulting in a five-coordinated heme iron within the
enzyme-substrate complex [(b), R-H…heme(Fe III )].
23

MATERIALS AND METHODS
24
Table 5 Oxidative and non-oxidative reactions catalyzed by P450 enzymes(Guengerich 2001, Isin and Guengerich 2007, Mansuy 1998, Sono et al. 1996)
INTRODUCTION

INTRODUCTION
25

INTRODUCTION
MATERIALS AND METHODS
26

INTRODUCTION
The latter accepts a first electron that is derived from NAD(P)H and supplied via
flavin/ferredoxin or diflavin reductases to form the ferrous state of the enzyme [(c), (R-
H)…heme(Fe II )] (Groves 2006, Ullrich and Hofrichter 2007). The ferrous complex is
reactive enough to add dioxygen (O 2 ) resulting in the formation of a ferrous dioxycomplex
[(d), R-H…heme(Fe II )-O 2 ], which then accepts the second electron (Groves 2006,
Ullrich and Hofrichter 2007). This results in formation of a ferric peroxy ion [(e), (R-
H)…heme(Fe III -O - 2 )], which is protonated to form the hydroperoxy complex (also referred
as Compound 0, [(f), (R-H)…heme(Fe III -O-OH)]. Compound 0 undergoes heterolytic
cleavage between the oxygen atoms giving rise to the putative oxo-ferryl state [(g), (R-
H)…heme(Fe IV =O) •+ ], referred to as Compound I (Rittle and Green 2010).
Figure 5 The catalytic cycle of cytochrome P450s monooxygenases, including the peroxide “shunt”
pathway. RH is the substrate and ROH the product of the reaction. The rhombus symbolizes the
heme. Modified according to Ullrich and Hofrichter (Ullrich and Hofrichter 2007) and Werck-
Reichhart and Feyereisen (Werck-Reichhart and Feyereisen 2000). (a) native (hydro)ferric enzyme
(resting state), (b) ferric heme-substrate complex, (c) ferrous heme-substrate complex, (d) ferrousdioxygen
complex, (e) ferric peroxy anion complex, (f) ferric hydroperoxy complex (Compound
0), (g) putative oxy-ferryl radical complex (Compound I), (h) product-ferric enzyme complex.
27

MATERIALS AND METHODS
This electron-deficient complex may abstract a hydrogen atom or one electron from the
substrate to form a sigma complex (g) with the substrate. A subsequent collapse of the
intermediate or an intermediate pair in step (g) generates the product. (In the case of a
hydrogen abstraction mechanism, step (g) is referred to as an “oxygen rebound.”) In step
(h), the product dissociates from the enzyme, and the cycle can restart (Guengerich 2001,
Isin and Guengerich 2007). The so called “shunt” pathway is a remarkable side reaction of
many P450s, in the course of which substrate is directly oxidized by hydrogen peroxide (or
an organic peroxide) to the hydroperoxo-ferryl state (f) (Ullrich and Hofrichter 2007).
1.3 Peroxygenases
Since February 2011, a second sub-subclass (EC 1.11.2.x) of peroxidases (EC 1.11.x) has
been listed in the Enzyme Nomenclature system. The enzymes of this sub-subclass are
named peroxygenases and catalyze reactions with a peroxide as acceptor, one oxygen atom
of which is incorporated into the product
(http://www.chem.qmul.ac.uk/iubmb/enzyme/EC1/11/2/). This means peroxygenases
catalyze the direct transfer of one oxygen atom from a peroxide to the substrate (→ monoperoxygenation).
The peroxide (preferably hydrogen peroxide, H 2 O 2 ) is reduced and the
substrate is oxidized (Hanano et al. 2006):
RH + H
2
O2
→ ROH + H
2O
On a functional level, peroxygenases can be regarded as hybrids of cytochrome P450
monooxygenases and "classic" heme peroxidases.
The sub-subclass of peroxygenases contains four entries: EC 1.11.2.1 (unspecific
peroxygenase, the key enzyme of this group), EC 1.11.2.2 (myeloperoxidase), EC 1.11.2.3
(plant seed peroxygenase) and EC 1.11.2.4 (fatty-acid peroxygenase). However, it has to
be mentioned that in the other sub-subclass (EC 1.11.1: peroxidases) an enzyme is listed -
chloroperoxidase (CfuCPO, EC 1.11.1.10) - that catalyzes, in addition to halogenation
reactions, also a number of peroxygenase-type reactions (Allain et al. 1993, Omura 2005).
CfuCPO, which was already discovered 50 years ago [(Hager et al. 1966), belongs as do
the P450s to the heme-thiolate proteins and is apparently produced only by the
ascomycetous sooty mold Caldariomyces fumago (Omura 2005).
28

INTRODUCTION
All unspecific peroxygenases 2 characterized so far are also heme-thiolate proteins
secreted by agaric basidiomycetes (colloquially called mushrooms). They are very stable,
highly glycosylated proteins that have been shown to [per]oxygenate and oxidize various
organic substances (for details, see 1.3.1 below). Myeloperoxidase (MPO; until 2011,
listed under EC 1.11.1.7) contains calcium and a covalently bound heme with histidine as
proximal ligand (Booth et al. 1989, Davey and Fenna 1996). It is present in
mammalian/human phagosomes of neutrophils and monocytes (Klebanoff 1999). MPO
catalyzes the hydrogen peroxide mediated peroxygenation of halide ions (chloride,
bromide, iodide) and of the pseudohalide thiocyanate, according to the following reaction
(Harrison and Schultz 1976):
H
2
O
2
X = Cl
+ X
−
−
, Br
+ H O
−
, I
−
3
+
, SCN
→ HOX
−
+ 2H
2
O
Products of this reaction (hypohalous acids) and their following products are strong
oxidants and responsible for killing phagocytized bacteria and viruses. MPO differs from
CPO (EC 1.11.1.10) in its preference for formation of free hypochlorite over the
chlorination of organic substrates under physiological conditions (pH 5-8). Hypochlorite in
turn forms a number of antimicrobial products (Cl 2 , chloramines, hydroxyl radical, and
singlet oxygen). In the absence of halides, MPO can oxidize phenols and exhibits a
moderate peroxygenase activity towards styrene (Fiedler et al. 2000, Klebanoff 2005,
Tuynman et al. 2000).
Plant seed peroxygenase is a heme protein with a calcium binding motif of the caleosintype.
Enzymes of this type include membrane-bound proteins found in seeds of different
plants. It catalyzes the direct transfer of one oxygen atom from an organic hydroperoxide
(R-OOH), which is reduced to the corresponding alcohol, to a substrate that will be
oxidized. Reactions catalyzed include hydroxylation, epoxidation and sulfoxidation.
Preferred substrate and co-substrate are unsaturated fatty acids and fatty acid
hydroperoxides, respectively. Plant seed peroxygenase is thought to be involved in the
synthesis of cutin (Hanano et al. 2006, Lequeu et al. 2003).
Fatty acid peroxygenase is a bacterial, cytosolic heme-thiolate protein with sequence
homology to P450 monooxygenases. Unlike the latter, it needs neither NAD(P)H,
2 In earlier publications, unspecific peroxygenase was also referred to as haloperoxidase, haloperoxidaseperoxygenase,
mushroom/fungal peroxygenase, AaP (Agrocybe aegerita peroxidase/peroxygenase), CrP
(Coprinellus radians peroxygenase) or APO (aromatic peroxygenase). In February 2011, these enzymes were
classified as unspecific peroxygenase in the Enzyme Nomenclature system (EC 1.11.2.1; systematic name:
substrate:hydrogen peroxide oxidoreductase, RH-hydroxylating or -epoxidizing; for more details see
following web page: http://www.chem.qmul.ac.uk/iubmb/enzyme/EC1/11/2/1.html).
29

MATERIALS AND METHODS
dioxygen nor specific reductases for function. Enzymes of this type are produced by
eubacteria (e.g. Sphingomonas paucimobilis, P450 SPα and Bacillus subtilis, P450 BSβ ) and
catalyze the hydroxylation of long chain fatty acids (e.g. myristic acid) using hydrogen
peroxide as oxidant/electron acceptor to produce hydroxylated fatty acids (Imai et al. 2000,
Lee et al. 2003, Shoji et al. 2010).
Caldariomyces fumago chloroperoxidase (CfuCPO) catalyzes the unspecific chlorination,
bromination, and iodation of a variety of electrophilic organic substances via hypohalous
acids as actual halogenating agent. In the absence of halide, CfuCPO mimics
peroxygenases and P450s by catalyzing allylic and benzylic hydroxylations, alcohol
oxidations, heteroatom dealkylations, and oxo transfer to alkenes, alkynes, alkyl- and
arylamines, and sulfides; however, CfuCPO does not peroxygenate aromatic rings or
alkanes (Hofrichter and Ullrich 2006, Hofrichter et al. 2010, Hrycay and Bandiera 2012).
1.3.1 Fungal unspecific peroxygenases
Like CfuCPO, fungal peroxygenases are functional hybrids of P450 monooxygenases and
heme-peroxidases, but with a more pronounced oxygen transfer activity. Agrocybe
aegerita aromatic peroxygenase (AaeAPO) was the first enzyme of this type discovered
in 2004 and is, for the time being, the best-characterized aromatic/unspecific
peroxygenase. At first, this enzyme was reported to oxidize aryl alcohols to the
corresponding aldehydes and hence to the corresponding benzoic acids. AaeAPO was
found to catalyze also the conversion of typical peroxidase substrates, such as ABTS (2,2’-
azinobis-(3-ethylbenzothiazoline-6-sulfonate) or DMP (2,6-dimethoxyphenol);
furthermore, it brominates MCD (monochlordimedon) and phenol (Ullrich et al. 2004).
Later it turned out that the enzyme epoxidizes/hydroxylates toluene and naphthalene over a
wide pH range (3-10) with an optimum around pH 7 (Kluge et al. 2007, Ullrich and
Hofrichter 2005). Further studies showed that AaeAPO has a very broad reaction spectrum
that includes, for example, sulfoxidations (e.g. of dibenzothiophene, thioanisole), N-
oxidations (e.g. of pyridine), the O-dealkylation of diverse ethers (e.g. of tetrahydrofuran,
diethyl ether, alkyl aryl ethers), the O-demethylenation of benzodioxoles, N-dealkylations
(e.g. of secondary and tertiary amines), the epoxidation/hydroxylation of polycyclic
aromatic hydrocarbons (PAHs), aliphatic hydroxylations (e.g. of n-alkanes) and the
epoxidation of alkenes (Aranda et al. 2009a, Aranda et al. 2009b, Kinne et al. 2009a,
Kinne et al. 2009b, Kinne et al. 2008, Kinne et al. 2010, Kluge et al. 2007, Kluge et al.
2012, Peter et al. 2011, Poraj-Kobielska et al. 2011b, Ullrich 2008, Ullrich et al. 2008,
Ullrich and Hofrichter 2007). In the course of several studies, numerous basidiomycetes
have been screened for peroxygenase acitivity. As an outcome, the second true
30

INTRODUCTION
peroxygenase was discovered in 2006 in the inky caps Coprinopsis verticillata (CveAPO)
and Coprinellus radians (CraAPO) (Anh 2008). The latter enzyme was characterized in
2007 and shown to oxidize veratryl and benzyl alcohol as well as ABTS, DMP and
guaiacol into the corresponding aldehydes/benzoic acids and quinones, respectively.
CraAPO is able to brominate phenol and to regioselectively epoxidize/hydroxylate
naphthalene to form preferably 1-naphthol (Anh et al. 2007). Further studies demonstrated
that CraAPO catalyzes the sulfoxidation and hydroxylation of dibenzothiophene, the
[per]oxygenation of various PAHs, as well as the O- and N-dealkylation of many
pharmaceuticals (Aranda et al. 2009a, Aranda et al. 2009b, Poraj-Kobielska et al. 2011b).
An additional interesting peroxygenase was found in the pinwheel mushroom, Marasmius
rotula. This enzyme (MroAPO), which is smaller than the other peroxygenases (32 kDa
instead of 40-45 kDa), can oxidize all typical peroxidase and peroxygenase substrates such
as ABTS, DMP, veratryl and benzyl alcohols, naphthalene, and pyridine. Furthermore, it
hydroxylates and O-/N- dealkylates many pharmaceuticals including steroids that are not
converted by the other peroxygenases (Gröbe et al. 2012, Yarman et al. 2012) (Poraj-
Kobielska, 2012 unpublished results).
The full-length sequences of the peroxygenases of A. aegerita, C. radians and M. rotula
have recently been obtained. The sequences of AaeAPO, CraAPO and MroAPO do not
show any homology to classic heme peroxidases (such as lignin, manganese or horseradish
peroxidase) or to cytochrome P450 monooxygenases, and display less than 30% sequence
homology to CfuCPO (Pecyna et al. 2009). The CraAPO sequence shows 59% identity and
69% similarity to AaeAPO, and 27% identity and 42% similarity to CfuCPO. The sequence
of MroAPO differs considerably from those of the other known fungal peroxygenases,
showing only 33% identity and 49% similarity to AaeAPO, 29% identity and 43%
similarity to CfuCPO, and 31% identity and 45% similarity to CraAPO (Pecyna 2012).
Last but not least it should be mentioned that hundreds of peroxygenase-like sequences
from all classes of true fungi (Eumycota: Basidio-, Asco-, Glomero-, Zygo-,
Chytridiomycota) as well as from Oomycota (e.g. Phytophtora spp.) can be found in
genetic data banks (Hofrichter et al. 2010, Pecyna et al. 2009). The significance of these
putative peroxygenases remains to be investigated.
1.4 Aims and objectives
In recent years, increasing scientific attention has been given to pharmaceuticals and their
metabolites. Both their effects on the human body and the environment are the subject of
physiological, toxicological and ecotoxicological research approaches, dealing in particular
with the fate of drugs. To bring more light into this field, it is important to know the
31

MATERIALS AND METHODS
properties of drug metabolites and their effects on living organisms. On the other hand,
there is a need to remove such substances from the environment in order to reduce their
risk potential. In many organisms, cytochrome P450 enzymes play a key role in the
conversion and detoxification of bioactive compounds including pharmaceuticals.
However, P450s are intracellular biocatalysts and consequently relatively unstable. They
need expensive cofactors and are difficult to purify, which hampers their broad use in an
isolated form. For a few years it has been known that fungal peroxygenases can catalyze
similar reactions to those catalyzed by P450s, and several studies have demonstrated their
great biotechnological potential for oxyfunctionalizations. The peroxygenases are
extracellular - i.e. they are actively secreted by the fungus - and highly stable, and they do
not need cofactors other than hydrogen peroxide. Peroxygenases catalyze a wide variety of
reactions, including hydroxylations, epoxidations, and dealkylations. These catalytic
properties suggest that they may also be an interesting alternative for the enzymatic
conversion of pharmaceuticals both in terms of metabolite preparation and drug removal.
Accordingly, my PhD work has addressed the following questions:
1. Are fungal peroxygenases (APOs) capable of oxygenating complex
pharmaceuticals? If so, what are the limitations in terms of substrate structure and
size?
2. What are the products of these conversions?
3. Are these products similar to human drug metabolites formed by P450s?
4. Are illicit drugs and narcotics subject to peroxygenase attack?
5. Do peroxygenases of different fungal origin produce different patterns of
metabolites?
6. Are peroxygenases able to oxidize pharmaceuticals at low, environmentally
relevant concentrations?
7. Can peroxygenases be used in long term experiments over several days/weeks? If
so, what is their performance under such conditions?
Finally, on the basis of the answers to the questions posed above, this study has aimed to
develop perspectives for the application of fungal peroxygenases in the pharmaceutical
industry and the water treatment sector.
32

MATERIALS AND METHODS
2 Materials and Methods
2.1 Chemicals and reactants
2.1.1 Pharmaceuticals and their metabolites
Metoprolol, oseltamivir phosphate, 4-hydroxytolbutamide, 4'-hydroxydiclofenac, 5-
hydroxydiclofenac, 3-hydroxyacetaminophen, omeprazole, and sildenafil were obtained
from Chemos GmbH (Regenstauf, Germany). 3-Hydroxycarbamazepine, 1-
hydroxyibuprofen, 2-hydroxyibuprofen, 1-oxoibuprofen and N-desmethylsildenafil were
purchased from Toronto Research Chemicals Inc. (Toronto, Canada), and α-
hydroxymetoprolol, glycinexylidide and monoethylglycinexylidide from TLC
PharmaChem. Inc. (Vaughan, Canada). 17α-Ethinylestradiol, (R)-naproxen, 4-
hydroxypropranolol, 5-hydroxypropranolol, (S)-N-desisopropylpropranolol and O-
desmethylmetoprolol were obtained from Fluka (St. Gallen, Switzerland), from Shanghai
FWD Chemicals Ltd. (Shanghai, China), Biomol GmbH (Hamburg, Germany), SPIbio,
Bertin Group (Montigny Le Bretonneux, France), ABX Advanced Biochemical
Compounds (Radeberg, Germany) and Sandoo Pharmaceuticals & Chemicals Co., Ltd.
(Zhejiang, China), (S)-mephenytoin, S-desmethyl mephenytoin and 4’-hydroxy
mephenytoin were purchased from Santa Cruz Biotechnology Inc. (Heidelberg, Germany)
respectively. H 18 2 O 2 (90 atom %, 2% wt/vol) was a product of Icon Isotopes (New York,
USA). All other chemicals were purchased from Sigma-Aldrich (Schnelldorf, Germany).
Phenacetin-d 1 (N-(4-[1- 2 H]ethoxyphenyl)acetamide) was prepared from acetaminophen
(N-(4-hydroxyphenyl)acetamide) and ethyl iodide-d 1 as described previously for
phenacetin-d 3 (Garland et al. 1977). The reaction product of the synthesis (phenacetin-d 1 )
was identified by comparison of retention time, UV absorption spectra, and mass spectra
relative to authentic phenacetin (Nottebaum and McClure 2010); mass spectrum (m/z, %):
180 (100, - H), 138 (62, - C 2 H 2 O), 110 (12, - C 4 DH 6 O), 109 (96, - C 3 DH 2 O 2 ), 108 (100, -
C 3 DH 6 ON), 80 (16), 43 (18).
2.1.2 Drugs and their metabolites
Codeine hydrochloride, dextroamphetamine hydrochloride, (R,S)-Methamphetamine, 3,4-
methylenedioxyamphetamine (tenamphetamine), norcodeine, benzoylecgonine, lysergic
acid diethylamne, flurazepam, midazolam and Δ9-tetrahydrocannabinol were purchased
from Lipomed GmbH (Weil am Rhein, Germany). All other drugs and metabolites were
obtained from Sigma-Aldrich (Schnelldorf, Germany).
33

MATERIALS AND METHODS
2.1.3 Steroids and anabolic agents
Cortisone, pregnenolone and cholesterol were purchased from TCI Deutschland GmbH
(Eschborn, Germany). Testosteron was obtained from Fisher Scientific GmbH (Schwerte,
Germany). All other steroids, anabolic steroids and metabolites were obtained from Sigma-
Aldrich (Schnelldorf, Germany).
2.1.4 Benzodioxole derivatives
Safrole, sesamol, myristicin, 3,4-methylenedioxymethamphetamine (MDMA, ecstasy) and
5-chloro-1,3-benzodioxole were purchased from Sigma-Aldrich (Schnelldorf, Germany).
5-Nitro-1,3-benzodioxole (3,4-methylenedioxynitrobenzene, MDONB, NBD), 4-Bromo-
1,2-methylenedioxybenzene (3,4-methylenedioxybromobenzene) were obtained from TCI
Deutschland GmbH (Eschborn, Germany). Pisatin was obtained from Apin Chemicals Ltd.
(Abington, United Kingdom).
2.2 Fungal cultivation and purification of peroxygenases
2.2.1 Agrocybe aegerita and its peroxygenase
Agrocybe aegerita (Black poplar mushroom) is a typical agaric fungus, whose fruiting
bodies consist of a stalk and a cap with lamellae (gills) on the underside (Fig. 6; Foto by
René Ullrich). The fruiting bodies are edible and
have been cultivated in Mediterranean countries
for centuries. A. aegerita preferably colonizes
hardwood (Populus spp., Acer spp.) where it
causes an atypical white-rot, but also mulch and
straw-like materials (Stamets 1993).
The extracellular peroxygenase of A. aegerita
(AaeAPO; isoform II, 44 kDa) was produced and
purified as described previously (Ullrich et al.
2004). Agrocybe aegerita (fungal strain: A1,
isolated from a straw-pile in Jena/Wöllnitz in
1970) was cultivated on agar plates and in liquid
cultures, the culture liquid of which was
ultrafiltered and concentrated.
Figure 6 Agrocybe aegerita (Photo by René Ullrich)
34

MATERIALS AND METHODS
These crude enzyme preparations were further purified by three steps of ion exchange
chromatography using the strong cation exchangers SP Sepharose FF and Mono S,
and the strong anion exchanger Mono Q. All chromatographic steps were performed
with an ÄKTA FPLC system (Fast Protein Liquid Chromatography, GE Healthcare
Europe GmbH, Freiburg, Germany). After every exchange step, enzyme-containing
fractions were collected, ultrafiltered and desalted by a 150-mL or 20-mL stirred cell
system (10 kDa cut-off modified polyethersulphone membrane, Pall Life Sciences,
Dreieich, Germany). After the purification, the molecular mass of enzyme isoforms was
determined by SDS-PAGE (Novex XCell; Invitrogen GmbH, Karlsruhe, Germany) using a
12% NuPage gel. Protein concentration was determined with a low-molecular-mass
calibration kit served as a protein standard (MBI Fermentas®, St. Leon Roth, Germany).
The same electrophoresis system was used for analytical isoelectric focusing (IEF).
Electrophoretically separated protein bands were visualized with the Colloidal Blue
Staining Kit (Invitrogen). Final enzyme preparations were sterile-filtered, portionized and
frozen at -80°C.
2.2.1.1 Fungal cultivation
Fungal stock cultures (A. aegerita strain A1; DSM 22459) were maintained on 2% maltextract
agar (MEA) plates and kept in the dark at 4°C (Pecyna et al. 2009). For subsequent
studies, fungal precultures were prepared by excising one agar plug (∅ 1 cm) from the
stock culture, transferring it onto fresh MEA plates and keeping the cultures at 24°C for 2-
3 weeks. Afterwards, 27 500-mL flasks containing 6 g of soybean flour (Hensel, Magstadt,
Germany) in 200 mL of destilled water, were sterilized and inoculated with the contents of
one fungal preculture homogenized in 80 mL sterile water (0.05 % v/v). Fungal cultures
were agitated on a rotary shaker (100 rpm) at 24°C for 2-3 weeks.
2.2.1.2 Ultrafiltration
The fungal mycelium was removed by filtration (filter GF6; Schleicher & Schuell, Dassel,
Germany) and the enzyme-containing liquid was concentrated 60-fold by two steps of
ultrafiltration using a tangential-flow cassette system (Ultrasette, cut-off 10 kDa, Pall
GmbH, Distributor VWR GmbH, Darmstadt, Germany).
2.2.1.3 SP Sepharose FF separation
The pH of the crude enzyme preparation was adjusted to 4.75 and the proteins were
separated on the strong cation exchanger SP Sepharose Fast Flow (column XK 26/20, GE
35

MATERIALS AND METHODS
HealthCare). Sodium acetate buffer (10 mM, pH 4.75) was used as a mobile phase and 10
mM sodium acetate buffer containing 2 M sodium chloride (pH 4.75) was used as the
eluent. The column was operated at a flow rate of 13 mL min -1 and fractions (10 mL) were
collected. The elution of protein was performed with a linear gradient from 0% to 50%
eluent in 8 column volumes (modified according to (Ullrich et al. 2004)).
2.2.1.4 Mono Q separation
The pH of the enzyme preparation was adjusted to 6.0, and the mixture separated by a
Mono Q 10/100 GL column. Separations were carried out with sodium acetate buffer (10
mM, pH 6.0) as mobile phase and sodium acetate buffer (10 mM, pH 6.0) containing
sodium chloride (2 M) as eluent. Fractions (2 mL) were collected at a flow rate of 4 mL
min -1 . The elution of protein was achieved with a linear gradient from 0% to 30% eluent in
7 column volumes.
2.2.1.5 Mono S separation
The pH of the enzyme preparation was adjusted to 4.75, and the mixture separated by a
Mono S 10/100 GL column. Separations were carried out with sodium acetate buffer (10
mM, pH 4.75) as solvent, and sodium acetate buffer (10 mM, pH 4.75) containing sodium
chloride (1 M) as eluent. Fractions (2 mL) were collected at a flow rate of 6 mL min -1 . The
elution of protein was performed with a linear gradient from 0% to 50% eluent in 10
column volumes.
Figure 7 Elution profile recorded after ion exchange on the strong cation exchanger Mono S.
Separation of different AaeAPO isoforms (I, II, III, IV). For all further studies with AaeAPO, the
isoform II was used.
36

MATERIALS AND METHODS
2.2.1.6 Sodium dodecyl sulfate polyacrylamide gel
electrophoresis
(SDS-PAGE)
Molecular weights of proteins were determined by sodium dodecyl sulfate (SDS)-
polyacrylamide gel electrophoresis (PAGE) in a Mini-Cell system (XCell SureLock TM ,
Invitrogen GmbH, Karlsruhe, Germany) with Novex® Pre-Cast gels. The method is
described in the manufacturer’s instructions. After electrophoresis, the gels were stained
and protein bands were visualized with Colloidal Blue® (Invitrogen). A low molecular
weight protein calibration kit (MBI Fermentas, St. Leon Roth, Germany) was used as the
standard (Ullrich et al. 2004). Figure 8 shows exemplary electrophoretic separations: an
SDS-PAGE gel and an IEF gel of purified AaeAPO.
Figure 8 Separated AaeAPO isoform II visualized with the Colloidal Blue® Staining Kit in an
SDS-PAGE gel (left) and an IEF gel (right). M: marker, M W : molecular weight, pI: isoelectric
point.
2.2.1.7 Isoelectric focusing (IEF)
IEF separation was done in a Mini-Cell system (XCell SureLock TM , Invitrogen GmbH,
Karlsruhe, Germany) applying pre-cast IEF gels (pH 3-7) with IEF markers pH 3–10 and
37

MATERIALS AND METHODS
relevant buffers (Invitrogen GmbH, Karlsruhe, Germany), according to the manufacturer’s
instructions, and previous studies (Ullrich et al. 2009, Ullrich et al. 2004).
2.2.1.8 Summarized properties of the AaeAPO preparation
The enzyme preparation was homogeneous by SDS-PAGE, had a molecular mass of 46
kDa, a pI of 5.1 and exhibited an A 418 /A 280 ratio of 1.75 (1.86 3 , 1.68 4 ). The specific
activity of the AaeAPO preparation was 117 U mg -1 (65 U mg -1 3 , 85 U mg -1 4 ) where 1 U
represents the oxidation of 1 µmol of 3,4-dimethoxybenzyl alcohol (veratryl alcohol) to
3,4-dimethoxybenzaldehyde (veratraldehyde) in 1 min at 23 °C (Ullrich et al. 2004).
2.2.2 Coprinellus radians and its peroxygenase
This fungus prefers nitrogen-rich habitats and grows both on dung, straw-like materials
and wet wood. It is found not only outdoors, but also indoors on wet wood in basements
and bathrooms (Uljé 2001).
The strain used here, DSM 888 (ATCC 20014) isolated from horse dung, was purchased
from the DSMZ, Braunschweig,
Germany (Deutsche Sammlung von
Mikroorganismen und Zellkulturen
GmbH 2012). The unspecific
peroxygenase of Coprinellus radians
(CraAPO) was produced and purified as
described previously (Anh et al. 2007).
The fungus was cultivated on agar plates
and transferred to liquid cultures, and
the culture liquid thus obtained was
ultrafiltered and concentrated.
Figure 9 Fruiting bodies of Coprinellus radians (Photo by Hung Anh)
Crude enzyme preparations were further purified by two steps of anion exchange
chromatography (AEX) using Q Sepharose FF and Mono Q as separation media
followed by size exclusion chromatography (SEC, Superdex 75). All chromatographic
steps were performed with an ÄKTA FPLC system (Fast Protein Liquid
Chromatography, GE Healthcare Europe GmbH, Freiburg, Germany). After every
3 second purification
4 Isoform III that was used in long-time experiments (see 2.8.3)
38

MATERIALS AND METHODS
exchange step, enzyme fractions were collected, ultrafiltered and desalted with 150-mL or
20-mL stirred cell systems (10 kDa cut-off modified polyethersulphone membrane, Pall
Life Sciences, Dreieich, Germany). After purification, the enzyme was characterized. The
molecular mass of enzyme isoforms was determined by SDS-PAGE (Novex XCell;
Invitrogen GmbH, Karlsruhe, Germany) using a 12% NuPage gel. Protein concentration
was determined with a low-molecular-mass calibration kit (MBI Fermentas ® , St. Leon
Roth, Germany). The same electrophoresis system was used for analytical isoelectric
focusing (IEF). Electrophoretically separated protein bands were visualized with the
Colloidal Blue Staining Kit (Invitrogen GmbH, Karlsruhe, Germany). The final enzyme
preparations were sterile-filtered, portionized and frozen at -80°C.
2.2.2.1 Fungal cultivation
Fungal stock cultures of C. radians (DSM 888) were cultivated on MEA plates and kept in
the dark at 4°C. For subsequent studies, fungal precultures were prepared by excising one
agar plug (∅ 1 cm) from the stock culture, transferring it onto fresh MEA plates and
keeping the cultures at 24°C for 2-3 weeks. Afterwards, 27 500-mL flasks containing 6 g
soybean flour and 8 g glucose in 200 mL destilled water, were sterilized and inoculated
with the contents of a fungal preculture homogenized in 80 mL sterile water (0.05% v/v).
Fungal cultures were agitated on a rotary shaker (100 rpm) at 24°C for 2-3 weeks.
2.2.2.2 Ultrafiltration
The fungal mycelium was removed by filtration (filter GF6; Schleicher & Schuell, Dassel,
Germany) and the enzyme-containing liquid was concentrated 60-fold by two steps of
ultrafiltration using a tangential-flow cassette system (Ultrasette, cut-off 10 kDa, Pall
GmbH, Distributor VWR GmbH, Darmstadt, Germany).
2.2.2.3 Q Sepharose FF separation
The pH value of crude enzyme was adjusted to 6.0, and the proteins were separated on the
strong anion exchanger Q Sepharose Fast Flow (column XK 26/20, GE HealthCare) and
fractionated. As the mobile phase, 10 mM sodium acetate buffer (pH 6.0) was used and 10
mM sodium acetate buffer with 2 M sodium chloride (pH 6.0) served as the eluent. The
fraction size was 8 mL at a flow rate of 13 mL min -1 . The elution of protein was achieved
by a linear gradient from 0% to 50% eluent in 10 column volumes.
39

MATERIALS AND METHODS
2.2.2.4 Mono Q separation
The pH value of the enzyme preparation was adjusted to 5.5 and the mixture separated on a
Mono Q 10/100 GL column and fractionated. Separations were carried out with sodium
acetate buffer (10 mM, pH 5.5) as solvent, and sodium acetate buffer (10 mM, pH 5.5)
with sodium chloride (2 M) as eluent. The fraction size was 2 mL at a flow rate of 4 mL
min -1 . The elution of protein was performed with a linear gradient from 0% to 30% eluent
in 7 column volumes.
2.2.2.5 Size exclusion chromatography (SEC) using
Superdex 75
The pH value of the enzyme preparation was adjusted to 6.5, and the mixture separated by
a HiLoad 26/60 Superdex 75 preparative grade column and fractionated. As the mobile
phase and eluent, 50 mM sodium acetate buffer (pH 6.5) with 0.1 M sodium chloride (pH
6.5) was used. The fraction size was 4 mL at a flow rate of 2.5 mL min -1 . The elution of
proteins occurred under isocratic conditions.
Figure 10 Elution profile of CraAPO recorded
after size exclusion chromatography on a
Superdex TM 75 column.
2.2.2.6 Sodium dodecyl sulfate polyacrylamide gel
electrophoresis
(SDS-PAGE) and isoelectric focusing (IEF)
The methods used were the same as described for AaeAPO (2.2.1.6 and 2.2.1.7). Figure 11
shows as an example the SDS-PAGE gel of the purified Coprinellus radians peroxygenase
(CraAPO).
40

MATERIALS AND METHODS
Figure 11 Purified CraAPO visualized in an SDS-PAGE gel
with the Colloidal Blue staining kit. M: Marker, M W :
Molecular weight.
2.2.2.7 Summarized properties of the CraAPO preparation
The enzyme preparation was homogeneous by SDS-PAGE, had a molecular mass of 44
kDa, a pI of 4.2, and exhibited an A 419 /A 280 ratio of 1.04. The specific activity was 25.8 U
mg -1 , where 1 U means the oxidation of 1 µmol of 3,4-dimethoxybenzyl alcohol (veratryl
alcohol) to 3,4-dimethoxybenzaldehyde (veratraldehyde) in 1 min at 23 °C (Anh et al.
2007).
2.2.3 Marasmius rotula and its peroxygenase
Marasmius rotula (Pinwheel
mushroom) grows in deciduous
forests and fruits in groups or
clusters on dead wood (especially
beech), woody debris such as
twigs or sticks, and occasionally
on rotting leaves. M. rotula
peroxygenase was a gift from K.
Scheibner and G. Gröbe (Lausitz
University of Applied Sciences,
Senftenberg, Germany).
Figure 12 Marasmius rotula (Foto by René Ullrich)
41

MATERIALS AND METHODS
Cultivation of the fungus as well as enzyme purification and characterization were
described in (Gröbe et al. 2012). The enzyme had a molecular mass of 32 kDa, a pI
between 4.97 and 5.27, and a specific activity of 12.8 U mg -1 .
2.3 UV-Vis spectroscopy
2.3.1 Veratryl alcohol assay for peroxygenases
Figure 13 Veratryl
alcohol assay
The assay was routinely performed in quartz cuvettes with a light path of 10 mm. The
reaction mixture contained in a total of 1 mL the ingredients given in Table 5. The reaction
was started by addition of H 2 O 2 and followed at 310 nm and room temperature for at least
20 seconds. Enzyme activity was calculated using the initial linear increase in absorbance
and the extinction coefficient for veratryl aldehyde at 310 nm (ε 310 = 9,300 M -1 cm -1 ) (Tien
and Kirk 1984, Ullrich et al. 2004). Note that the same assay is used to measure the activity
of lignin peroxidase (LiP, EC 1.11.1.14), except that acidic tartrate buffer (pH 3.0) is used
for Lignin peroxidase instead of neutral phosphate buffer (Hofrichter et al. 2010).
Table 5 Composition of veratryl alcohol assay.
Compound
Solution
concentration (mM)
Amount (µL)
End
concentration (mM)
Potassium phosphate 100 500 50
buffer pH 7
Veratryl alcohol 50 100 5
H 2 O 290-389
Sample/ Enzyme
1-100
solution
H 2 O 2 100 10 1
42

MATERIALS AND METHODS
2.3.2 ABTS assay for laccase
Figure 14 ABTS assay for laccases
The assay was routinely performed in quartz cuvettes with a light path of 10 mm. The
reaction mixture contained in a total of 1 mL the ingredients listed in Table 6. The reaction
was started by the addition of the enzyme-containing sample to the mixture and was
followed at 420 nm and room temperature for over 60 seconds. Enzyme activity was
calculated using the initial linear increase in absorbance and the extinction coefficient for
the ABTS cation radical at 420 nm (ε 420 = 36,000 M -1 cm -1 ) (Wolfenden and Willson
1982).
Table 6 Composition of the ABTS [2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid)] assay.
Compound
Solution
concentration (mM)
Amount (µL)
End
concentration (mM)
Sodium citrate buffer 100 500 50
pH 4.5
ABTS 6 50 0.3
H 2 O 350-445
Sample/Enzyme
solution
5-100
2.3.3 Nitrobenzodioxole assay for peroxygenases
2.3.3.1 Spectrophotometric activity measurement at neutral pH
Figure 15 Nitrobenzodioxole assay for peroxygenases
This is a new assay developed in the course of this work (see 3.10) and it has been
published recently (Poraj-Kobielska et al. 2011a). Peroxygenases of Agrocybe aegerita,
43

MATERIALS AND METHODS
Coprinellus radians, Coprinopsis verticillata 5 , Marasmius rotula as well as recombinant
peroxygenase of Coprinopsis cinerea 6 (0.05-2 Units mL -1 ; ~0.02-10 µM) were individually
dissolved together with 5-nitro-1,3-benzodioxole [NBD; 1,2-(methylenedioxy)-4-
nitrobenzene (0.125-0.5 mM) 7 ] in potassium phosphate buffer (50 mM, pH 7.0) containing
acetonitrile (10% vol/vol) and, if the reaction products were to be quantified by HPLC,
ascorbic acid 8 (4.0 mM). After adding the co-substrate H 2 O 2 (1 mM), the reaction mixture
immediately changed its color to bright yellow (at pH 7 and 23 °C).
The assay was routinely performed in quartz cuvettes with a light path of 10 mm. The
reaction mixture contained in a total of 1 mL the ingredients given in Table 7. The reaction
was started by the addition of H 2 O 2 and followed at 425 nm and room temperature for 15-
30 seconds. Peroxygenase activity was calculated using the initial linear increase in
absorbance and the extinction coefficient for 4-nitrocatechol at 425 nm (ε 425 = 9,700 M -1
cm -1 ).
Table 7 Composition of the nitrobenzodioxole (NBD) assay.
Compound
Potassium phosphate
buffer pH 7
Solution Amount (µL)
End
concentration (mM)
concentration (mM)
100 500 50
NBD 50 100 5
H 2 O 290-389
Sample 1-100
H 2 O 2 100 10 1
2.3.3.2 Colorimetric detection under alkaline conditions
The colorimetric detection of 4-nitrocatechol can be performed as an end-point
determination and requires the adjustment of the assay solution mentioned above, after a
certain reaction time (5-10 min), to a pH >12, e.g. by addition of 100 µL sodium lye (10
M). In the presence of 4-nitrocatechol, the assay mixture will immediately change its color
to red and peroxygenase activity can be calculated by detecting the absorbance at 514 nm
(ε 514 = 11,400 M −1 cm −1 ).
5 gift of E. Aranda and R. Ullrich (International Graduate School of Zittau)
6 gift of H. Lund (Novozymes A/S, Copenhagen, Denmark)
7 NBD is hardly soluble in water and aqueous solvent mixtures at concentrations higher than 0.5 mM. Stock
solutions can be prepared by using pure acetonitrile as solvent.
8 Ascorbic acid is added to prevent the further oxidation of the product 4-nitrocatechol, which proceeds via
the peroxidative activity of A. aegerita peroxygenase (Kinne et al. 2009a, Kinne et al. 2008). The addition of
the radical scavenger is not necessary for the qualitative colorimetric detection of peroxygenases.
44

MATERIALS AND METHODS
2.3.3.3 Tests with environmental samples
Fresh samples of leaf-litter from a riparian Alnus forest near Zittau (Germany) as well as of
chopped wheat straw and beech wood were homogenized and 2 g of them extracted in
potassium phosphate buffer (50 mM, pH 7.0) for 24 h. The crude extracts were centrifuged
and the supernatants used for the experiment. The peroxygenase activity messurements in
the extracts showed no positive results. The test was performed analogue to the
nitrobenzodioxole assay (2.3.3.1) in quartz cuvettes with a light path of 10 mm. The
reaction mixture contained in a total of 1 mL the ingredients given in Table 8. The reaction
was started by the addition of H 2 O 2 and followed at 425 nm and room temperature for 15-
30 seconds. Peroxygenase activity was calculated using the initial linear increase in
absorbance and the extinction coefficient for 4-nitrocatechol at 425 nm (ε 425 = 9,700 M -1
cm -1 ).
Table 8 Composition of the nitrobenzodioxole (NBD) assay with environmental samples.
Compound
Solution
concentration (mM)
Amount (µL)
End
concentration (mM)
Potassium phosphate 100 500 50
buffer pH 7
NBD 50 100 5
extract 300
H 2 O 70
AaeAPO 20 7.2 U(VA) mL -1 / 13.2
U (NBD) mL -1
H 2 O 2 100 10 1
2.3.4 Determination of protein concentration
The determination of protein concentration was performed using the Bradford assay. This
is a colorimetric protein assay based on an absorbance shift of the dye Coomassie Brilliant
Blue G-250, in which under acidic conditions the red form of the dye is converted into its
blue form to bind with the protein being assayed. The (bound) form of the dye has an
absorption maximum historically held to be at 595 nm. The cationic (unbound) forms are
green or red and absorb at 450 nm.
For the calibration curve, standard protein (Albumin, Roth Chemie GmbH, Karlsruhe,
Germany) was prepared in following concentrations: 5, 10, 20, 30, 40, 50, 60, 80, 100 µg
mL -1 . 5 mL of the original dye solution Roti ® -Nanoquant (Roth Chemie GmbH, Karlsruhe,
Germany) was diluted in 20 mL of distilled water. A Greiner 96-well Flat Transparent
Plate was filled as follows: wells A1, A2, and A3 with 250 µL of water (blank), lines B to
H (1, 2, and 3) and lines A and B (4, 5, and 6) each with 50 µL of the calibration stock
45

MATERIALS AND METHODS
solutions. All calibration wells were filled up with 200 µL of the dye solution. 50 µL of the
protein samples were pipetted in one of the free wells and filled up with 200 µL of the dye
solution. The color was visually checked and compared with a calibration row. If the color
fitted into the calibration row, the sample was pipetted into three following wells (if not,
the sample was first diluted) and filled up with 200 µL dye solution. The plate was
measured with a multimode reader Infinite M200, TECAN® (Tecan Group Ltd.,
Männedorf, Germany) with the following method: shaking for 5 seconds at 44.3 rpm,
measurement at 590 nm, followed by measurement at 450 nm. The results were read as the
quotient 590/450 as mean concentration in µg mL -1 .
2.3.5 Difference spectra
Difference spectra were recorded in quartz cuvettes (light path 10 mm) in the scanning
range from 200 to 800 nm at room temperature using a Cary 50 spectrophotometer
(Varian, Darmstadt, Germany). The mixture contained 50 mM potassium phosphate buffer
(pH 7), 40 U mL -1 (13.75 µM) AaeAPO and 0.625 mM to 4.6 mM of the ligand. The
substrate ligands were phenol (as reference (Kinne 2010)), diclofenac, propranolol and
dextromethorphan. The instrument baseline was set to zero with the reference cuvette in
the sample beam. The reference cuvette contained 50 mM potassium buffer (pH 7) and 40
U mL -1 (13.75 µM) of AaeAPO.
2.4 Standard reaction conditions for peroxygenase conversions
Typical reaction mixtures (0.2-1.0 mL) contained purified peroxygenase (1.0-2.0 U mL -1 =
0.04-0.08 µM), substrate to be oxidized (0.5-2.0 mM), potassium phosphate buffer (50
mM, pH 7.0), and ascorbic acid (4.0-6.0 mM, to inhibit further oxidation of any phenolic
products that were released (Kinne et al. 2009a, Kinne et al. 2008). The reactions were
started by the addition of limiting H 2 O 2 (2.0-4.0 mM) and stirred at room temperature.
They were stopped by addition of sodium azide (1 mM) or trichloroacetic acid (TCA) (5%)
after 3 min. In some cases, H 2 O 2 was added continuously with a syringe pump and the
reactions were stopped after 4-6 h, when chromatographic analyses showed that product
formation was complete.
Aliphatic aldehydes were analyzed as their 2,4-dinitrophenylhydrazones after addition of
0.2 volume of 0.1% 2,4-dinitrophenylhydrazine solution in 0.6 N HCl to each reaction
mixture.
46

MATERIALS AND METHODS
2.5 Product identification
Reaction products of interest were detected, generally with baseline resolution, by high
performance liquid chromatography (HPLC) using an Agilent Series 1200 instrument
equipped with a diode array detector (DAD) and an electrospray ionization mass
spectrometer (MS) (Agilent Technologies Deutschland GmbH, Böblingen, Germany).
Formic acid was analyzed by ion exchange chromatography (IC) using a Dionex Series
ICS 1100 system (Thermo Fisher GmbH, Idstein, Germany).
2.5.1 HPLC method I
For the experiments with acetaminophen, acetanilide and diclofenac, a Synergi 4µ Fusion
reversed-phase column (4.6 mm diameter by 150 mm length, 4 µm particle size, 80Ǻ,
Phenomenex, Aschaffenburg, Germany) was used. The column was run at 40°C and a flow
rate of 1 mL min -1 with a mixture of aqueous phosphoric acid solution (15 mM, pH 3) and
acetonitrile, 95:5, for 5 min, followed by a 20-min linear gradient to 100% acetonitrile.
2.5.2 UHPLC method
For the experiments with low concentrated propranolol, a Kinetex 2.6 µ C18 column (2.1
mm diameter by 150 mm length, 2.6 µm particle size, 100 Ǻ, Phenomenex, Aschaffenburg,
Germany) was used. The column was eluted at 0.3 mL min -1 and 40°C with aqueous
0.01% vol/vol ammonium formate (pH 3.5)/acetonitrile, 90:10, followed by a 3-min linear
gradient to 70% acetonitrile.
2.5.3 HPLC method for chiral separation
Chiral separation was done using the HPLC apparatus above, but using a Whelk-O 5/100
Kromasil column (4.6 mm diameter by 250 mm length, Regis Technologies (Morton
Grove, USA). The isocratic mobile phase consisted of 80% vol/vol methanol and 20% of
15 mM phosphate buffer. The columns were operated at 40°C and 1 mL min -1 for 30 min.
2.5.4 HPLC-MS method I
Unless otherwise stated, reversed-phase (RP) chromatography of reaction mixtures was
performed on a Luna C18 column (2 mm diameter by 150 mm length, 5 µm particle size,
100 Ǻ, Phenomenex, Aschaffenburg, Germany), which was eluted at 0.35 mL min -1 and
40°C with aqueous 0.01% vol/vol ammonium formate (pH 3.5)/acetonitrile, 95:5 for 5
min, followed by a 25-min linear gradient to 100% acetonitrile (for mass spectrometer
parameters see 2.5.7).
47

2.5.5 HPLC-MS method II
MATERIALS AND METHODS
The derivatized products were analyzed using a Luna C18 column. The column was eluted
with aqueous 0.1% vol/vol ammonium formate (pH 3.5)/acetonitrile, 70:30 for 5 min,
followed by a 20-min linear gradient to 100% acetonitrile (for mass spectrometer
parameters see 2.5.7)
2.5.6 HPLC-MS method III
For metoprolol, a Gemini-Nx 3µ 110A C18 reversed-phase column (2 mm diameter by
150 mm length, 3 µm particle size, Phenomenex) was used. The column was eluted at
45°C and a flow rate of 0.3 mL min -1 with a mixture of aqueous 0.2% vol/vol ammonium
(pH 10.5) and acetonitrile, 95:5, for 1 min, followed by a 20-min linear gradient to 80%
acetonitrile, followed by 21-min linear gradient to 100% acetonitrile (for mass
spectrometer parameters see 2.5.7).
2.5.7 Mass spectrometry for HPLC-MS methods
Mass spectrometric determinations were made in the positive or negative ESI mode
(electrospray ionization) in a mass range from 70 to 500, at a step size of 0.1, a drying gas
temperature of 350°C, a capillary voltage of 4,000 V for the positive mode and 5,500 V for
the negative mode. Reaction products were identified relative to authentic standards, based
on their retention times, UV absorption spectra, and mass spectral [M + H] + or [M - H] -
ions.
2.5.8 IC method for organic acids
Formic acid was analyzed by ion exchange chromatography (IC) on a Dionex Series ICS
1100 HPLC system fitted with a IonPac® ICE-AS1 column (4 mm diameter by 250 mm
length, Dionex, Germany). The column was eluted at 30°C and 1.6 mL min -1 with an
aqueous perfluorobutyric acid solution (1 mM) for 20 min. The product was identified
relative to an authentic standard by its retention time.
2.6 Stoichiometrical analyses
2.6.1 Sildenafil
Stoichiometrical experiments on sildenafil N-demethylation were conducted in stirred
reactions (0.20 mL) that contained 2 U mL -1 (0.144 µM) peroxygenase, potassium
phosphate buffer (50 mM, pH 7.0) and the substrate (0.25 mM). The reactions were
initiated by 0.0023-0.037 mM H 2 O 2. The resulting product (N-desmetyl sildenafil) was
48

MATERIALS AND METHODS
quantified by HPLC-MS as described above (HPLC-MS method I, 2.5.4) using the
external standard curve prepared with the authentic standard.
2.6.2 Safrole
The stoichiometrical experiment on safrole demethylenation occurred in a stirred reaction
mixture (0.20 mL) that contained 2 U mL -1 (0.144 µM) of peroxygenase, potassium
phosphate buffer (50 mM, pH 7.0), acetonitrile (10% vol/vol), ascorbic acid (4.0 mM) and
the substrate (0.25 mM). The reactions were initiated by 0.0125-0.200 mM H 2 O 2 . The
product (formic acid) was quantified by IC as described above, using an external standard
curve. The standard curve had linear regression values of R 2 >0.99.
2.7 Enzyme kinetics
2.7.1 Pharmaceuticals
The kinetics of propranolol hydroxylation as well as metoprolol and phenacetin O-
dealkylation were analyzed in stirred microscale reactions (0.10 ml, 23°C) that contained
0.40 µM peroxygenase (AaeAPO), potassium phosphate buffer (50 mM, pH 7.0), ascorbic
acid (4 mM) and 0.010-5.000 mM substrate. The reactions were initiated with 2.0 mM
H 2 O 2 . Reactions with propranolol or phenacetin were stopped with 0.010 mL of a 50%
TCA solution after 5 seconds, and reactions with metoprolol were stopped with 0.2 volume
of 0.1% 2,4-dinitrophenylhydrazine solution in 0.6 N HCl after 10 seconds. The resulting
products were quantified by HPLC as described above using the external standard curves
prepared with authentic standards.
The kinetics of acetanilide hydroxylation were analyzed in stirred reactions (0.5 mL, 23
°C) that contained 0.18 µM AaeAPO, potassium phosphate buffer (50 mM, pH 7.0),
ascorbic acid (4 mM) and 0.010-5.000 mM substrate. The reactions were initiated by 2.0
mM H 2 O 2 and stopped by 0.050 mL of a sodium azide solution (1 mM) after 5 seconds.
The initial velocity of acetaminophen formation was then determined from the increase in
absorbance at 290 nm (ε = 1,100 M -1 cm -1 ) using a Cary 50 UV/visible spectrophotometer.
The kinetic parameters were determined by nonlinear regression using the Lineweaver-
Burk model in the ANEMONA program (Hernandez and Ruiz 1998).
2.7.2 Nitrobenzodioxole
The kinetics of 5-nitrobenzo-1,3-dioxole demethylenation were analyzed in stirred reaction
mixtures (0.60 mL, 23°C ) that contained 0.036 µM (0.5 U mL -1 ) of AaeAPO, potassium
phosphate buffer (50 mM, pH 7.0), and 0.125-1.000 mM of the substrate. The reactions
49

MATERIALS AND METHODS
were initiated with 0.063-0.100 mM H 2 O 2 , and the initial velocity of 4-nitrocatechol
formation was measured by the increase in absorbance at 425 nm (ε = 9700 M -1 cm -1 )
using a Cary 50 UV-Vis spectrophotometer. Three kinetic data sets were obtained for each
pair of substrate concentrations. Kinetic parameters were determined by nonlinear
regression using the ping-pong model in the ANEMONA program (Hernandez and Ruiz
1998).
2.8 Experiments with 18 O-isotopes
2.8.1 Pharmaceuticals
The reaction mixtures (0.20 mL, stirred at room temperature) contained 2 U mL -1 (0.08
µM) of AaeAPO, potassium phosphate buffer (50 mM, pH 7.0), and 0.5 mM substrate
(tolbutamide, acetanilide, or carbamazepine). Reactions with tolbutamide or acetanilide
were initiated by a single addition of 2.0 mM H 18 2 O 2 (final concentration). For reactions
with carbamazepine, the same quantity of H 18 2 O 2 was added continuously with a syringe
pump over 6 hours. A portion of each completed reaction was then analyzed by HPLC-MS
as described above (HPLC-MS method I, 2.5.4). For each m/z value, the average total ion
count within the metabolite (4-hydroxytolbutamide, acetaminophen, 3-
hydroxycarbamazepine) peak was used after background correction to generate the ion
count used for mass abundance calculations.
2.8.2 Nitrobenzodioxole
The reaction mixtures (0.20 mL, stirred at room temperature) contained 2 U mL -1 (0.144
µM) of the peroxygenase, potassium phosphate buffer (50 mM, pH 7.0), ascorbic acid (4.0
mM, to inhibit further oxidation of the phenolic products that were released (Kinne et al.
2009a, Kinne et al. 2008) and 0.25 mM nitrobenzodioxole. The reaction was initiated with
2.0 mM H 18 2 O 2 . A portion of the mixture was analyzed by HPLC-MS as described above
(HPLC-MS method I, 2.5.4). Calculation of 18 O-incorporation was performed by dividing
the sum of the natural species abundance and the isotope abundance with the latter. The
10% of natural abundance H 2 O 2 in the H 18 2 O 2 preparation was taken into account in these
calculations.
2.9 Deuterium isotope effect experiment
The reaction mixture (0.2 mL) contained purified AaeAPO (2 U mL -1 , 0.08 µM),
potassium phosphate buffer (50 mM, pH 7.0), phenacetin-d 1 and ascorbic acid (4.0 mM).
The reaction was started by the addition of H 2 O 2 (2.0 mM) and stirred at room
50

MATERIALS AND METHODS
temperature. The reaction was stopped by addition of 10% sodium azide (10 mM) after 10
seconds. The reaction products were analyzed as described above (HPLC-MS method I,
2.5.4). For each m/z value, the average total ion count within the acetaminophen peak was
used after background correction to generate the ion count used for mass abundance
calculations.
2.10 Experiment with different peroxides and peroxide sources
The reaction mixture (0.20 mL, stirred at room temperature) contained: 2 U mL -1 (0.144
µM) of AaeAPO, potassium phosphate buffer (50 mM, pH 7.0), 4 mM ascorbic acid, 0.5
mM of diclofenac and 2 mM of glucose by test with glucose oxidase. Reactions were
initiated in three different ways: i) single addition, ii) multiple additions (four times in four
minutes), and iii) continuous addition with a syringe pump; the following peroxides or
peroxide-generating systems were tested:
1. 2 mM of hydrogen peroxide (H 2 O 2 ),
2. 2 mM of meta-chloroperoxybenzoic acid (mCPBA),
3. glucose oxidase in excess (~20 U mL -1 , in the presence of 2 mM glucose),
4. 2 mM of L-cysteine,
5. 2 mM of tert-butyl hydroperoxide, and
6. 2 mM of Cu(II) in form of copper(II) perchlorate hexahydrate [Cu(ClO 4 ) 2 × 6
H 2 O].
2.11 In vivo incubation experiment
Eighteen 500-mL round bottom flasks were filled with the two culture media described in
Table 9 (nine flasks of each medium, 200 mL).
Table 9 Composition of culture media
Culture medium (each 1,800 mL)
Soybean medium
Czapek-Dox medium
54 g (3%) soybean flour 1.8 g glucose
1,800 mL distilled water 5.4 g sodium nitrate
1.8 g potassium hydrogen
phosphate
0.9 g potassium chloride
0.018 g ferrous sulfate heptahydrate
0.9 g yeast extract
1,800 mL distilled water
51

MATERIALS AND METHODS
All eighteen flasks were autoclaved and numbered. Single flasks were prepared as
described in Table 10.
Table 10 Preparation of different cultures. a precultures of A. aegerita were prepared on MEA
plates and homogenized as described under 2.2.1.1; b instead, 9 mL distilled water was
supplemented; c 70% ethanol was used for the preparation of stock solution; d stock solutions: 100
mM of the substance in ethanol (70%)
Flask
no.
Czapek-Dox medium
Agrocybe
aegerita
homogenate a
(9 mL)
Culture type and
additional
substance
Flask
no.
Soybean medium
Agrocybe
aegerita
homogenate a
(9 mL)
Culture type and
additional
substance
4 + control, 1 mL of
1 + control, 1 mL of
70% ethanol c 70% ethanol c
2 - b control, 1 mL
propranolol d on
inoculation day
3 - b control, 1 mL
diclofenac d on
inoculation day
7 + sample, 1 mL
propranolol d on
inoculation day
8 + sample, 1 mL
diclofenac d on
inoculation day
11 + sample,1 mL
propranolol d one
week after
inoculation
12 + sample, 1 mL
diclofenac d one
week after
inoculation
15 + sample, 1 mL
propranolol d two
weeks after
inoculation
16 + sample, 1 mL
diclofenac d two
weeks after
inoculation
5 - b 1 ml propranolol d
on inoculation day
6 - b control,1 mL
diclofenac d on
inoculation day
9 + sample, 1 mL
propranolol d on
inoculation day
10 + sample, 1 mL
diclofenac d on
inoculation day
13 + sample, 1 mL
propranolol d one
week after
inoculation
14 + sample, 1 mL
diclofenac d one
week after
inoculation
17 + sample,1 mL
propranolol d two
weeks after
inoculation
18 + sample,1 mL
diclofenac d two
weeks after
inoculation
The culture flasks were incubated over sixteen days and agitated on a rotary shaker (100
rpm) at 24°C. Sampling (1 L) of liquid from fungal cultures and controls was done in a
clean bench on the day of substrate addition and the two following days. The cultures 7, 8,
9, and 10 were sampled one week after inoculation. All samples were centrifuged, 500 µL
were used for enzyme activity measurements, the rest was filtrated through Acrodisc® LC
52

MATERIALS AND METHODS
13 Syringe filters (13 mm with 0,2 µM PVDF membrane; Pall GmbH, Distributor VWR
GmbH, Darmstadt, Germany) and analyzed by the HPLC-MS method I (2.5.4).
2.12 Conversion of propranolol at environmentally relevant
concentration
Propranolol (1.5 mL; 5 µM = 5,000 nM) was added into 1,440 mL of distilled water. Six
500-mL Schott® flasks were filled each with 240 mL of this diluted solution, and 9 mL of
distilled water was added into three control flasks; into the other three flasks, 50 Units of
AaeAPO dissolved in 9 mL water was added. The six flasks were incubated at 24°C
overnight under mixing and continuous dosage of hydrogen peroxide (end concentration 2
mM) with a syringe pump. In the next step, remaining proteins were removed from the
samples; for this, the content from all six flasks was ultrafiltered through an Amicon Cell®
(with 10 kDa cut-off membrane). The membrane was conditioned with 5 µM propranolol
(to prevent/reduce absorption) and the pH was adjusted to 12 with 10 M NaOH in order to
facilitate subsequent extraction. The ultrafiltered samples were transferred to separation
funnels, extracted with dichloromethane by vigorous shaking for 5 minutes, and left
overnight for completion of phase separation. The separated organic phase was removed
and concentrated in the evaporator workstation TurboVap® (Zymark, USA) to a volume of
1 mL; then 2 mL of acetonitrile were added and the mixture concentrated again. In the last
step, 2 mL of water was added and the mixture concentrated again. The aqueous samples
were analyzed by HPLC as described above (2.5.2 and 2.5.4).
2.13 Long term experiments
2.8.2 Preliminary test with Vivaspin® (fed-batch method)
For this experiment, Vivaspin® 2 Centrifugal Concentrators (2 mL, with 10 kDa cut-off;
VivaScience Sartorius Group, Distributor VWR GmbH, Darmstadt, Germany) were used.
Two Vivaspins were filled each with 10 Units of AaeAPO. Two reaction mixtures (10 mL
each) were prepared, both containing 50 mM potassium phosphate buffer (pH 7), 0.50 mM
propranolol, and 2 mM of hydrogen peroxide and one of them in addition 4 mM of
ascorbic acid. Of both mixtures 1 mL was filled in parallel into the Vivaspins that were
then centrifuged with a table-centrifuge at 10,397 x g (Allegra 64R with Beckmann-Rotor
C1015; Beckmann-Coulter, Krefeld, Germany). This procedure was repeated eight times
on the first day and six times afterwards with one-day breaks in between (each with fresh
mixtures). The Vivaspins with the enzyme were repeatedly filled with fresh potassium
53

MATERIALS AND METHODS
phosphate buffer after the first day and samples were stored at 4°C. After every
centrifugation step, the enzyme activity was measured with the veratryl alcohol assay
(2.3.1) in the permeate, as were the product (5-hydroxypropranolol) and substrate
(propranolol) concentrations by HPLC-MS (method I; 2.5.4).
2.8.3 Long term experiment using a hollow fiber cartridge
Four 250-mL Schott ® flasks were filled with following solutions: (i) 2.5 mM propranolol;
(ii) 100mM potassium phosphate buffer, pH 7; (iii) 8 mM ascorbic acid; (iv) 6.7 mM
hydrogen peroxide. The outlet-tubes of the four flasks were connected with a quaternary
HPLC pump (Hewlett Packard series 1050) and the solutions pumped with a flow rate of
0.2 ml min -1 (25% of each solution) through a hollow fiber cartridge (UFP-5-C-MM01A,
GE Healtcare Europe GmbH, Freiburg, Germany). The cartridge was loaded before with
30 Units (0.354 mg; A spec =84.8) of AaeAPO. For two hours, from the very beginning,
samples were collected and the propranol and 5-hydroxypropranolol concentrations
measured every 15 minutes by HPLC-MS (Method I; 2.5.4). This procedure was repeated
for fourteen days. Every day fresh solutions (i), (ii), (iii), and (iv) were prepared and
pumped through the cartridge, but the enzyme in the cartridge was the same throughout the
whole experiment.
54

RESULTS
3 Results
Previous studies have shown that fungal peroxygenases catalyze various reactions similar to
P450 enzymes. Pharmaceuticals and illicit drugs are one important group of P450 substrates.
Their conversion by P450s usually increases solubility and bioavailability, which is
prerequisite for their detoxification or further degradation in the environment. On the other
hand, the drug metabolites formed are of relevance for drug development and toxicological
studies. In this section, the results of comprehensive tests using peroxygenase and a
representative number of pharmaceutically active substances are summarized and classified
according to the reaction type. Most substances were treated with peroxygenases from three
different fungal species. Thereafter, the results of kinetic experiments, stoichiometrical
analyses as well as of 18 O-labeling and deuterium-effect studies are presented. The next two
subchapters deal with a new enzymatic assay for the determination of peroxygenase activity
and experiments with selected steroids, in the course of which some catalytic limitations were
observed. The last part of the results section describes complementary experiments with
propranolol at low, environmentally relevant concentrations as well as long-term and in vivo
studies. Due to space limitations, additional information including exemplary chromatograms
and mass spectra can be found in the appendix section.
3.1 Aromatic ring hydroxylation
3.1.1 Ring hydroxylation of pharmaceuticals
Figure 16 provides an overwiew of pharmaceuticals converted by AaeAPO and CraAPO. In
many cases, ring hydroxylation was observed and sometimes the preferred reaction was
catalyzed by both APOs with high regioselectivity. For example, AaeAPO preferentially
oxidized propranolol (I) to 5-OH-propranolol (Kinne et al. 2009a), acetanilide (II) to
acetaminophen (paracetamol), carbamazepine (III) to 3-OH-carbamazepine, and diclofenac
(IV) to 4’-OH-diclofenac (Kinne et al. 2009a). CraAPO catalyzed the same reactions but with
lower regioselectivity. Both enzymes also hydroxylated ketoprofen, sulfadiazine and
chlorpromazine. Tamoxifen (table 15, XXVII) was oxidized by both peroxygenases yielding
low amounts of 4-OH-tamoxifen along with some other products, which we were unable to
resolve sufficiently by HPLC to permit quantifications.
55

RESULTS
Figure 16 Structures of pharmaceuticals and their ring-hydroxylated products. Chemical structures in
brackets refer to structures with uncertain positions of the introduced oxygen.
The observed products were also quantified, inso far as respective analytical standards were
available. The results of quantifications are summarized in table 11.
56

Table 11 Products identified by mass spectrometry after oxidation of selected pharmaceuticals by AaeAPO and CraAPO. The following data are given:
m/z values of the major observed diagnostic ion for substrates and products, consumed substrate (SC), yield (Y) and regioselectivity (RS).
Substrate Reaction Product
Discussed as HDM
AaeAPO (%) CraAPO (%)
m/z
in Ref. no.
SC Y RS SC Y RS
I 23 11
5-OH-Propranolol (Otey et al. 2006) [M+H] + 276 21 91 6 53
[M+H] + 260 4-OH-Propranolol (Otey et al. 2006) [M+H] + 276 − 2 13
N-desisopropylpropranolol (Otey et al. 2006) [M+H] + 218 traces 3 22
II 89 20
Acetaminophen (Greenberg and Lester 1946) [M-H] − 150 80 90 13 65
[M-H] − 134 3-OH-Acetaminophen (Greenberg and Lester 1946) [M-H] −
166 5 5 1 2
III 22 15
3-OH-Carbamazepine (Pearce et al. 2002) [M+H] + 253 13 61 6 40
[M+H] + 237 di-OH-Carbamazepine (Pearce et al. 2002) [M+H] + 269 n.d. n.d.
IV 78 15
[M+H] + 296 4'-OH-Diclofenac (Bort et al. 1999) [M+H] + 312 68 87 5 30
V 30 5
OH-Ketoprofen
[M+H] + 255
(Alkatheeri et al. 1999, Skordi
et al. 2005)
[M+H] + 271 n.d. n.d.
di-OH-Ketoprofen n.a. [M+H] + 287 n.d. n.d.
VI 94 13
[M+H] + 251 OH-Sulfadiazine (Winter and Unadkat 2005) [M+H] + 267 n.d. n.d.
VII 99 15
OH-Chlorpromazine (Perry et al. 1964) [M+H] + 335
[M+H] + 319 di-OH-Chlorpromazine n.a. [M+H] + 351 n.d
n.d.
N-desalkyl-OH-Chlorpromazine n.a. [M+H] + 250
(n.d.) = not determined due to poor resolution or no standard, (−) = not detected, (n.a.) = not applicable
RESULTS
57

RESULTS
3.1.2 Ring hydroxylation of drugs of abuse
Figure 17 provides an overview of controlled and illegal drugs that were ring-hydroxylated by
fungal peroxygenases in the presence of H 2 O 2 . The benzodiazepine drugs VIII-X were
hydroxylated by AaeAPO and CraAPO to the same products, though the yields of products
were lower in case of CraAPO. MroAPO oxidized only oxazepam (X).
Figure 17 Structures of controlled and illicit drugs and their hypothetical ring-hydroxylated products.
Chemical structures in brackets are uncertain with respect to the position of the hydroxyl group on the
aromatic ring where no analytical standard was available.
58

RESULTS
As already mentioned in material and methods section, ascorbic acid was added to the
reaction mixtures, to prevent the oxidative coupling and polymerization of phenolic products,
by scavenging the phenoxyl radicals produced via the peroxidative activity of AaeAPO.
Methamphetamine (XI), amphetamine (XII) and LSD (XIII) were also ring-hydroxylated, but
the yields were very low (

RESULTS
Figure 18 Structures of pharmaceuticals and their products that were oxidized at alkyl side chains.
Chemical structures in brackets are hypothetical with regard to the position of the introduced oxygen
(no analytical standard available).
Figure 19 shows the structures of the APO substrate Δ 9 -tetrahydrocannabinol (THC, XVIII)
and its products proposed on the basis of mass spectral and literature data (ElSohly and Feng
1998).
Figure 19 Structures of Δ 9 -tetrahydrocannabinol and its proposed products.
60

Table 13 Products identified by mass spectrometry after oxidation of the listed pharmaceuticals by AaeAPO and CraAPO.
The following data are given: m/z values for the major observed diagnostic ions for substrates and products, consumed substrate (SC), yield (Y) and regioselectivity (RS).
Substrate Reaction Product
Discussed as HDM
AaeAPO (%) CraAPO (%)
m/z
in Ref no.
SC Y RS SC Y RS
XIV 87 98
[M+H] + 207
2-OH-Ibuprofen
(Hamman et al. 1997, Holmes et al.
2007)
[M+H] + 223 21 24 74 75
1-OH-Ibuprofen (Holmes et al. 2007) [M+H] + 223 7 8 −
1-oxo-Ibuprofen n.a. [M+H] + 221 traces −
XV a 25 20
[M-H] − 271 4-OH-Tolbutamide (Hansen and Brøsen 1999) [M-H] − 287 15 60 13 62
4-keto-Tolbutamide n.a [M-H] − 285 n.q. n.q.
XVI 98 96
OH-Gemfibrozil (Murai et al. 2004) [M-H] − 265
[M-H] − 249 di-OH-Gemfibrozil (Murai et al. 2004) [M-H] − 281
keto-Gemfibrozil n.a. [M-H] − 263
n.q.
n.q.
carboxy-Gemfibrozil (Murai et al. 2004) [M-H] − 279
XVII 98 96
OH-Omeprazole (Hoffmann 1986) [M+H] + 362
n.q.
Omeprazole-sulfone (Andersson et al. 1993) [M+H] + 362
[M+H] + 346 keto-Omeprazole n.a. [M+H] + 362
O-desmethyl-OH-Omeprazole n.a. [M+H] + 348
n.q.
O-desmethyl-Omeprazole-sulfone n.a. [M+H] + 348
carboxy-Omeprazole (Andersson et al. 1993) [M+H] + 376
XVIII 99 99
[M+H] + 315
OH-THC (ElSohly and Feng 1998) [M+H] + 331
di-OH-THC (ElSohly and Feng 1998) [M+H] + 347
n.q. n.q.
a Mass spectral data indicate the formation of corresponding carbonyls as described previously (Kinne et al. 2010); (n.q.) = not quantified, no standard available; (−) = not
detected, (n.a.) = not applicable
RESULTS
61

RESULTS
Table 13 summarizes the m/z values of substrates and detected products, the amounts of
consumed substrate as well as the product yields and regioselectivity of reactions. The
conversion of all substrates into the products shown in Figures 18 and 19 (XIV-XVIII)
applies both to AaeAPO and CraAPO (though the product yields were different in some
cases); the quantitative results are summarized in the Table 13.
3.1 O-Dealkylation
1.3.1 O-Dealkylation of pharmaceuticals
AaeAPO and CraAPO cleaved alkyl aryl ether linkages in various drugs to yield phenolic
products (Kinne et al. 2009b). Thus, naproxen (XIX) was selectively demethylated to O-
desmethylnaproxen, dextromethorphan (XX) to dextrorphan, and metoprolol (XXII) to O-
desmethylmetoprolol.
Figure 20 Structures of O-dealkylated pharmaceuticals and their products. Chemical structures in
brackets are hypothetical and were deduced from mass spectral and literature data.
62

Table 14 Products identified by mass spectroscopy after oxidation of the listed pharmaceuticals by AaeAPO and CraAPO in the presence of H 2 O 2 . The m/z values for the major
observed diagnostic ion of substrates and products are given in each case along with the percentages of consumed substrate (SC), the yield (Y) and the regioselectivity (RS).
Substrate Reaction Product
Discussed as HDM
AaeAPO (%) CraAPO (%)
m/z
in Ref no.
SC Y RS SC Y RS
XIX a 60 10
[M-H] − 229 O-Desmethylnaproxen (Segre 1975) [M-H] − 215 57 95 9 85
XX 17 15
[M+H] + 272 Dextrorphan (Jacqz-Aigrain et al. 1993) [M+H] + 258 16 95 8 53
XXI 34 16
Acetaminophen (Brodie and Axelrod 1949) [M+H] + 152 23 66 13 80
[M+H] + 136
3-OH-Acetaminophen (Brodie and Axelrod 1949) [M+H] + 168 2 5 −
XXII 82 73
O-Desmethylmetoprolol (Hoffmann et al. 1987) [M+H] + 254 17 20 4 5
[M+H] + 268
α-OH-Metoprolol (Hoffmann et al. 1987) [M+H] + 284 2 2 1 1
XXIII b − 80
[M+H] + 313 Oseltamivir carboxylate (McClellan and Perry 2001) [M+H] + 285 − 71 88
RESULTS
a A syringe pump was used for hydrogen peroxide supply; (−) = not detected
63

RESULTS
Phenacetin (XXI) was deethylated to acetaminophen. Yields and regioselectivities were
generally greater with AaeAPO. The above phenolic reaction products all tended to
undergo further oxidation to quinones and/or coupling products because of the APOs’
peroxidase activity that generates phenoxyl radicals from phenols via one-electron
oxidations (Kinne et al. 2009a, Kinne et al. 2009b, Kinne et al. 2008, Ullrich and
Hofrichter 2007). Undesired coupling reactions were partially inhibitable by addition of the
radical scavenger ascorbate to the reactions. DNPH derivatization of the reaction mixtures
showed that the methyl group was released as formaldehyde in each case. The structures of
substrates and proposed products are summarized in the figure 20. In case of naproxen
(XIX) and phenacetin (XXI) O-dealkylation, AaeAPO and CraAPO showed significant
differences in the amount of converted substrate and the yield of products.
3.1.2 Ester cleavage
Ester cleavage by peroxygenase can be regarded as a special case of O-dealkylation.
CraAPO was capable of catalyzing this reaction. The enzyme selectively cleaved the
antiviral drug oseltamivir (XXIII) to oseltamivir carboxylate and acetaldehyde in high
yield. Interestingly, oseltamivir was not converted at all by AaeAPO.
3.2 N-dealkylation
3.2.1 N-dealkylation of pharmaceuticals
APOs catalyzed the oxidative N-dealkylation of several drugs that contained secondary or
tertiary amino groups. For example, AaeAPO regioselectively N-demethylated sildenafil
(XXIV) at the tertiary N of the piperazine ring to give N-desmethylsildenafil and
formaldehyde in high yield. In contrast to that, CraAPO was ineffective in catalyzing this
oxidation (figure 21). Among the other N-dealkylations showed in figure 21 are the
conversion of lidocaine (XXVI) to monoethylglycinexylidide and glycinexylidide, of 4-
dimethylaminoantipyrine (XXVI) to 4-aminoantipyrine and of tamoxifen (XXVII) to N-
desmethyltamoxifen.
64

Table 15 Products identified by mass spectrometry after oxidation of the listed pharmaceuticals by AaeAPO and CraAPO in the presence of H 2 O 2 . The m/z value for the major
observed diagnostic ion of substrates and products, consumed substrate (SC), the yield (Y) and the regioselectivity (RS) are shown in each case.
Discussed as HDM
AaeAPO (%) CraAPO (%)
Substrate Reaction Product
m/z
in Ref no.
SC Y RS SC Y RS
XXIV 82 80
[M+H] + 475 N-Desmethylsildenafil (Hyland et al. 2001) [M+H] + 461 82 99 4 5
XXV 60 45
Monoethylglycinexylidide (Orlando et al. 2004) [M+H] + 207 25 41 32 70
[M+H] + 235
Glycinexylidide (Orlando et al. 2004) [M+H] + 179 18 30 5 11
XXVI 68 38
[M+H] + 232 4-Aminoantipyrine (Banerjee et al. 1967) [M+H] + 204 16 23 19 48
XXVII 25 20
[M+H] + 372 N-Desmethyltamoxifen (Jin et al. 2005) [M+H] + 358
4-OH-Tamoxifen (Jin et al. 2005) [M+H] + 388
n.d. n.d.
Endoxifen (Jin et al. 2005) [M+H] + 374
(n.d.) = not determined due to poor resolution
RESULTS
65

RESULTS
Figure 21 Structures of pharmaceuticals that were subject to N-dealkylation. Chemical structures in
brackets are hypothetical.
Most of the products shown in Figure 21 could be quantified; the data are summarized in
Table 15.
3.2.2 N-dealkylation of drugs of abuse
N-Dealkylation was also observed for controlled and illicit drugs including some narcotics.
The structures of the tested substrates and proposed products are shown in Figure 22.
Codeine (XXVIII) and morphine (XXIX) were converted by MroAPO. The major
metabolite of codeine oxidation was the N-demethylation product norcodeine. Only traces
of morphine were formed, i.e. O-demethylation was much less pronounced than N-
demethylation; codeine was neither a substrate of AaeAPO nor of CraAPO. Unexpectedly
morphine was not subject to oxidation by MroAPO under the same conditions. Only in the
absence of ascorbic acid, a conversion of morphine was observed, which – according to
mass spectroscopic data - resulted in a dimerization of the molecule. This reaction might
be based on the formation of phenoxyl radicals and was also observed for AaeAPO and
CraAPO when ascorbate was omitted.
66

RESULTS
Figure 22 Structures of N-dealkylated controlled pharmaceuticals and illicit drugs and their
products. Chemical structures in brackets are hypothetical.
The reaction of APO with cocaine (XXX) was performed under slightly acidic conditions
(pH 5) to prevent the spontaneous hydrolysis of the substrate.
Table 16 Products identified by mass spectrometry after oxidation of the listed pharmaceuticals by
AaeAPO, CraAPO and MroAPO in the presence of H 2 O 2 . The m/z values of the major observed
diagnostic ion for substrates and products (P), and the amount of substrate consumed (SC) are
shown in each case; (n.d.) = not determined due to poor resolution.
Substrate
XXVIII
[M + H] + 300
XXIX
[M + H] + 286
XXX
[M + H] + 304
XXXI
[M + H] + 388
AaeAPO CraAPO MroAPO
SC P SC P SC P
(%) m/z (%) m/z (%) m/z
traces n.d traces n.d. 99
[M + H] + 286
[M + H] + 286
76
[M + H] + 569
[M + H] + 272
58
[M + H] + 569
[M + H] + 272
0 - traces [M + H] + 290 9
42
[M + H] + 360
[M + H] + 332
30
[M + H] + 360
[M + H] + 332
72
99
[M + H] + 569
[M + H] + 272
[M + H] + 290
[M + H] + 290
[M + H] + 360
[M + H] + 332
[M + H] + 374
67

RESULTS
Benzoylecgonine and norcocaine were found to be the major metabolites formed by
MroAPO and, in traces, by CraAPO. Flurazepam (XXXI) conversion was catalyzed by all
tested peroxygenases, and single and double N-dealkylated products were detected as
products. MroAPO turned out to be the best enzyme with a substrate conversion of almost
100%. The results of this study are summarized in the Table 16.
3.3 O-Demethylenation
AaeAPO was found to oxidize diverse benzodioxoles (XXXII-XXXVIII; table 17). The
reaction products were catechol derivatives and formic acid, which were identified by LC-
MS and IC, respectively.
Table 17 Structures of benzodioxoles and their products identified by mass spectrometry after
treatment with AaeAPO in the presence of H 2 O 2 . The m/z values of the major observed diagnostic
ion of the products, consumed substrate (SC), and product yield (Y) are given in each case.
Substrate
SC (%) Yield (%); mass ion (m/z)
Name
XXXII
4-Nitrobenzodioxole
XXXIII
4-Bromobenzodioxole
XXXIV
4-Chlorobenzodioxole
XXXV
Safrole
XXXVI
Myristicin
XXXVII
MDMA („ecstasy“)
O 2 N
Br
Cl
O
O
O
Structure
O
O
O
O
O
O
O
O
O
O
O
NH
96
99
99
78
99
88
Demethylenated
product
95
[M-H] - 154
13
[M-H] - 188
75
[M-H] - 143
n.d.
[M-H] - 149
n.d.
[M-H] - 179
n.d.
[M+H] + 182
formic
acid
95±5.0
68±2.1
94±3.2
75±7.0
98±1.1
83
O
XXXVIII
Pisatin
O
OH
90
n.d.
[M-H] - 301
50
O
O
68

RESULTS
Examples of oxidized benzodioxole derivatives are the entactogenic drug ecstasy
(XXXVII, MDMA, 3,4-methylenedioxymethamphetamine), the phytoalexin pisatin
(XXXVIII) from Pisum sativum as well as the nutmeg-seed ingredients safrole (XXXV)
and myristicin (XXXVI). Although no authentic standards of all reaction products were
available, the m/z values for these metabolites were the same as those expected for the
respective catechols with an m/z loss of 12 relative to the mass of the substrate (table 17).
Moreover, we used 18 O-labeled H 2 O 2 as co-oxidant and did not find an incorporation of
18 O into the catechol products. Our attempts to detect 18 O-labeled formic acid failed. The
reaction proceeded rapidly with total conversions between 78 and 99% of the substrate and
product yields up to 98%. For example, when we incubated 250 µM of 4-
nitrobenzodioxole with AaeAPO under standard reaction conditions and non-limiting
H 2 O 2 , the total conversion of the substrate was 240 µM (SC = 96 %) and we obtained
237.5 µM of 4-nitrocatechol and formaldehyde (Y=95%). Some of the products were
further oxidized; thus when 4-bromo- and 4-chlorobenzodioxole were converted to 4-
bromo- and 4-nitrocatechol, we noticed the formation of additional ring-hydroxylated
products indicated by an m/z of +16 in the mass spectrum (Appendix, Figure 8).
Formaldehyde was only released from pisatin and myrisiticin, and may originate from the
APO-catalyzed O-demethylation of the respective methoxy groups (Kinne et al. 2009b).
3.4 Oxidation of steroids
Initial attempts to convert steroids (17α-ethinyl estradiol and Reichstein’s substance S)
with AaeAPO and CraAPO did not give positive results (Poraj-Kobielska et al. 2011b).
Later these experiments were repeated with MroAPO and extended to other steroids.
MroAPO turned out to oxidize at least some steroids including important hormones,
anabolics and pharmaceuticals. In default of appropriate metabolite standards, the products
formed are given by their m/z values in Table 18.
69

RESULTS
Table 18 Conversion of steroids by MroAPO. Chemical structures of the substrates, the m/z values
for the major observed diagnostic ion of substrates and products, and the percentage of consumed
substrates are given.
70

RESULTS
3.5 Stoichiometrical analyses
3.5.1 Sildenafil
A quantitative analysis of AaeAPO-catalyzed sildenafil oxidation in the presence of
limiting H 2 O 2 revealed that one equivalent of N-desmethylsildenafil was formed per
equivalent of co-oxidant supplied (Table 19).
71

RESULTS
Table 19 Stoichiometry of sildenafil (XXIV) oxidation by AaeAPO. The initial sildenafil
concentration was 250 µM.
H 2 O 2 added
N-desmethylsildenafil
produced
Ratio
N-desmethylsildenafil/H 2 O 2
2.3
4.6
9.2
18.3
36.6
µM
2.12
4.27
8.75
20.49
32.37
0.92
0.93
0.96
1.12
0.88
3.5.2 Safrole
A quantitative analysis of safrole cleavage in the presence of a limiting amount of the cooxidant
H 2 O 2 gave an atypical result. Only half an equivalent of N-desmethylsildenafil was
formed per consumed equivalent of H 2 O 2 (table 20).
Table 20 Stoichiometry of safrole oxidation by A. aegerita peroxygenase. The initial safrole
concentration was 250 µM.
H 2 O 2 added Formic acid produced Formic acid/H 2 O 2
10.5
17.5
22.0
43.8
87.5
µM
5.70
9.02
11.53
21.68
45.53
0.54 ± 0.01
0.52 ± 0.01
0.52 ± 0.06
0.54 ± 0.06
0.52 ± 0.03
3.6 18 O-Isotope incorporation
Using AaeAPO and three pharmaceuticals (acetanilide, carbamazepine, tolbutamide), we
investigated the source of the oxygen introduced during hydroxylation. When the oxidation of
acetanilide (II) was conducted with H 18 2 O 2 in place of H 2 O 2 , mass spectral analysis of the
resulting acetaminophen showed that the principal [M-H] − ion had shifted from its natural
abundance m/z of 150 to m/z 152. Likewise, the hydroxylation of carbamazepine into 3-
hydroxycarbamazepine under these conditions resulted in a shift of the [M+H] + ion from its
natural abundance m/z of 253 to m/z 255. Accordingly, hydrogen peroxide was the source of
the introduced oxygen in both reactions.
A mass spectral experiment with tolbutamide and H 18 2 O 2 as the substrates showed that the
resulting 4-hydroxytolbutamide exhibited a principal [M-H] − ion with an m/z of 289 rather
than 287, again showing that the peroxide was the source of transferred oxygen (Figure 23).
72

RESULTS
Figure 23 HPLC elution profiles of the
conversion of tolbutamide (XV) in the
presence of ascorbic acid and AaeAPO
(background profile), control without
enzyme (front profile). Insets of mass spectra
show the incorporation of 18 O from H 2 18 O 2
into the alcohol group of 4-
hydroxytolbutamide (XVa). Upper inset:
MS of the product obtained with natural
abundance H 2 O 2 . Lower inset: MS of the
product obtained with 90 atom% H 2 18 O 2 . The
peak XVb conformed to the mass expected
for 4-ketotolbutamide, an oxidation product
of 4-hydroxytolbutamide (see Table 13)
(Kinne et al. 2010).
3.7 Kinetics experiments
3.7.1 Michaelis-Menten kinetics
Using AaeAPO, apparent kinetic constants were determined for some representative
pharmaceuticals. The apparent K m of the the enzyme for propranolol was 1,340 µM and the
k cat was 55 s -1 . For acetanilide, the constants were 1,310 µM and 1,925 s -1 . The apparent K m of
AaeAPO for phenacetin was 998 µM and the apparent k cat was 33 s -1 . For metoprolol, an
apparent K m of 2,330 µM and a k cat of 96 s -1 were detected. All kinetic constants of AaeAPO
determined here are summarized in Table 21 and compared with data obtained with other
substrates of AaeAPO in previous studies.
Table 21 Apparent kinetic constants of AaeAPO for selected pharmaceuticals in comparison to the
data for other substrates tested in previous studies (Kluge et al. 2007, Ullrich et al. 2004).
Substrate
Product
K m
(µM)
V max
[µM min -1 ]
k cat (s -1 )
k cat /K m
(M -1 s -1 )
ABTS (Ullrich et al. 2004) [ABTS] +· 37 283 7.7 × 10 6
Acetanilide Paracetamol 1,310 21077 1925 1.5 × 10 6
(Kluge et al.
Naphthalene
2007) 1-Naphthol 320 166 5.2 × 10 5
4-Nitro-benzodioxole 4-Nitrocatechol 1,190 457 3.8 × 10 5
73

RESULTS
Propranolol
Metoprolol
5-Hydroxypropranolol
Desmethylmetoprolol
1,340 3781 55 4.1 × 10 4
2,330 1047 96 4.1 × 10 4
(Ullrich et
Veratryl alcohol
al. 2004) Veratraldehyde 2,367 85 3.6 × 10 4
Phenacetin Paracetamol 999 361 33 3.3 × 10 4
3.7.2 Bisubstrate kinetics
The demethylenation of nitrobenzodioxole was chosen as a model reaction for initial rate
kinetics experiments at pH 7.0, in the course of which the concentration both of the substrate
and the co-substrate was varied. In contrast to the determination of the above apparent kinetic
data, steady state conditions can be assumed under these conditions. The calculation of kinetic
data was done using a nonlinear regression method and their double reciprocal plots gave
parallel lines (figure 24), which is consistent with an enzymatic ping-pong mechanism. The
kinetic parameters were as follows: k cat of 457 s -1 and a K m for H 2 O 2 of 2,160 µM and a K m for
4-nitrobenzodioxole of 1,190 µM (k cat /K m = 3.8×10 5 M -1 s -1 ).
Figure 24 Double reciprocal plots of the
kinetic data for nitrobenzodioxole oxidation
by AaeAPO. The H 2 O 2 concentrations used
were 62.5 µM (■), 75 µM (●) and 100 µM
(▲). The kinetic parameters reported in the
text were calculated by a nonlinear
regression method (Hernandez and Ruiz
1998)
74

RESULTS
3.8 Deuterium isotope effect experiments
Phenacetin (XXI) was found to be deethylated by AaeAPO to give acetaminophen and
acetaldehyde (table 14, figure 20). Since phenacetin has a symmetrical site at its α-carbon, it is
a suitable substrate to determine whether a catalyzed etherolytic reaction exhibits an
intramolecular deuterium isotope effect. HPLC-MS analysis of DNPH-derivatized reaction
mixtures showed that the AaeAPO-catalyzed cleavage of N-(4-[1- 2 H]ethoxyphenyl)acetamide
(phenacetin-d 1 ) resulted in a preponderance of [ 2 H]acetaldehyde 2,4-dinitrophenylhydrazone
(m/z 224, [M - H] - ) over natural abundance acetaldehyde 2,4-dinitrophenylhydrazone (m/z
223, [M - H] - ) (Figure 25). The observed mean intramolecular deuterium isotope effect
[(k H /k D )obs] from three experiments was 3.1 ± 0.2.
Figure 25 Oxidative cleavage of N-(4-[1-
2 H]ethoxyphenyl)acetamide (phenacetin-d 1 ) by
AaeAPO. The mass spectrum of the reaction mixture
containing the products acetaldehyde 2,4-
dinitrophenylhydrazone ([M - H] – , 123) and [ 2 H]
acetaldehyde 2,4-dinitrophenylhydrazone ([M - H] – ,
126) obtained from the oxidation of phenacetin-d 1 is
shown. The mass spectrum is one of three used to
calculate the observed mean intramolecular isotope
effect.
3.9 Spectrophotometric nitrobenzodioxole assay
Peroxygenases from A. aegerita, C. radians, Coprinopsis verticillata and M. rotula as well as
recombinant peroxygenase of Coprinopsis cinerea were dissolved together with
nitrobenzodioxole (NBD) in potassium phosphate buffer containing acetonitrile (10% vol/vol)
and, if the reaction products were to be quantified by HPLC, ascorbic acid. After adding the
co-substrate H 2 O 2 , the reaction mixture immediately changed its color to bright yellow (figure
26, inset A, Figure 27).
The metabolite causing color change was identified as 4-nitrocatechol by HPLC-MS relative
to an authentic standard with complete baseline resolution (compare also Table 17). The
reaction proceeded selectively with almost total conversions up to 99% of NBD. The
concentration-dependent change of absorbance in the reaction mixture (caused by the
formation of 4-nitrocatechol) at wavelengths in the visible region and at different stages of the
reaction (figure 26 A) was used to develop a direct spectrophotometric assay. A
spectrophotometer was applied to follow this rapid reaction over 15-30 seconds, which
corresponded with the linear initial slope (Figure 26 B). To prove the applicability of the new
enzyme assay, we followed the secretion of A. aegerita aromatic peroxygenase (AaeAPO) in
75

RESULTS
liquid cultures spectrophotometrically with the NBD assay and compared the results with the
classic peroxygenase assay 9 (Figure 27).
Figure 26 Concentration of 4-nitrocatechol in reactions followed by HPLC vs. absorbance at 425 nm
measured spectrophotometrically. Parallel reactions of the AaeAPO-catalyzed conversion of NBD
(0.125 mM) were stopped after different times of incubation (1: 5 s; 2: 15 s; 3: 30 s; 4: 60 s; 5: 120 s 6:
240 s; 7: 360 s) using sodium azide (1 mM). Insets show the time course of 4-nitrocatechol production
(A) and the spectral changes during the conversion (B).
It was found that the absorbance increase of the reaction mixture at 425 nm measured
spectrophotometrically correlated well (R 2 =0.99) with the concentration of produced 4-
nitrocatechol detected by HPLC in the range from 10 to 100 μM (Figure 26). Based on this
result and additional spectrophotometric measurements with NBD and 4-nitrocatechol
standards, we calculated a corrected extinction coefficient 10 of 9,700 M −1 cm −1 for 4-
nitrocatechol at 425 nm under the reaction conditions used.
9 1 U (unit) of peroxygenase represents the oxidation of 1 µmol of 3,4-dimethoxybenzyl
alcohol into 3,4-dimethoxybenzaldehyde in 1 min at 23 °C (Ullrich et al. 2004)
10 The actual extinction coefficient was calculated from calibration curves spectrophotometrically recorded for 4-
nitrocatechol at 425 nm (ε = 9,900 M -1 cm -1 ) and corrected by subtraction of the extinction coefficient of NBD at
425 nm (ε = 200 M -1 cm -1 ).
76

RESULTS
Figure 27 Parallel measurements of AaeAPO activity during liquid cultivation of the fungus A.
aegerita (days 9 to 23) in a soybean medium (2% w/vol) using the veratryl alcohol assay (I) (Ullrich et
al. 2004) and the new NBD assay (II). The inset shows the change of color before (A; pH = 7) and
after the addition of sodium hydroxide (B; pH >12).
The results show that the measured volume activity of the enzyme in the supernatant of the
culture liquid correlates well with the color change over a broad activity range. The activity of
A. aegerita peroxygenase measured with NBD was app. 80% higher than that with veratryl
alcohol (may be due to a higher turnover number of the enzyme for NBD). After alkalization
(pH >12) of the yellow reaction mixture with 10% (vol/vol) of 1 M sodium hydroxide, its
color changed to deep red (Figure 27, inset B). The corresponding UV-Vis spectrum of 4-
nitrocatechol showed an absorption maximum at 514 nm (ε 514 = 11,400 M −1 cm −1 , Appendix
Figure 9) (Cooper and Tulane 1936a). To confirm the specificity of the assay, we tested a
number of secreted oxidoreductases, such as horseradish peroxidase, soybean peroxidase,
chloroperoxidase from L. fumago and laccase from Trametes versicolor, none of which
showed any activity towards NBD. On the other hand, the aromatic peroxygenases from C.
radians, C. verticillata and M. rotula as well as the recombinant enzyme of C. cinerea
converted NBD into 4-nitrocatechol (Appendix Figure 7).
To prove the applicability of the NBD assay for environmental samples (at pH 7), we added
extracts (300 µL) from forest litter, wheat straw and beech wood to the assay mixture (0.7
mL) that contained a defined amount of AaeAPO (0.067 µM = 0.166 U mL -1 ) (see Chapter
2.3.3.3). Relative to the control assay, 87 ± 3% of added enzyme activity was detected in the
77

RESULTS
presence of the forest litter extract as well as 74 ± 5% and 55 ± 8% in the presence of the
straw and beech wood extracts, respectively.
3.10 Conversion of a set of human P450 substrates
A set of twelve common human P450 substrates (pharmaceuticals, xenobiotics, plant
ingredients), that is routinely used to distinguish between different CYP types (Walsky and
Obach 2004), was tested with our three model peroxygenases (AaeAPO, CraAPO, MroAPO).
The results are summarized in Table 22.
Coumarin (LVII; specific for CYP2A6) and felodipine (LV; CYP3A) were not or hardly
(

RESULTS
Table 22 Results of the treatment of twelve common substrates of human cytochrome P450s (CYPs)
by the fungal peroxygenases from A. aegerita, C. radians and M. rotula. P450/CYP products were
adopted from following references: (Walsky and Obach 2004). 1 (+) P450 metabolites were found,
(±) indication for the formation of P450 metabolites was found but standard was lacking, (-) no P450
metabolites were found; # rapid further conversion into the corresponding ketone product.
Substrate AaeAPO CraAPO MroAPO
LII
Conversion 6 15 16
Amodiaquine (%)
Cl N P450-
product
(CYP2C8)
desethylamodiaquine
desethylamodiaquine
Cl
N
H O
LIII
Bupropion
O
NH
NH
APO
Yield (%)
desethylamodiaquine
Conversion
(%)
P450-
product
(CYP2B6)
+
4
+
10
+
11
27 26 18
Cl
Cl
LIV
Chlorzoxazone
H
N
O
O
O
LV
Felodipine
Cl
Cl
O
N
H
LVI
Midazolam
N
N
N
F
O
O
APO ±
(3 hydroxylated
products)
Conversion
hydroxybupropion
hydroxybupropion
hydroxybupropion
(%)
P450-
product
(CYP2E1)
±
(2 hydroxylated
products)
58 38 8
6-hydroxy
chlorzoxazone
APO ±
(1 hydroxylated
product)
Conversion
(%)
P450-
product
(CYP3A)
6-hydroxy
chlorzoxazone
±
(1 hydroxylated
product)
-
6-hydroxy
chlorzoxazone
±
(1 hydroxylated
product)

RESULTS
Coumarin (%)
O O
P450-
product
(CYP2A6)
7-hydroxycoumarin
7-hydroxycoumarin
7-hydroxycoumarin
APO - - -
LVIII Conversion 15 no data 11
S-Mephenytoin (%)
H
P450- 4-hydroxy-Smephenytoimephenytoin
no data 4-hydroxy-S-
N O
product
N (CYP2C19)
APO + (traces) + (traces)
XXI Conversion 34 16 traces
Phenacetin
(%)
O
P450- acetaminophen acetaminophen acetaminophen
NH
product
(CYP1A2)
O
APO/
+
+
-
Yield(%) 23
13
0
IV
Conversion 78 15 68
Diclofenac
(%)
Cl
P450-
product
4’-hydroxydiclofenac
4’-hydroxydiclofenac
4’-hydroxydiclofenac
NH
(CYP2C9)
Cl
O
OH
APO/
Yield(%)
4-hydroxytolbutamide
XV
Tolbutamide
Conversion
(%)
P450-
product
(CYP2C9)
O HN
APO/
S NH O yield(%)
O
XX
Conversion
Dextromethorphan (%)
N H
P450-
product
(CYP2D6)
APO/
yield(%)
+
68
+
5
+
4
25 20 29
4-hydroxytolbutamide
4-hydroxytolbutamide
+
+
+
15 # 13 #

RESULTS
3.11 Difference spectroscopy
Heme containing enzymes can be characterized using a number of spectroscopic methods.
Thus it is known that the addition of various substrates or inhibitors can cause characteristic
changes in the optical absorption spectrum of P450 enzymes. The spectral changes reflect the
binding of these compounds near the active site, which causes changes in the heme
environment and influences the electron density (Lewis 2001).
Lewis described three characteristic types of substrate binding spectra for P450s, namely type
I (e.g. with phenol or propranolol), type II (e.g. with cyanide or pyridine), and the reverse type
I (sometimes referred as modified type II, e.g. with ethanol or caffeine) (Lewis 2001,
Schenkman et al. 1972, Schenkman et al. 1981).
⎯ Type I is characterized by a decrease in intensity of the Soret absorption peak around
418 nm, coupled with a concominant increase in the band intensity around 390 nm.
Consequently, a type I spectrum shows that the binding of substrate within the P450
heme locus brings about a shift in the iron(III) spin equilibrium from low-spin to highspin
(Lewis 1996).
⎯ Type II is typical for compounds acting as P450 inhibitors via ligation of the heme
iron (Lewis 1996). The UV spectrum is characterized by decrease in absorption around
390 nm with concominant increase around 430 nm. Consequently, there is a shift in
spin-state equilibrium from high-spin to low-spin iron.
⎯ Reverse type I is a "mirror-image" change to type I, where increase of absorption at
420 nm is coupled with a decrease in the 390 nm peak (Lewis 2001).
The results in Figure 28 and Table 23 show that the substrates propranolol, diclofenac and
dextromethorphan (Kinne et al. 2009a, Poraj-Kobielska et al. 2011b) as well as phenol
(Ullrich et al. 2004) caused a type I spectral change with the formation of maxima between
385 and 398 nm and minima around 420 nm.
Table 23 Difference absorption spectra of AaeAPO obtained by the addition of three common
pharmaceuticals and phenol as a reference.
Ligand λ max (nm) λ min (nm) Spectral type
Phenol 1 388 421 I
Dextromethorphan 389 422 I
Propranolol 386 422 I
Diclofenac 386 417 I
1 (Kinne 2010)
81

RESULTS
Figure 28 Difference absorption spectra of AaeAPO obtained by the addition of phenol (A),
dextromethorphan (B), propranolol (C) und diclofenac (D).
3.12 Different peroxides and peroxide-generating systems
The conversion of diclofenac into 4'-hydroxydiclofenac by AaeAPO was used to test the
suitability of different peroxides and peroxide-generation systems. The best result was
obtained with hydrogen peroxide (H 2 O 2 ) added in a few small doses, the lowest conversion
was observed with the system of L-cysteine. The organic peroxides mCPBA and tert-butyl
82

RESULTS
hydroperoxide were less efficient than H 2 O 2 but nevertheless enabled AaeAPO to catalyze the
substantial oxidation of diclofenac. The results of this study are summarized in Table 24.
Table 24 Comparison of different peroxides and peroxide-generating systems. The experiments were
performed with diclofenac (0.5 mM), AaeAPO (2 U mL -1 ) in the presence of ascorbic acid and 2
mM of each peroxide. Details are described in Chapter 2.10.
Peroxide/
peroxide generating system
Yield of 4'-OH-
Diclofenac (%)
H 2 O 2 (one time addition) 42
H 2 O 2 (few doses) 45
H 2 O 2 (syringe pump) 34
mCPBA 25
mCPBA (few doses) 32
mCPBA (syringe pump) 27
Glucose/glucose oxidase 36
L-cysteine 6
tert-butyl hydroperoxide 13
Copper(II) perchlorate hexahydrate/ascorbic acid 12
3.13 In vivo experiments
Growing liquid cultures of A. aegerita were supplemented with propranolol or diclofenac
(500 µM; 500 ppm); controls lacked either the pharmaceuticals or the fungus. Pictures of the
cultures are shown in Figure 29.
A
B
Figure 29 Liquid cultures of A. aegerita: on day 8 (A): soybean medium, one day after addition of
diclofenac (left) and control without diclofenac (right); (B): Czapek-Dox medium, one day after
addition of diclofenac (right) and without diclofenac (left).
Figure 30 summarizes the results of this in vivo experiment, in the course of which both
propranolol and diclofenac disappeared. This effect was more pronounced in soybean cultures
that produce much more peroxygenase than Czapek-Dox cultures do (Ullrich et al. 2004), but
where also adsorption processes may significantly contribute to the substrate removal.
83

84
RESULTS
Figure 31 HPLC elution profiles of samples from fungal cultures containing diclofenac (IV) or propranolol (I) in soybean medium. (A) Culture of A. aegerita with diclofenac on
day 14 when the pharmaceutical was added and on day 16, compared to a control without fungus; (B) Fungal culture with propranolol on day 7 (when substrate was added) and
the day after, compared to a control without fungus. (II) and (III) refer to medium components. The insets show the DAD-spectra of: (I) propranolol with structure; (II) and (III)
unidentified soybean medium components; (IV) diclofenac with structure.

RESULTS
Figure 30 Propranolol and diclofenac concentration in in-vivo experiments with liquid cultures of A.
aegerita: soybean medium (red), Czapek-Dox (green).
No indications for the formation of the typical peroxygenase metabolites were found.
However, both substrates disappeared rapidly in the fungal culures, particularly in those
containing soybean flour (where the enzyme activity was near its maximum around day 14;
Figure 30). Interestingly the detectable concentration of both substrates in the soybean
cultures decreased by more than 50% immediately after their addition (also in the controls
without fungus). This was probably the result of substrate absorption to the soybean particles
(figure 29 A). Figure 31 shows, as an example, the changes in the HPLC elution profiles
during the incubation of propranolol and diclofenac with A. aegerita in the soybean medium.
3.14 Tests with environmentally relevant concentrations of
propranolol
The concentration of pharmaceuticals in the environment (e.g. waste water, activated sludge)
is usually rather low (i.e. between 0.1-1000 ppb). Thus the testing of environmetally relevant
propranolol concentrations required the optimization of the analytical HPLC method and
sample preparation (2.5.2 and 2.12.). As a result, it was possible to identify and quantify
85

propranolol in the nM range (M W : 259 g mol -1 , 1 nM = 0.259 µg L -1 ). The outcome of this
experiment was that about 50% of the substrate propranolol at a concentration as low as 5 nM
(1.3 ppb) could be removed by AaeAPO (figure 32).
Figure 32 HPLC elution
profiles showing the
disappearance of
propranolol at an
environmentally relevant
concentration (5 nM = 1.3
ppb = 1.3 µg L -1 ) after
treatment with 50 units of
AaeAPO for 20 hours.
3.15 Long-term experiments
3.15.1 Pre-study with Vivaspin® (fed-batch method)
A Vivaspin® centrifugal concentrator (molecular mass cut-off 10 kDa) was used to determine
whether AaeAPO is stable enough to be repeatedly used in enzymatic conversions. Since the
concentrator holds back proteins, the enzyme, can in principle be used several times (Figure
33). Experiments with propranolol as the substrate demonstrated that this approach is feasible.
Surprisingly, the experiment without ascorbic acid gave more
product (5-OH-propranolol) than the experiment in its presence
(Figure 34), indicating – despite the reduction of phenoxyl radicals
that ascorbate provides – a general lowering of the overall
efficiency of the peroxygenase system by the radical scavenger.
A
B
Figure 33 Vivaspin tubes after treatment of propranolol with A. aegerita
peroxygenase (A) in the absence of ascorbic acid and (B) in its presence.
86

RESULTS
In the absence of ascorbic acid, the formation of dark substances was observed in the
Vivaspins®, which can be attributed to the oxidative coupling of phenoxyl radicals formed
from 5-OH-propranolol by the peroxidase activity of AaeAPO. The results of this experiment
are summarized in Figure 34.
Figure 34 Production of 5-OH-propranolol (green columns) and presence of remaining propranolol
(red columns) in Vivaspin experiments with AaeAPO. Reactions with (left) and without (right)
ascorbic acid.
3.15.2 Long-term experiment using a hollow fiber cartridge
Figure 35 Production of 5-OH-propranolol in a long-term experiment using AaeAPO. Daily amount of
propranolol added (green columns); cumulative amount of formed 5-OH-propranolol (orange
columns).
The preliminary experiments with Vivaspin® demonstrated that AaeAPO is highly stable and
can be used in long-term experiments. For that, the ultrafiltration membrane of a fiber
87

cartridge was loaded with AaeAPO protein and continuously used for propranolol
hydroxylation over 14 days. Figure 35 illustrates the progress of this experiment. In all, 13.85
mg of 5-OH-propranolol was produced from 217.56 mg of propranolol using 354 µg enzyme
over a period of fourteen days.
3.16 Scope of peroxygenase reactions concerning
pharmaceuticals and drugs
Figure 36 Substrates that were not converted by fungal peroxygenases.
88

RESULTS
In the course of this PhD project, some limitations of fungal peroxygenases concerning the
substrate spectrum have been ascertained. No conversion was observed for cocaine (XXX),
codeine (XXVIII), testosterone (XLI), 17α-ethinylestradiol (XLII), Reichstein’s substance S
(XLIII) and felbamate (LIX) when treated with AaeAPO or CraAPO. Furthermore, AaeAPO
was not able to convert oseltamivir (XXIII). The experimental results furthermore indicate
that APOs often fail to discriminate between chiral centers in the pharmaceutical substrates.
For example, chiral HPLC separation of the two O-desmethylnaproxen entantiomers that
resulted from naproxen oxidation showed that neither enantiomer predominated in the endproduct
mixture. In case of MroAPO, no product formation was observed for temazepam
(IX), oxazepam (X), diazepam (VIII) and amphetamine (XII). All three enzymes were unable
to convert felodipine (LV) and coumarine (LVII). The structures of the non-oxidizable
substrates are given in Figure 36.
89

DISCUSSION
4 Discussion
4.1 Preparation of human drug metabolites
Human drug metabolites (HDMs) are valuable chemicals needed for the development of
safe and effective pharmaceuticals. They are frequently used as substrates and authentic
standards in studies of drug bioavailability, pharmacodynamics and pharmacokinetics
(Baillie et al. 2002, Nelson 1982, Peters et al. 2007). In vivo, the C-H bonds of
pharmaceuticals are predominantly oxygenated by liver cytochrome P450-
monooxygenases (P450s) to yield more polar HDMs that are excreted directly or as
conjugates (Gibson and Skett 2001). This directed incorporation of an oxygen atom into a
complex organic structure is one of the most challenging reactions in synthetic organic
chemistry (Ullrich and Hofrichter 2007). Thus low yields and the need for laborious
removal of byproducts have prevented the cost-effective preparation of HDMs by purely
chemical methods (Kinne et al. 2009a). A preferable route to prepare human drug
metabolites would be the use of isolated human P450s, but these complex multiprotein
systems are membrane-bound, poorly stable, cofactor-dependent, and generally exhibit
low reaction rates (Eiben et al. 2006, Urlacher et al. 2004). More promising results have
been reported for human cell cultures (Acikgöz et al. 2009), microbial whole-cell systems
that heterologouly express human P450s (Peters et al. 2007) and laboratory-evolved
bacterial P450s (Damsten et al. 2008, Sawayama et al. 2009). The latter are capable of
catalyzing the H 2 O 2 -dependent hydroxylation of pharmaceuticals via the ”peroxide shunt”
pathway (Otey et al. 2006). Recent studies, however, have demonstrated that this approach
needs further optimization, in particular regarding turnover numbers and peroxide stability
(Sawayama et al. 2009). Alternatively, modified hemoproteins such as microperoxidases
might be used to catalyze hydroxylations by a P450-like oxygen transfer mechanism, but
so far these catalysts have not reached the necessary performance and selectivity (Caputi et
al. 2005, Dallacosta et al. 2003, Dorovska-Taran et al. 1998, Osman et al. 1996, Prieto et
al. 2006, Veeger 2002).
A few years ago, a new heme-thiolate enzyme was discovered in the wood- and litterdwelling
black poplar mushroom, Agrocybe aegerita. This enzyme turned out to be a true
peroxygenase (aromatic/unspecific peroxygenase, APO; EC 1.11.2.1), efficiently
transferring oxygen from peroxides to various organic substrates including aromatic,
heterocyclic and aliphatic compounds (Hofrichter et al. 2010, Ullrich and Hofrichter 2005,
90

DISCUSSION
Ullrich et al. 2004). In the following years, a few similar enzymes have been found in
cultures of other agaric basidiomycetes such as Coprinellus radians, Coprinopsis
verticilata (Anh et al. 2007) and Marasmius rotula (Gröbe et al. 2012) These stable,
secreted enzymes are promising oxidoreductases for biotechnological applications
including the synthesis of human drug metabolites. Depending on the particular substrate
and the reaction conditions, APOs catalyze in addition to peroxygenations, also oneelectron
oxidations (e.g. of phenols) and unspecific halogenations (bromination), and for
all these reactions, they need merely a peroxide as co-substrate to function.
Previous studies with diverse organic compounds showed that APOs catalyze several types
of oxyfunctionalization such as epoxidation, aromatic and benzylic hydroxylation, ether
cleavage (O-dealkylation), N-dealkylation, sulfoxidation and N-oxidation (Aranda et al.
2009a, Aranda et al. 2009b, Hofrichter et al. 2010, Kinne et al. 2009a, Kinne et al. 2009b,
Kinne et al. 2008, Kinne et al. 2010). All these reactions are also typical of P450s and part
of drug transformations in the human liver (Guengerich 1987, Ortiz de Montellano 2005).
Promising results were achieved in the prestudies with the pharmaceuticals propranolol
and diclofenac, which were converted regioselectively into the respective hydroxylated
human drug metabolites (Kinne et al. 2009a). All of this background has contributed to the
here developed approach: namely that aromatic peroxygenases (APOs) from the
basidiomycetes A.aegerita (AaeAPO), C. radians (CraAPO) and M. rotula (MroAPO) can
be used as an "artificial liver" to produce diverse HDMs.
To this end, a representative selection of widely used pharmaceuticals and illicit drugs was
tested with the above mentioned peroxygenases. In the first phase of this study, only
AaeAPO and CraAPO were available, later also MroAPO was included in the in vitro
experiments. The results have shown that all fungal peroxygenases (APOs) tested
catalyzed the hydroxylation or dealkylation of diverse pharmaceuticals and drugs of abuse,
in most cases with high regioselectivity. Moreover, MroAPO was also found to catalyze
the conversion of some steroids that were not accessible to the other APOs. The findings
of this study confirm previous work, which suggested that extracellular APOs share
functional characteristics with intracellular P450s but utilize the peroxide-dependent shunt
pathway with much higher efficiency (Guengerich 2001, Hofrichter and Ullrich 2010,
Ortiz de Montellano and de Voss 2005, Shoji et al. 2007). These peroxygenations are
thought to be initiated when the enzyme heme is oxidized by H 2 O 2 to give an iron species
(oxo-ferryl iron, compound I) that carries one of the peroxide oxygens, which
subsequently oxidizes a C-H bond in the substrate (Rittle and Green 2010).
91

DISCUSSION
Figure 37 Molecular structures of pharmaceuticals and drugs converted by AaeAPO. The substrates
are divided into three different groups according to the relative effectiveness of their oxidation.
Arrows indicate the points of enzymatic attack. SC – substrate conversion
Figures 37-39 give an overview of pharmaceutically relevant peroxygenase substrates
classified by the effectiveness of their conversion. What at first glance (Figure 37) attracts
the attention is the better (60-100%) conversion by AaeAPO of relatively small aromatic
substances or less complex molecules. However, more complex structures with exposed
aromatic rings, with secondary or tertiary amino groups or methoxy groups were
92

DISCUSSION
nevertheless oxidized by the enzyme to extents ranging from 20 to 59%. Amphetamines
were only hydroxylated at the aromatic ring and after long reaction times, and even then
the conversion rate was relatively low (below 15%). CraAPO has a similar substrate
spectrum as AaeAPO but the conversion rates were considerably lower in most cases
(Figure 38). On the other hand, compared to AaeAPO, an overview of the MroAPO
substrates suggests a preferance for more complex non-aromatic structures by the enzyme
as found, for example, with steroids or codeine. The latter substrates were hardly or not at
all oxidized by AaeAPO and CraAPO, which indicates considerable differences in the
heme channels and/or the active sites of the enzymes.
Furthermore, most of the observed oxidations matched those catalyzed by human liver
P450s, e.g. propranolol (I) to 5-hydroxypropranolol (CYP2D6) (Masubuchi et al. 1994),
tolbutamide (XV) to 4-hydroxytolbutamide (CYP2C9) (Miners and Birkett 1996),
naproxen (XIX) to O-desmethylnaproxen (CYP1A2) (Miners et al. 1996), acetanilide (II)
to acetaminophen (CYP1A2) (Liu et al. 1991, Miners et al. 1996) and sildenafil (XXIV) to
N-desmethylsildenafil (CYP3A4) (Ku et al. 2008). Particularly interesting results will be
discussed in detail in the next part of this chapter.
Evidently, APOs have some advantages over P450s - including currently developed,
laboratory-evolved P450s - where reaction yields are concerned (Kinne et al. 2009a). For
example, the transformation of diclofenac (IV) to 4-hydroxydiclofenac by mutants of
P450 cam (CYP101A1) resulted in total conversions between 15 and 44% and yields around
10% (Weis et al. 2009). By contrast, the wild-type AaeAPO used here gave a total
conversion of 78% and a yield of 68% for the same reaction. For the metabolism of
propranolol (II) in humans, CYP2D6 and CYP1A2 are responsible, forming three major
products: 5-hydroxypropranolol, 4’-hydroxypropranolol and desisopropylpropranolol
(Masubuchi et al. 1994). Positive results concerning the production of propranolol
metabolites were reported for laboratory-evolved bacterial P450s that use the shunt
pathway to hydroxylate the pharmaceutical. Thus, an engineered mutant D6H10 from
P450 BM3 (Bacillus megaterium, expressed in E.coli) was optimized for the synthesis of
propranolol metabolites, especially the 5-hydroxylated product. The optimization resulted
in a yield of 0.5% of 5- hydroxypropranolol (6.5 mg L -1 ) when 5 mM (1,295 mg L -1 )
propranolol was converted (Otey et al. 2006). Analogous experiments with AaeAPO gave
yields up to 13.6% (176 mg L -1 ), with regioselectivity higher than 90%. Interestingly,
CraAPO produced the other human P450 metabolites of propranolol, too. The analysis
93

DISCUSSION
showed the formation of the following metabolites in the stated yields: 6% of 5-
hydroxypropranolol, 2% of 4’-hydroxypropranolol and 3% of desisopropylpropranolol.
The conversion of ibuprofen (XIV) is another example for the formation of different
metabolite ratios by different peroxygenases. In humans, more than 70% of ibuprofen is
hydroxylated. The formation of metabolites is dependent on the P450 system and
comprises 2-hydroxy-, 3-hydroxy-, and 1-hydroxyibuprofen (Hamman et al. 1997, Holmes
et al. 2007). AaeAPO converted ibuprofen to 2-hydroxyibuprofen and 1-hydroxyibuprofen
with yields of 21% and 7%, respectively. The reaction with CraAPO showed high
regioselectivity and gave only 2-hydroxyibuprofen with a yield of 72%.
Not only hydroxylations but also N-dealkylations were differently catalyzed by different
peroxygenases. For example, the oxidation of lidocaine (XXV) catalyzed by AaeAPO gave
25% of monoethylglycinexylidide and 18% of glycinexylidide (ratio 1.4:1), whereas
CraAPO formed 32% of monoethylglycinexylidide (with a regioselectivity of 70%) but
only 5% of glycinexylidide (ratio 6.4:1). Both substances are major human metabolites
when lidocaine is used as local anesthetic (Orlando et al. 2004, Tam et al. 1990).
The different specificities of fungal peroxygenases are an interesting aspect, suggesting
that a set of them might be used as a "biocatalytic toolbox". Depending on what
reaction/metabolite is needed, the appropriate enzyme could be chosen and applied under
appropriate conditions. This interesting approach will be clarified by the following
examples.
The regioselective O-dealkylation of the anti-inflamatory drug naproxen (XIX) leads to the
formation of 6-desmethylnaproxen as the only metabolite in the human liver (Segre 1975).
In catalyzing this reaction, AaeAPO (yield of 57 %) was more effective than CraAPO (9%
yield). On the other hand, CraAPO was the only peroxygenase to catalyze the unusual
cleavage of an ester bond in the antiviral substance oseltamivir, which led to the formation
of the human metabolite oseltamivir carboxylate (yield 71%; formed in humans by hepatic
carboxylesterases; (Lindegardh et al. 2006, McClellan and Perry 2001), which is the
bioactive form of the drug (McClellan and Perry 2001). Sildenafil (XXIV) is an inhibitor
of the human cGMP-specific phosphodiesterase type 5 and widely used as an antiimpotence
drug (Ballard et al. 1998, Boolell et al. 1996). The compound was N-
dealkylated by all tested peroxygenases at the piperazine ring to form N-
desmethylsildenafil, the major metabolite circulating and operating in vivo. (Walker 1999).
The conversion rates were quite high in all cases (>80 %), but the yields of N-
desmethylsildenafil differed noticeably with only 4% for CraAPO and almost 90% for
94

DISCUSSION
AaeAPO and MroAPO. Other pathways of the metabolism of sildenafil lead in humans to
the formation of piperazine-N,N-deethylation and aliphatic hydroxylation products
(Muirhead et al. 2002, Walker 1999).
The benzodiazepine compounds diazepam (VIII), temazepam (IX) and oxazepam (X)
used as psychoactive drugs in the treatment of mental deseases were efficiently
hydroxylated (m/z +16) by AaeAPO.
Figure 38 Molecular structures of pharmaceuticals and drugs converted by CraAPO. The
substrates are divided into three different groups according to the relative effectiveness of their
oxidation. Arrows indicate the points of enzymatic attack. SC – substrate conversion
95

DISCUSSION
The reaction of diazepam with human P450s gives also a hydroxylated derivative as the
major metabolite (temazepam, hydroxylated at the heterocyclic 7-ring) and N-
demethylated diazepam (N-desmethyldiazepam) as a second metabolite (Andersson et al.
1994). However, since the hydroxylated product of the AaeAPO reaction did not have the
same retention time as temazepam, and taking into account the preference of AaeAPO for
aromatic substrates (Aranda et al. 2009a, Aranda et al. 2009b), the hydroxylation of one of
the carbons of the phenyl moiety would be a plausible explanation (most probably in parapostion).
Temazepam and oxazepam were seemingly hydroxylated by AaeAPO in a
similar way, and in the case of the latter compound, also a double hydroxylated product
was detected (m/z +32). By contrast, P450s preferably N-demethylate temazepam to form
oxazepam (Schwarz 1979), which in turn is almost exclusively conjugated with glucuronic
acid to improve excretion (Kellermann and Luyten-Kellermann 1979).
The antitussive morphine-derivative codeine (XXVIII), which was not a substrate of
AaeAPO or CraAPO, was rapidly converted by MroAPO into the N-dealkylated product
norcodeine and traces of the O-dealkylation product morphine (XXIX). Unexpectedly, no
product was found when morphine was treated with MroAPO under standard conditions;
only in the absence of ascorbic acid, a reaction was observed leading to the formation of a
coupling product (dimer, m/z 569) that was later also observed for AaeAPO and CraAPO.
This fact indicates that peroxygenases attack the phenolic group of morphine via oneelectron
oxidation resulting in the formation of a phenoxyl radical that in turn can dimerize
(Figure 22, Figure 50). Since the morphine molecule and its coupling product are relatively
bulky, the oxidation may occur at the enzyme surface rather than in the heme channel.
P450s (e.g. human CYP2D6) preferably O-demethylate codeine to form morphine in Phase
I metabolism, and this product is in turn conjugated with glucuronic acids and excreted
(Kilpatrick and Smith 2005, Williams et al. 2002). Tests with human plasma samples
showed that codeine is primarily conjugated at the 6-position. The P450-catalyzed
demethylation clearance rates of codeine and norcodeine were found to be similar in
magnitude but about four times lower compared to 6-glucuronidation. Since the partial
clearance to norcodeine was normal in poor human metabolisers, it would be reasonable to
infer that the two demethylation processes (N-demethylation to norcodeine, O-
demethylation to morphine) are mediated by different cytochrome P-450 isoenzymes
(Chen et al. 1991). In contrast to morphine, the psychoactive, illicit drug
tetrahydrocannabinol (THC, XVIII) was efficiently converted by AaeAPO and CraAPO
(conversion rate almost 100%) resulting in the formation of two hydroxylated products, the
96

DISCUSSION
major of which is probably 11-hydroxy-THC; proposed structures are shown in the results
section (Figure 19).
Figure 39 Molecular structures of pharmaceuticals and drugs converted by MroAPO. The
substrates are divided into three different groups according to the relative effectiveness of their
oxidation. Arrows indicate the points of enzymatic attack. SC – substrate conversion.
In other experiments where side chains were attacked the best results were noted for
MroAPO but in the case of THC, no positive result was observed. In the human liver, THC
is metabolized mainly to 11-hydroxy-THC, which is further oxidized to the corresponding
carboxylic acid (11-nor-9-carboxy-THC); both reactions are thought to be catalyzed by
P450s (CYP2C9, CYP2C19, CYP3A4) (Bland et al. 2005, Lemberger 1973, Watanabe et
al. 2007).
97

DISCUSSION
None of the steroids tested were attacked by CraAPO and AaeAPO; however, MroAPO
again was able to convert these bulky molecules. According to the effectiveness of the
conversion, two groups of steroid substrates can be distinguished. Substrates of the first
group were converted by more than 95% (e.g. Reichstein's substance S, cortisone,
prednisone, medrol), while those of the second group were oxidized with lower rates (2-
70% conversion; e.g. progesterone, testosterone, nandrolone, pregnenolone). All steroids
of the first group bear a 1-oxo-2-hydroxyethyl side chain (-CO-CH 2 -OH), whereas those of
the second group have a 1-oxoethyl (-CO-CH 3 ) or shorter (e.g.
-CH 3 ) side chain, or no side chain at all (e.g. -OH). This finding indicates that the primary
alcohol functionality of the oxohydroxyethyl group somehow facilitates the conversion of
steroids by MroAPO (and maybe it is also the target of oxidation). Unfortunately, the mass
spectral data of the products are difficult to interpret, so that incorporation of oxygen (m/z
+16) could be deduced in only a few cases, an (e.g. for Reichstein's substance S,
pregnenolone, progesterone; see also Table 18 in the results section). Nevertheless the
mass data indicate another interesting reaction that led to the formation of a progesteronelike
product (m/z 315) from pregnenolone (m/z 317). The reaction could proceed via a
second hydroxylation of the C3-position yielding a geminal alcohol that eliminates water
to form a keto group. However, it is unclear whether and if so how the double bound is
transferred from C5 to C4. In animals and humans these reactions are catalyzed by the 3-βhydroxysteroid
dehydrogenase/Δ-4-5 isomerase (EC 1.1.1.145), the only enzyme in the
adrenal pathway of steroid synthesis that is not a member of the P450 superfamily
(Thomas et al. 2004). Various other human P450s are involved in steroid transformation
and the synthesis of hormones; among others, they hydroxylate different rings (e.g.
CYP11B1, CYP17), the side chain (e.g. CYP21) and the C18-methyl group (CYP11B2),
and as well they are involved in the aromatization of the A ring, thereby converting
androgens to estrogens (CYP19) (Hall 1986).
The results of steroid conversion clearly display some limitations and drawbacks of APO
catalysis. One is that the enzymes have relatively narrow catalytic clefts (albeit the hemechannel
is considerably wider than those of other heme peroxidases), which prevent access
of markedly bulky substrates to the active site (Piontek et al. 2010). The negative results
we obtained when treating felbamate (LIX), 17α-ethinylestradiol (XLII), testosterone
(XLI), Reichstein's substance S (XLIII) and other steroid structures with AaeAPO or
CraAPO may reflect this limitation. Furthermore, all three tested peroxygenases were
98

DISCUSSION
unable to mimic some characteristic reactions of human P450s such as felodipine (LIX)
and coumarin (LVII) oxidation. Why the small and hydrophobic coumarin molecule was
not attacked remains an enigma. Although MroAPO had no problems with the conversion
of steroids, it was not able to catalyze the hydroxylation of benzodiazepines, which on the
other hand were good substrates of AaeAPO and CraAPO.
In conclusion, fungal APOs may be better suited than most P450s for synthetic
applications: (i) they utilize a relatively cheap co-substrate, H 2 O 2 ; (ii) they do not require
costly co-reactants such as pyridine nucleotides, flavin reductases, or ferredoxins; (iii) they
are secreted enzymes and thus can be cost-effectively produced (no cell disruption
required); and (iv) they are stable and soluble due to their high degree of glycosylation
(Pecyna et al. 2009, Ullrich et al. 2009).
4.2 Mechanistic studies
4.2.1 Source of oxygen and stoichiometry.
The results of this work and the previous studies with APOs showed that an oxygen atom
from hydrogen peroxide is incorporated into the phenolic or alcoholic functionalities
produced after conversion of alkyl aromatics (Aranda et al. 2009a, Kinne et al. 2009a,
Kinne et al. 2008, Kluge et al. 2009), and also into the carbonyls produced after oxidation
of alcohols or oxidative cleavage of ethers (Kinne et al. 2009b). According to the above
results, oxygen incorporation from H 2 O 2 should be quantitative when the substrate is
oxidized. The data on APO-catalyzed oxidation of pharmaceuticals agree with this picture,
since almost 100% of oxygen present in newly generated phenolic or benzylichydroxylation
products - e.g. in the phenolic group of acetaminophen or in the alcohol
moiety of 4-hydroxytolbutamide - was 18 O-labeled when the experiment was conducted
with H 18 2 O 2 . This result correlated well with the reaction mechanism reported for
engineered P450 BM3 or P450 cam , which can use the so-called “peroxide shunt” pathway,
in the course of which also one oxygen atom from hydrogen peroxide is incorporated into
the substrate to yield an oxygenated product without the need of a reductase domain,
dioxygen or NADPH; (Joo et al. 1999, Otey et al. 2006). Moreover, the stoichiometrical
experiment with sildenafil showed that one equivalent of N-desmethylsildenafil was
formed per equivalent of H 2 O 2 supplied. This agrees with the two-electron oxidation
expected from a peroxygenative mechanism (Section 1.2.7; (Gorsky et al. 1984)). The
99

DISCUSSION
stoichiometrical experiment with benzodioxoles seems to follow another mechanism and
that is described in one of the following subchapters (4.2.7).
4.2.2 Deuterium isotope effect
Historically, the determination of deuterium isotope effects has been a powerful tool to
help unravel the intricacies of carbon-hydrogen bond cleavage and define the mechanism
of specific chemical reactions. The intrinsic primary isotope effect, k H /k D , is the magnitude
of the isotope effect for a reaction on the rate constant for the bond-breaking step (C–H
versus C–D) of the reaction and is related to the symmetry of the transition state for that
step (Nelson and Trager 2003). There are two designs to follow deuterium isotope effects.
Intermolecular design depends on complex determinations of rate constants, whereas
intramolecular design is kinetically independent of all steps besides the C-H bond-breaking
step, and is thereby simpler. In an isotope effect experiment of intramolecular design, a
substrate that is susceptible to enzymatic attack at either of two symmetrically equivalent
sites is chosen (Hjelmeland et al. 1977, Nelson and Trager 2003). One site contains
deuterium and the other retains its normal complement of hydrogen. The enzyme then has
the choice of attacking a deuterated or an equivalent protio site within the same molecule.
The observed isotope effect (k H /k D ) obs is measured as the ratio of product resulting from
attack at the protio site versus product resulting from the attack at the deutero site (Kinne
2010, Nelson and Trager 2003). The intramolecular deuterium isotope effect that was
observed for phenacetin-d1 oxidation by AaeAPO, is consistent with a P450-like
mechanism. The determined value of (k H /k D ) obs around 3 is close to the values of (k H /k D ) obs
near 2 that have been observed for the P450-catalyzed O-dealkylation of this substrate
(Garland et al. 1976). A value of 2 is considerably smaller than the intrinsic isotope effect
near 10 expected for a hydrogen abstraction mechanism, and may indicate that phenacetin
oxidation by APOs may proceed instead via electron transfers, as proposed earlier for the
P450-catalyzed reaction (Yun et al. 2000). By contrast, the O-dealkylation of 1,4-
dimethoxybenzene-d 3 by AaeAPO probably does proceed via hydrogen abstraction,
because it was found to exhibit a much higher (k H /k D ) obs near 12 (Groves and McClusky
1978, Kinne et al. 2009b, Yun et al. 2000).
4.2.3 Kinetics
The apparent kinetic data determined in this study for the action of AaeAPO on a variety
of pharmaceuticals suggest that peroxygenases may be a useful alternative to P450s for the
100

DISCUSSION
regioselective preparation of human drug metabolites (HDMs). Although some P450s are
known to bind pharmaceutical substrates more strongly than APOs, exhibiting K m values
between 1 and 70 µM, they generally exhibit relatively low k cat values in the vicinity of 0.2
s -1 or less (Upthagrove and Nelson 2001, Yun et al. 2000). As a result, the catalytic
efficiencies [k cat /K m ] of AaeAPO for pharmaceutical oxidations - despite relatively high K m
values - lie in the same range as those of the P450s, as exemplified by our values for
propranolol hydroxylation (1.32 × 10 5 M -1 s -1 ), metoprolol O-dealkylation (4.1 × 10 4 M -1 s -
1 ) and phenacetin O-dealkylation (3.3 × 10 4 M -1 s -1 ). Moreover, the k cat /K m value we
obtained for acetanilide (1.5 × 10 6 M -1 s -1 ) is much higher than the corresponding value
found for the P450-catalyzed reaction (Liu et al. 1991).
4.2.3.1 Bisubstrate kinetics
In some cases, the kinetic investigation of APO-catalyzed reactions has encountered some
experimental problems. For example, a previous study showed that, under saturating H 2 O 2
concentrations, AaeAPO exhibits an interfering catalase activity, which can lead to an
underestimation of kinetic efficiency (Ullrich et al. 2008). A better option for the
evaluation of kinetic parameters would be bisubstrate systems using an experimental
design that varies the concentration of both substrates (Segel 1994). Initial velocity
patterns are usually obtained by making reciprocal plots for one substrate (variable
substrate) at different fixed concentrations of one of the others (changing fixed substrate),
while keeping all other substrates, if there are any, at constant concentration. If the
mechanism does not include alternate reaction sequences and no substrates react with more
than one enzyme form, the intercepts always have to be a linear function of the reciprocal
concentration of the changing fixed substrate. As a result, there are two possible initial
velocity patterns: parallel lines when no reversible connection exists, or lines intersecting
to the left of the vertical axis when such a connection does exist (Cleland 1963). Data from
bisubstrate kinetics under steady state conditions, which were spectrophotometrically
recorded for the AaeAPO-catalyzed reactions at varying concentrations of 4-
nitrobenzodioxole and H 2 O 2, gave in fact parallel lines for double reciprocal plots of the
data. This result is consistent with an enzymatic ping-pong mechanism (Bisswanger 2000,
Cleland 1963, Segel 1994). The mechanism can be explained as follows. The substrate (A)
binds to the free enzyme (E) to form the enzyme-substrate complex (EA). Substrate (A)
converts the enzyme to the modified enzyme-product complex (FP). Only after the
substrate (A) is released in the form of (P) can the second substrate (B) bind to the
101

DISCUSSION
modified enzyme (F), yielding the enzyme-substrate complex (FB), and then react with the
modified enzyme, finally regenerating the free enzyme (E) with release of the product (Q)
(Bisswanger 2000, Kinne 2010, Segel 1994).
Figure 40 Reaction sequence of a "ping-pong bi bi" mechanism. The enzyme constantly bounces
back and forth between the two states of the enzyme (E) and (F); substrate (A), second substrate
(co-substrate) (B), product (Q), enzyme-substrate complex (EA), complex of the modified enzyme
and the second substrate (FB), modified enzyme-product complex (FP), modified enzyme (F)
The kinetic parameters determined in this way for the demethylenation of the model 4-
nitrobenzodioxole by AaeAPO were in the same range as the data obtained for selected
pharmaceuticals. Thus, the K m for 4-nitrobenzodioxole of 1,190 µM is between the value
calculated for the N-demethylation of phenacetin (XXI; K m =999) and the hydroxylation of
acetanilide (II; K m = 1,310 µM). The K m for the demethylenation of methylenedioxymethamphetamine
(XXXVII) catalyzed by P450s is by two orders of magnitude lower
(K m =1.7-3 µM) (Tucker et al. 1994). The catalytic efficiency for nitrobenzodioxole O-
demethylenation (k cat /K m = 3.81×10 5 M -1 s -1 ), lies in the same range as that for
naphthalene oxidation (k cat /K m = 5,2 × 10 5 M -1 s -1 ; see Chapter 4.2.3 and 3.9.1).
4.2.4 Assays for human cytochrome P450 activities
Twelve validated assays for human hepatic cytochrome P450s have been established in the
last decades. Using this set of assays and substrates, six subfamilies, including nine
individual P450s can be characterized (Anzenbacher and Anzenbacherová 2001, Walsky
and Obach 2004, Wrighton and Stevens 1992). All twelve assays were applied here for the
identification of specific P450-like activities of three fungal peroxygenases (AaeAPO,
CraAPO and MroAPO; see Table 22, Chapter 3.12).
In the liver, CYP1A2 metabolizes about 15% of all incoming pharmaceuticals (Gibson and
Skett 2008). The activity of this enzyme is measured with the phenacetin (XXI) O-
deethylase assay; only MroAPO was unable to perform this reaction. Two additional
substrates of CYP1A2 and AaeAPO/CraAPO are naproxen (XIX) and 3,4-
aminoantipyrine (XXVI) (Anzenbacher and Anzenbacherová 2001). CYP2A6 contributes
102

DISCUSSION
about 5% of cytochrome P450-dependent drug metabolism in the liver. One of its
characteristic catalytic properties, a coumarin 7-hydroxylase activity, is seemingly lacking
in APOs. Other substrates of CYP2A6 are biogenic amines including tobacco-specific
nitrosamines with pro-carcinogenic potential (Guengerich and Montellano 2005).
However, CYP2A6 is generally considered less important in drug metabolism than other
P450s (Walsky and Obach 2004). CYP2B6 is responsible for only about 1% of hepatic
metabolism of pharmaceuticals. This enzyme is characterized by its bupropione
hydroxylase activity. Although AaeAPO and CraAPO converted bupropione to
hydroxylated products, the absence of an analytical standard did not allow a clear
conclusion as to whether the same carbon is attacked. Another marker reaction of CYP2B6
is the N-demethylation of S-mephenytoin, which was performed by the APOs only to a
low extent. CYP2C8 converts less than 5% of incoming pharmaceuticals in the liver and
has a generally low catalytic activity towards the known substrates of the CYP2C
subfamily (Guengerich and Montellano 2005); its characteristic amodiaquine N-
deethylase activity was detected for all APOs as well. CYP2C9 is involved in the
metabolism of about 20% of pharmaceuticals and drugs, and is one of the major enzymatic
metabolizers in the liver (Guengerich and Montellano 2005). It is characterized by
hydroxylating activities towards diclofenac and tolbutamide, both excellent substrates of
AaeAPO and also convertable by the other APOs (but with lower yields). Additional
substrates are carbamazepine (III) sildenafil (XXIV) (Warrington et al. 2000) and
ibuprofen (XIV). S-mephenytoin 4’-hydroxylation is the activity characteristic for
CYP2C19, which participates in less than 5% of hepatic drug metabolism. APOs
seemingly share this activity with CYP2C19, and also the ability to efficiently oxidize
diazepam (VIII), omeprazole (XVII) and propranolol (I). The CYP2E subfamily is present
in all mammals in the form of the characteristic representative CYP2E1. This enzyme is
well-known for its involvement in the metabolism of ethanol (Quertemont 2004, Zimatkin
and Deitrich 1997). Concerning pharmaceutical drugs, it was shown to convert
acetaminophen, phenacetin (XXI) and chlorzoxazone (LIV), the latter of which is the
specific assay substrate for CYP2E1 (Anzenbacher and Anzenbacherová 2001).
Chlorzoxazone (LIV) was found to be a good substrate of AaeAPO and CraAPO but, in
the absence of an appropriate analytical standard, it was not possible to confirm that the
same product was formed. The CYP3A subfamily with four human members contributes
to about 50% of the hepatic metabolism of drugs and pharmaceuticals (Anzenbacher and
Anzenbacherová 2001, Guengerich and Montellano 2005). The substrate spectrum of these
103

DISCUSSION
enzymes is very broad and includes, inter alia, various statins, steroids, and codeine
(XXVIII), but also APO substrates such as sildenafil (XXIV), lidocaine (XXV),
acetaminophen, carbamazepine (III), dextromethorphan (XX, also converted by
CYP2D6), diazepam (VIII), omeprazole (XXII), and tamoxifen (XXVII). The typical
CYP3A substrates felodipine and midazolam (both belonging to the twelve key assay
substrates) were not at all or only slowly converted by the APOs. Another CYP3A
substrate, testosterone, was only oxidized by MroAPO.
The results of this work clearly show that APOs share catalytic activities with all relevant
human P450s. Since the substrate spectra of P450s overlap with each other, it is not
possible to select a particular P450 enzyme that shows the most similarities with APOs at
the reaction level. Some limitations of APO catalysis and differences with human P450s
were also acertained (coumarin and felodipine were converted by none of the APOs,
testosterone only to some extent by MroAPO), and may be attributable to issues of
molecule size or charge, and/or special features of the 3-dimensional structure of the
substances (see 4.5 for more details). Table 38 summarizes the results of this study.
Table 38 Summary of the measured activities by P450s and APOs; +(red)- assay metabolite was
found; (+)(orange)- metabolite with the same ion mass was found, but analitycal standard was
lacking; *- N-dealkylation; **- hydroxylation
4.2.5 Peroxides and peroxide generation systems
In the literature, there are many ways described to add or generate peroxides to initiate
peroxidation/peroxygenation reactions. As a part of this work, three different hydrogen
peroxide generating systems and three peroxides were tested in the AaeAPO-catalyzed
conversion of diclofenac to 4'-hydroxydiclofenac.
One approach that goes back to studies in the 1950s uses the the oxidation of ascorbic acid
by dioxygen in the presence of Cu(II) perchlorate or L-cysteine, which generates
104

DISCUSSION
dehydroascorbic acid and hydrogen peroxide (Davies 1984, Khan and Martell 1967,
Udenfriend et al. 1954). Though the details of H 2 O 2 formation are not fully clear, two
possible mechanisms may be considered for metal-catalyzed peroxide formation via
ascorbyl radical intermediates. In the first proposed mechanism a metal-ascorbate complex
is suggested in a pre-equilibrium step. This is followed by a rate-determining electron
transfer within the complex from the ascorbate anion to the metal ion. The reduced
complex dissociates in a fast step to the lower valence metal ion and a semiquinone.
Afterwards follows reoxidation of the reduced metal ion by O 2 to produce superoxide,
which then generates H 2 O 2 via dismutation.
•
2
fast
2HO
⎯⎯→H
O + O
2
2
2
In the second suggested mechanism, at first a metal-ascorbate complex is formed. This is
followed by an attack of dioxygen on the metal-ascorbate complex with the formation of
an oxygen complex in a second pre-equilibrium step (Khan and Martell 1967).
AscH
M
−
( n−1)
+
+ O ⎯⎯→ Asc + HO
+ fast
+ HO H ⎯⎯→M
fast
2
•
2
+
•
2
n+
+ H
In the case of the cysteine system, it is thought that the initial oxidation of the thiolate
group of the amino acid into a thiyl radical initiates peroxide generation (Law et al.
1983) 11 . Another peroxide-generating system, which has widely been used in enzymatic
conversions for decades and was also successfully tested here, is that of glucose/glucose
oxidase (Hofrichter et al. 1998, Paice et al. 1993, Ullrich et al. 2004). tert-
Butylhydroperoxide and meta-chloroperoxybenzoic acid (mCPBA) were tested as organic
alternatives replacing hydrogen peroxide in this work. The former peroxide was shown to
be the best oxidant in sulfur/sulfide and olefin oxidation by chloroperoxidase and
horseradish peroxidase (Colonna et al. 1990, Hu and Hager 1999). Also mCPBA was used
as an oxidant in many previous studies, particularly when the oxidations with H 2 O 2 were
too fast (Kellner et al. 2002, MacKeith et al. 1994).
Recent studies with hydrogen peroxide have already shown that the dosage and the way of
adding it can considerably influence the efficiency of a APO-catalyzed reaction (Aranda et
al. 2009b, Poraj-Kobielska et al. 2011b). The results obtained here demonstrated that
hydrogen peroxide, mCPBA, and the system glucose/glucose oxidase are the best oxidants
2
O
2
11 Instead glutathione described in this paper, cysteine was used (Scheibner and Hofrichter 1998).
105

DISCUSSION
in terms of the diclofenac conversion rates (32-45%). A stepwise dosage of the oxidant
(with a pipette or a syringe pump) further increased product formation (by 3-7%) but was
not nessesarily required. The worst results were observed for the L-cysteine/ascorbate
system followed by the Cu(II) chlorate/ascorbate system and the usage of tertbutylhydroperoxide
(conversion rates for diclofenac 6-13%). This experiment showed that
different peroxides and sources of them can be used in APO-catalyzed reactions; the
effectiveness depends on the particular peroxide, its dosage and the reaction time.
4.2.6 Difference binding spectra
One of the characteristics of P450s is the influence of substrate binding on the appearance
of the overall spectrum, especially with respect to the positions and intensities of the major
absorption bands (Lewis 2001, Schenkman et al. 1981). AaeAPO shows difference binding
spectra that are similar to those of P450s. The pharmaceuticals tested with AaeAPO in this
respect were: propranolol (II), diclofenac (IV) and dextromethorphan (XX), as well as
phenol (C 6 H 5 -OH) as a reference (Kinne 2010, Lewis 2001). They all gave a binding type I
spectrum that is also characteristic for the binding exhibited by phenol (Hayhurst et al.
2001, Schenkman et al. 1981, Topham 1970, Wester et al. 2003). These results suggest
that the electronic environment of APOs, especially of AaeAPO, is similar to that of
P450s, although the amino acid residues in the heme channel and near the heme may be
totally different.
4.2.7 Phenoxyl radical scavenger
The high general ("classic") peroxidase activity of APOs can cause problems when the
desired products are phenols (e.g. after aromatic hydroxylation or O-demethylation of
methoxyaromatics), and necessitates the inclusion of a radical scavenger such as ascorbate
in the reactions.The peroxidase activity of APOs causes the formation of phenoxyl radicals
from phenolic compounds, which leads to undesired coupling reactions. Ascorbic acid is
present in an aqueous environment in a protonated form (at pH 7) as a monoanionic
species. The latter (probably as metal ion-ascorbate anion complex, see 1.2.5) can react
with a phenoxyl radical to give the ascorbyl radical that in turn may disproportionate to
dehydroascorbic acid and ascorbic acid. In other words, the phenoxyl radical
(R-C 6 H 5 -O • ) abstracts one electron from ascorbate followed by hydrogen/proton rebound
to reform the phenolic compound (R-C 6 H 5 -OH). Figure 41 shows the proposed phenoxyl
radical cycle in the presence of ascorbic acid with propranolol and 5-hydroxypropranolol
106

DISCUSSION
as APO substrate and phenolic product, respectively (Hofrichter et al. 2010, Kinne et al.
2009a, Poraj-Kobielska et al. 2011b).
Figure 41 Ascorbic acid as phenoxyl radical scavenger. The radical scavenging cycle is illustrated
with propranolol as APO substrate. AscH 2 -ascorbic acid, AscH - -monoprotonated (monoionic)
species of ascorbic acid, Asc 2- -diionic species, Asc •- -ascorbyl radical, Asc- dehydroascorbic acid.
4.2.8 Stoichiometry and proposed mechanism of demethylenation
The methylenedioxyphenyl moiety is found widely distributed in essential oils, alkaloids,
and other physiologically active compounds of natural origin. Compounds containing this
moiety are extensively utilized in food additives, perfumes, medicinals, topical
preparations, and insecticides (Fishbein and Falk 1969). The methylenedioxybenzenes
(1,3-benzodioxoles) contain a blocked catechol system, which is not considered to be
particularly sensitive to mild hydrolyzing, reducing, or oxidizing reagents (Hennessy
1965). Proposed metabolic pathways for the oxidation of methylenedioxyphenyl
compounds by P450s and APOs are summarized in Figure 42.
Oxidation at the methylene carbon atom (I) causes the generation of a putative 2-hydroxy-
1,3-benzodioxole intermediate (II). The intermediate gives rise to a carbene (IV) by
dehydration or to an o-hydroxyphenyl formate (V; formic acid ester). The base-catalyzed
proton abstraction would yield a carbenium ion (III) that could undergo an α-elimination
107

DISCUSSION
reaction to form also the carbene (Anders et al. 1984). The carbene may either generate
very stable 1,3-benzodioxol-2-carbene complexes (VI) with the heme of P450s in the
ferrous state, characterized by a Soret peak at 455 nm (Philpot and Hodgson 1972), or it is
hydrolyzed to the corresponding formate ester (V) (Murray 2000). Casida and co-workers
(Casida et al. 1966) have shown that the release of formate- 14 C upon scission of the
hydroxy-methylene- 14 C-dioxyphenyl group would lead to both the formation of 14 CO 2 ,
and also the introduction of formic acid- 14 C into the general metabolic pool. The
metabolism of 1,3-benzodioxoles to catechols (VII), carbon monoxide and formic acid is
consistent with hydroxylation of the carbene complex (VI) (Correia et al. 2005,
Simonneaux et al. 2006). During the conversion of 1,3-benzodioxoles with APOs,
catechols and formic acid were formed in equimolar amounts, i.e. the pathway seemingly
follows the reaction cascade from (I) via (II) and (V) to (VII) and formic acid.
The stoichiometrical experiments with AaeAPO and safrole showed that two hydrogen
peroxide molecules are consumed when one molecule of formic acid is formed (Table 20,
Chapter 3.7.2). The same ratio (H 2 O 2 :formic acid, 2:1) was obtained in stoichiometrical
experiments with 4-nitrobenzodioxole, in the case of which also the 4-nitrocatechol
formation was followed by HPLC; the molecular ratio of formic acid:4-nitrocatechol was
1:1 (Appendix, Table 1). This could mean that the conversion of 1,3-benzodioxoles to
catechols (VII) and formic acid proceeds in two enzymatic steps. A possible explanation
can be suggested (but with the proviso that also other reactions are conceivable). APOs
may catalyze the conversion of 1,3-benzodioxoles in parallel using two pathways. The first
pathway via the 2-hydroxy-1,3-benzodioxole intermediate (II) may give o-hydroxyphenyl
formate (V), which is hydrolyzed to a catechol (VII) and formic acid and in the other
pathway, it is hydroxylated to o-hydroxyphenyl formic acid (X) or the 2,2-dihydroxy-1,3-
benzodioxole intermediate (VIII) (at equilibrium with cyclic phenylene carbonate IX),
which then break down to a catechol (VII) and carbon dioxide. This would explain the
double consumption of hydrogen peroxide; however, the formation of carbon dioxide has
not been proved yet, and the observed molar ratio of formic acid and 4-nitrocatechol of 1:1
also contradicts this assumption. Considering the latter fact and the presence of ascorbic
acid in the reaction mixture, another explanation is possible. Part of the peroxide may be
"uselessly" consumed during the peroxidation of formed catechols whose phenoxyl
radicals in turn can be re-reduced by ascorbic acid (see also Figure 41). Eventually also the
catalase-like side activity of APOs that leads to the "unproductive" destruction of H 2 O 2
may have to be taken into consideration (R. Ullrich, personal communication).
108

DISCUSSION
Figure 42 Proposed mechanism of benzodioxole demethylenation by P450 and AaeAPO. I-1,3-
benzodioxole; II - 2-hydroxy-1,3-benzodioxole intermediate; III - carbanion; IV - carbene;V - o-
hydroxyphenyl formate; VI - 1,3-benzodioxol-2-carbene complexes; VII - catechol derivative;
VIII - 2,2-dihydroxy-1,3-benzodioxole intermediate; IX - cyclic phenylene carbonate; X - o-
hydroxyphenyl formic acid.
4.3 NBD assay
4-Nitrocatechol is a colorimetric, pH-selective indicator compound showing a yellow color
under acidic conditions and a red color under alkaline conditions in aqueous solutions
(Cooper and Tulane 1936b). This property has already been in use for a long time, in
various enzyme assays where 4-nitrocatechol is the diagnostic reaction product (e.g. P450
monooxygenase-catalyzed oxidation of 4-nitrophenol (Tassaneeyakul et al. 1993a,
Tassaneeyakul et al. 1993b) or sulfatase-catalysed hydrolysis of 4-nitrocatechol sulfate
(Baum et al. 1959, Dzialoszyński 1955)). Here, we developed an assay on the basis of 4-
nitrocatechol formation from nitrobenzodioxole (NBD), which is suitable to selectively
detect peroxygenase activities in complex growth media with high background absorption
such as soybean-peptone suspensions (Anh et al. 2007, Ullrich et al. 2004). Such growth
media are typically colored (high background absorption), which may interfere with
classic peroxygenase assays that follow substrate oxidation in the UV range.
The exact measurement of peroxygenase activities is an important prerequisite for studying
them in the lab, for example, during fungal fermentations in bioreactors, heterologous
expression studies, protein purification and biochemical enzyme characterization as well as
109

DISCUSSION
during the optimization of enzymatic in vitro reactions. All these approaches require
simple and rapid enzyme assays, which are selective and based on simple
spectrophotometric detection. So far, only two spectrophotometric peroxygenase assays
have been reported: (i) the oxidation of veratryl alcohol (3,4-dimethoxybenzyl alcohol) or
methyl 3,4-dimethoxybenzyl ether to veratraldehyde (3,4-dimethoxybenzaldehyde) (Kinne
et al. 2009b, Ullrich et al. 2004), which is followed in the UV range at 310 nm and lacks
full selectivity, since there is some overlap with lignin peroxidase activity under acidic
conditions (pH 3-5) (Hofrichter and Ullrich 2010, Schmidt et al. 1989, Tien et al. 1986),
and (ii) the oxidation of naphthalene to 1-naphthol (Kluge et al. 2007), which proceeds in
the UV range as well (at 303 nm). The assay established here is a simple method for the
rapid spectrophotometric detection (in the visible range) of fungal peroxygenase activity in
vivo and in vitro. The NBD assay can be applied for the spectrophotometric identification
and quantification of fungal peroxygenase activities, (i) in complex plant-based media
during fungal growth, (ii) in environmental samples, (iii) for reaction control during
enzymatic catalysis and (iv) for the detection of peroxygenase-positive fungi in the course
of high-throughput screenings of wild-type and recombinant strains as well as mutants.
4.4 Long-term and in-vivo experiments
The purification of peroxygenases is quite complicated and time-consuming, and would be
too expensive for most kinds of industrial applications (Gröbe et al. 2012, Ullrich et al.
2009). For this reason, it is important to find possibilities to apply the enzymes in a more
economic way, either by using crude preparations or extending their "life-span", for
example, by immobilization and/or re-use over several cycles (see also chapter 3.5). Initial
tests in this direction were performed by retaining and re-using the enzyme during
propranolol hydroxylation in Vivaspins® (special centrifugal concentrators with
ultrafiltration membranes). Later long-term experiments were carried out in hollow-fiber
modules holding back the enzyme from the circulating filtrate containing the reactants of
enzymatic conversion. AaeAPO turned out to be very stable under these conditions and
was successfully used over a period of two weeks. Over the whole time, the enzyme was
active and continuously hydroxylated the model substrate propranolol into 5-
hydroxypropranolol (see Figure 35, chapter 3.17.2). The cumulative molar amount of
product (50.4 µmol) and the amount of applied enzyme (7.9 nmol) show that one
peroxygenase molecule oxidized about 6,400 substrate molecules in the course of the
experiment. This is a promising basis for the further improvement of HDM production
110

DISCUSSION
with fungal peroxygenases. Existing methods use either multi-step organic synthesis
(Johansson et al. 2007, Oatis et al. 1981), whole-cell biotransformations with cell cultures
(e.g. human hepatocytes (Acikgöz et al. 2009, Li et al. 1999, Maurel 1996) or microbial
host cells (e.g. Schizosaccharomyces pombe, Nocardia autotropica or Curvularia lunata;
(Julsing et al. 2008, Peters et al. 2007) as well as, at least at lab scale, engineered P450
enzymes (Leemann et al. 1993, Schroer et al. 2010, Weis et al. 2009).
Another promising field of peroxygenase application could be the removal of
pharmaceuticals from waste water or other environmental media. The occurrence of watersoluble
and pharmacologically active organic micropollutants in the effluents of
wastewater treatment plants (WWTPs) has caused much attention over the recent years
(Lienert et al. 2007). It has been a matter of concern that the existing WWTPs are
inefficient in removing some of these pollutants and there is an increasing interest to
develop efficient techniques for achieving their destruction (Clara et al. 2005, Marco-Urrea
et al. 2009). A number of experiments with the white-rot fungus Trametes versicolor
showed that waste waters contaminated with several pharmaceuticals such as clofibric
acid, ibuprofen and carbamazepine could be cleaned to some extent by the fungal
mycelium. However, in these experiments, adsorption and transformation of
pharmaceuticals by the fungus were not clearly distinguished, nor were intra- and
extracellular enzymatic mechanisms (Blánquez et al. 2004). The concentrations tested in
this study (~10 mg L -1 ) were also three to five orders of magnitude higher than those
usually present in effluents of municipal sewage plants (0.1-50-µg L -1 range). Subsequent
studies dealing with the analgesic ketoprofen at an environmentally relevant concentration
(40 µg L -1 ) showed that the substrate was almost completely removed by T. versicolor
(Marco-Urrea et al. 2010b). Tests with environmentally relevant concentrations of
propranolol carried out herein demonstrated that AaeAPO is able to remove about 50% of
the compound at a concentration as low as 1.4 µg L -1 . The experiments were done with the
purified enzyme and respective controls, so that all other paths of substance removal such
as absorption or involvement of other enzymes than APO could be ruled out. The positive
result obtained, despite the fact that the K m for propranolol is rather high (1,340 µM), can
be explained by the reaction cycle of APO and the particular reaction conditions. Thus, the
continuous dosage of hydrogen peroxide with a syringe pump may have ensured the
(pre)formation of sufficient amounts of a reactive APO species (compound I; (Wang et al.
2012). The presence of a certain steady-state concentration of compound I (spontaneous
decomposition rate 1.4 s -1 ) combined with a sufficient reaction time (>12 hours) under
111

DISCUSSION
stirring may enable the oxidation of quite low amounts of the substrate independent on the
high apparent K m value.
In vivo experiments with A. aegerita were carried out to follow the fate of two
pharmaceutical model compounds (propranolol, diclofenac) in fungal cultures and to find
indications for the involvement of APOs in biodegradation processes under natural
conditions. The transformation of the anti-inflammatory drug diclofenac had already been
studied in liquid cultures of Phanerochaete chrysosporium under fed-batch conditions. The
fungus started to eliminate diclofenac (34 µM) already after two hours (77% removal) and
after 18 hours, 94% of the compound were converted (Rodarte-Morales et al. 2012). The
capability of diclofenac degradation by T. versicolor was demonstrated by Marco-Urrea et
al. (Marco-Urrea et al. 2010a). They observed an almost complete diclofenac removal
(>94%) within the first hour of fungal treatment in the presence of fungal pellets; the drug
was added both at relatively high (10 mg L -1 ) and low, environmentally relevant (45 µg L -
1 ) concentrations in a defined liquid medium. During this treatment, 4'-hydroxydiclofenac
and 5-hydroxydiclofenac were detected as metabolites, which disappeared within the next
24 hours (Cruz-Morató et al. 2012, Marco-Urrea et al. 2010a). Another example of using
the fungal mycelium as biodegradative agent is the so-called biologically advanced
oxidation of propranolol by T. versicolor (80% removal within 6 hours of incubation). In
this case, extracellular oxidizing species (e.g. hydroxyl radicals) were thought to be
produced through a quinone redox cycling mechanism catalyzed by an intracellular
quinone reductase and ligninolytic enzymes of Trametes versicolor, after addition of
lignin-derived 2,6-dimethoxy-1,4-benzoquinone and Fe 3+ -oxalate (Marco-Urrea et al.
2010c).
Diclofenac and propranolol also readily disappeared in liquid cultures of A. aegerita,
interestingly when a high peroxygenase level (ca. 600 U L -1 ) was present. The
experiments, carried out in parallel in two different growth media (one of which stimulated
peroxygenase production (Ullrich et al. 2004)) indeed indicated an involvement of
peroxygenase in this process. However, no transformation products were detectable by
HPLC, probably due to coupling/polymerization reactions, which could not be prevented
in contrast to in vitro experiments where ascorbic acid served as radical scavenger.
Furthermore, analyses of control cultures of A. aegerita (soybean medium) showed the
presence of numerous aromatic ingredients, which disappeared in the course of fungal
growth. UV-Vis spectra of these substances indicated a polyphenolic nature with
similarities to flavonoids (Figure 30) (Barková et al. 2011). One-electron oxidation of
112

DISCUSSION
these compounds would yield phenoxyl radicals, which may force the overall
polymerization process. The peroxidative activity of AaeAPO, or the activity of laccase
which is also secreted by A. aegerita (Ullrich et al. 2004), may be responsible for these
reactions. Interestingly, in both growth media (soybean medium, Czapek-Dox) the color
changed to violet when diclofenac was added on day six. The same color was observed in
in vitro studies with APOs in the absence of ascorbic acid (Appendix Figure 2). On the
basis of UV-Vis and mass spectra as well as literature data from Miyamoto at al. (1997)
we propose that this compound is the one-electron-oxidation product diclofenac-1',4'-
quinone imine (Figure 43) (Miyamoto et al. 1997, Waldon et al. 2010).
Figure 43 Proposed mechanism of diclofenac-1',4'-quinone imine formation by AaeAPO in vitro
without ascorbic acid and by by fungal cultures. In Czapek-Dox medium, conversion may be only
possible by intracellular 450s (no peroxygenase formation; laccase alone is not able to catalyze this
reaction) (Marco-Urrea et al. 2010a). In soybean medium, this pathway could be realized both by
intracellular P450s or secreted AaeAPO or a mixed AaeAPO-laccase system (Figure 48;
(Hofrichter et al. 2010, Ullrich and Hofrichter 2007, Waldon et al. 2010).
4.5 Homology model structures of APOs.
In order to better understand the catalytic properties of APOs and differences between
them, homology modelling was performed. The full-length sequences of the
peroxygenases from A. aegerita, C. radians and M. rotula have recently been obtained and
confirm the enzymes’ affiliation to the heme-thiolate proteins. The sequences revealed no
homology to classic heme peroxidases or cytochrome P450 enzymes, and only little
113

DISCUSSION
114
DISCUSSION
Figure 45 Homology model structures of CraAPO (left), AaeAPO (middle) and MroAPO (right). Homology modelling was performed with I-TASSER
software (Roy et al. 2010) using the crystal structure of unspecific peroxygenase of A. aegerita as basis [(Piontek et al. 2010), as well as unpublished data, the
same authors]. The resulting models of CraAPO and MroAPO were superimposed with the AaeAPO structure using the PyMOL program (2010), resulting in
a root-mean-square deviation (RMS) of 1.133 over 295 aligned residues for CraAPO, and a RMS of 1.381 over 177 aligned residues for MroAPO. Prepared
by Marek Pecyna and Marzena Poraj-Kobielska (2012).

DISCUSSION
homology (

DISCUSSION
For comparison, the structure of propranolol, which is smaller and a good substrate of the
AaeAPO, was placed into the active site and fit it well (Figure 42, examples of other
structures are given in the Appendix). The homology model of CraAPO shown in Figure
45 indicates that the entrance to the heme channel is about the size of that of AaeAPO and
the active site volume is even larger, but one phenylalanine seems to block the entrance
(Figure 49).
The position at the end of the polypeptide chain (C-terminus) indicates that this
phenylalanine may not be fixed and can move to some extent to open and close the
entrance of the heme channel, i.e. it may act as a sort of molecular "gatekeeper". This may
explain the overall lower conversion rates observed for CraAPO compared to AaeAPO, in
spite of a similar amino acids sequence and configuration (59% sequence identity and 69%
similarity; for comparison, MraAPO shares only about 30% sequence identity with both
enzymes) (Pecyna et al. 2009), personal communication). Of course, more precise
predictions will only be possible when the crystal structures of all three APOs will have
been published.
Figure 48 Amino acids that possess reactive groups and are found in the active sites of AaeAPO,
CraAPO and MroAPO. Tyr - tyrosine, Phe - phenylalanine, Glu – glutamic acid, Thr – threonine,
Ser – serine and Arg - arginine.
Figure 49 shows homology model structures of the active site of the three APOs studied
here.
The channels of AaeAPO and CraAPO are rich in aromatic amino acids, mainly
phenylalanines (Phe) and a tyrosine (Tyr), whereas in MroAPO, almost exclusively
aliphatic amino acids are found (Ile, Leu, Val, Ala). Phe is a hydrophobic aromatic amino
acid with a benzyl side chain; Tyr possesses also an aromatic ring that is however less
hydrophobic than that of Phe due to its phenolic OH-group.
116

DISCUSSION
117
Figure 46 Homology models of peroxygenase channels (solvent accessible surface calculation) leading to the active sites of AaeAPO (B) and MroAPO (C)
fitting codeine. Dimensions (in Ǻ) of the heme channels are shown in (A; AaeAPO) and (F; MroAPO). A 3-Dimensional model of codeine is given in D1, D2
and D3 from different perspectives. (E) illustrates the active site's architecture of MroAPO with codeine as ball-and-stick model: heme (red), magnesium ion
(green), codeine substrate (violet), amino acids (gray).

DISCUSSION
118
DISCUSSION
Figure 47 Homology models of heme channels with active sites fitting hydrocortisone. AaeAPO from the top (A) and from the side (B) with channel length in
Ǻ and MroAPO from the top (F) and side (E). (C) Electron surface density of hydrocortisone with the length of the structure, and (D) stick model of
hydrocortisone from two other perspectives (dimensions in Ǻ).

DISCUSSION
Figure 49 Heme channels leading to the active site of AaeAPO, CraAPO and MroAPO, including
important amino acid residues. Aromatic amino acids (orange): Phe - phenylalanine, Tyr - tyrosine;
functional amino acids (blue): Glu - glutamic acid, Ser – serine, Arg - arginine, Thr - threonine;
aliphatic amino acids (grey): Ile - isoleucine, Leu - leucine, Ala - alanine, Gly - glycine, Val -
valine and Pro - proline. One example of each kind of amino acid is labeled with the respective
abbreviation (model and figure prepared by M. Pecyna and M. Poraj-Kobielska, 2012).
119

DISCUSSION
Near the heme and the stabilizing magnesium, three aliphatic amino acids are found:
glutamic acid (Glu), arginine (Arg) and threonine (Thr; Figure 48)(Ophardt 2003). Glu has
a carboxylic side chain and can exist as its negatively charged deprotonated carboxylate
(Pecyna et al. 2009, Piontek et al. 2010, Sundaramoorthy et al. 1995). As an acid-base
catalyst, it may play a key role in peroxide binding and cleavage, and thereby in the
formation of AaeAPO compound I (Wang et al. 2012). Arg may act as a charge stabilizer
in this process as in CPO (Sundaramoorthy 1995) and Thr is probably the proton acceptor.
In MroAPO, serine (Ser) seemingly replaces Arg. In classical peroxidases, Glu is
substituted by a histidine (His), whereas in DyP-type peroxidases an aspartic acid (Asp)
replaces Glu while the charge stabilizer is again an Arg (Sugano et al. 2007).
4.6 Key findings
1. All tested fungal unspecific/aromatic peroxygenases (APOs from Agrocybe
aegerita, Coprinellus radians, Marasmius rotula) catalyzed the
oxygenation/hydroxylation or N-/O-dealkylation (Figure 50) of diverse
pharmaceuticals and illicit drugs, in most cases with high regioselectivity. Many of
the oxidation products were identical to human drug metabolites formed by P450s.
2. The peroxygenase of M. rotula (MroAPO) was moreover found to catalyze the
conversion of different steroids with preference for those bearing an oxohydroxyethyl
side chain.
3. The different substrate specificities and selectivities of individual APOs raise the
possibility to choose the appropriate enzyme according to the needed product
("catalytic toolbox" for oxyfunctionalizations).
4. The AaeAPO-catalyzed oxygenation of aromatic pharmaceuticals such as
propranolol, diclofenac or carbamazepine and the hydroxylation of benzylic
carbons in compounds such as tolbutamide resulted in the incorporation of an 18 O-
atom from H 18 2 O 2 into the reaction products (phenols, alcohols), which identifies
these reactions as true peroxygenations.
5. The stoichiometry of AaeAPO-catalyzed sildenafil oxidation showed that the
reaction was a two-electron oxidation yielding an N-dealkylated product (Ndesmethylsildenafil)
and an aldehyde (formaldehyde).
6. The deethylenation of phenacetin-d1 by AaeAPO showed an observed
intramolecular deuterium isotope effect (k H /k D ) obs of 3.1, which is close to the
values that have been observed for the P450-catalyzed O-dealkylation of this
120

DISCUSSION
substrate. This indicates that phenacetin oxidation by APOs may proceed via an
electron transfer rather than a hydrogen abstraction mechanism.
7. The kinetic data and particularly the catalytic turnover numbers (k cat ) determined
for the conversion of various pharmaceuticals by AaeAPO suggest that APOs may
be at least as efficient as the best P450s and hence represent useful alternatives for
the regioselective preparation of human drug metabolites (HDMs).
8. Steady-state bisubstrate kinetics of the AaeAPO-catalyzed demethylenation of
nitro-1,3-benzodioxole (NBD), which yielded catechol and formic acid, gave
parallel double reciprocal plots suggestive of a ping-pong mechanism.
9. The NBD assay established on the basis of this reaction is a simple method for the
rapid spectrophotometric detection of fungal peroxygenase activities in vivo and in
vitro.
10. The testing of twelve validated P450 assays with the three APOs showed that they
possess a substrate spectrum and an activity range that overlap with those of the
most important human P450s involved in drug metabolism.
11. Stoichiometry analysis of AaeAPO-catalyzed safrole oxidation showed that two
hydrogen peroxide molecules were consumed during the formation of one molecule
of products (catechol and formic acid). This indicates a divergent mechanism that
may involve two different pathways and/or unproductive peroxide-consuming side
reactions.
12. APOs are able to use, in addition to H 2 O 2 , different organic peroxides as cooxidant,
and they efficiently catalyze oxidation reactions in the presence of
different peroxide-generating systems.
13. AaeAPO was capable of oxidizing the pharmaceutical propranolol at a very low,
environmentally relevant concentration (in the ppb range).
14. APOs could be used in long-term in-vitro experiments over several days for the
constant synthesis of the HDM, 5-hydroxypropranolol.
15. Propranolol and diclofenac disappeared rapidly when added to growing cultures of
A. aegerita indicating fungal drug degradation also under natural conditions.
16. Homology models of APOs provide explanations for limitations observed with
regard to the substrate size and structure. The size of the entrance to the heme
channel, the amino acids there and the active site volume seem to be key
parameters determing peroxygenase performance.
121

DISCUSSION
122
DISCUSSION
Figure 50 Mechanistic routes of reactions catalyzed by fungal peroxygenases with pharmaceuticals and drugs. From left to right: demethylenation (ecstasy),
N-dealkylation (lidocaine), O-dealkylation (naproxen), aliphatic side chain hydroxylation (tolbutamide), aromatic epoxidation/hydroxylation (diclofenac),
phenol oxidation (morphine); modified according to (Hofrichter et al. 2010)

DISCUSSION
The most important finding of this work is that APOs are capable of oxidizing a large
variety of pharmaceuticals and other drugs, often yielding typical human drug metabolites.
This opens the fascinating possibility to produce diverse human drug metabolites in the
future using a "peroxygenase toolbox", figuratively speaking as an artificial liver in the
reaction tube.
4.7 Outlook
In the meantime, one of the most promising approaches in peroxygenase research, the
empirical comparison of as many APOs as possible to obtain a broad selection of
biocatalysts with different substrate specificities, has taken shape. Thus, recent genetic
investigations have resulted in a phylogenic tree, which impressively shows the widespread
occurrence of peroxygenases in the whole fungal kingdom (Pecyna et al. 2009). New wildtype
peroxygenases have been found in a number of basidiomycetes, for example, in
Coprinopsis verticillata (Anh et al. 2007), Agrocybe parasitica and Auricularia auriculajudae
(R. Ullrich 2012, personal communication). Furthermore, the first attempts to
heterologously produce peroxygenases in suitable hosts (e.g. Aspergillus spp.,
Saccharomyces cerevisiae) have been successful (H. Lund, Novozymes A/S & M. Alcalde,
CSIC Madrid, 2012; personal communications). The above developments open the
possibility to extend the APO-based “monooxygenation toolbox” for the production of
other HDMs and fine chemicals. Also the preparation of HDMs and fine chemicals even at
industrial scale seems to be reasonable taking into account the positive long-term
experiments presented here and assuming that the heterologous mass production of
peroxygenases will be successful in the near future. Environmental studies performed over
the last years indicate that there is a strong need to remove pharmaceuticals and their
metabolites from the environment. In this context, peroxygenases could become an
innovative biocatalytic tool to reduce the risk potential of pharmaceuticals. The properties
of peroxygenases such as: i) extracellular location, ii) active secretion by the fungi, iii)
high stability and iv) independence from complex, expensive cofactors, as well as the
positive results in the removal of low-concentration pharmaceuticals make the application
of APOs even in the water-treatment sector conceivable. Again, the heterologous largescale
production of peroxygenases, i.e. their availability in tons per year, will be de rigueur
to achieve this challenging goal.
123

DISCUSSION
Figure 51 Phylogenetic tree of peroxygenases (unpublished results, Kellner 2012).
124

REFERENCES
5 References
Acikgöz, A., Karim, N., Giri, S., Schmidt-Heck, W. and Bader, A. (2009), Two
compartment model of diazepam biotransformation in an organotypical culture of
primary human hepatocytes, Toxicol Appl Pharmacol 234, 179-91.
Alexander, N. J., Hohn, T. M. and McCormick, S. P. (1998), The TRI11 Gene of Fusarium
sporotrichioides Encodes a Cytochrome P-450 Monooxygenase Required for C-15
Hydroxylation in Trichothecene Biosynthesis, Appl Environ Microb 64, 221-25.
Alkatheeri, Wasfi and Lambert (1999), Pharmacokinetics and metabolism of ketoprofen
after intravenous and intramuscular administration in camels, J Vet Pharmacol
Ther 22, 127-35.
Allain, E. J., Hager, L. P., Deng, L. and Jacobsen, E. N. (1993), Highly enantioselective
epoxidation of disubstituted alkenes with hydrogen peroxide catalyzed by
chloroperoxidase, J Am Chem Soc 115, 4415-16.
Anders, M. W., Sunram, J. M. and Wilkinson, C. F. (1984), Mechanism of the metabolism
of 1,3-benzodioxoles to carbon monoxide, Biochem Pharmacol 33, 577-80.
Andersson, T., Miners, J. O., Veronese, M. E. and Birkett, D. J. (1994), Diazepam
metabolism by human liver microsomes is mediated by both S-mephenytoin
hydroxylase and CYP3A isoforms., Br J Clin Pharmacol 38, 131-37.
Andersson, T., Miners, J. O., Veronese, M. E., Tassaneeyakul, W., Tassaneeyakul, W.,
Meyer, U. A. and Birkett, D. J. (1993), Identification of human liver cytochrome
P450 isoforms mediating omeprazole metabolism, Br J Clin Pharmacol 36, 521-30.
Anh, D. H. (2008), Enzymatic and molecular characterization of haloperoxidaseperoxygenase
from litter- and dung-dwelling agaric fungi, International graduate
school Zittau, Zittau
Anh, D. H., Ullrich, R., Benndorf, D., Svatos, A., Muck, A. and Hofrichter, M. (2007), The
coprophilous mushroom Coprinus radians secretes a haloperoxidase that catalyzes
aromatic peroxygenation, Appl Environ Microbiol 73, 5477-85.
Ankley, G. T., Brooks, B. W., Huggett, D. B. and Sumpter, a. J. P. (2007), Repeating
History: Pharmaceuticals in the Environment, Environ Sci Technol 41, 8211-17.
Anzenbacher, P. and Anzenbacherová, E. (2001), Cytochromes P450 and metabolism of
xenobiotics, Cell Mol Life Sci 58, 737-47.
Appleby, A. C. (1967), A soluble haemoprotein P 450 from nitrogen-fixing Rhizobium
bacteroides, BIOCHIM BIOPHYS ACTA (2), 399-402.
Aranda, E., Kinne, M., Kluge, M., Ullrich, R. and Hofrichter, M. (2009a), Conversion of
dibenzothiophene by the mushrooms Agrocybe aegerita and Coprinellus radians
and their extracellular peroxygenases, Appl Microbiol Biot 82, 1057-66.
Aranda, E., Ullrich, R. and Hofrichter, M. (2009b), Conversion of polycyclic aromatic
hydrocarbons, methyl naphthalenes and dibenzofuran by two fungal
peroxygenases, Biodegradation 21, 267-81.
Baillie, T. A. (2009), Approaches to the assessment of stable and chemically reactive drug
metabolites in early clinical trials, CHEM RES TOXICOL 22, 263-66.
Baillie, T. A.et al. (2002), Drug metabolites in safety testing, Toxicol Appl Pharmacol 182,
188-96.
Ballard, S. A., Gingell, C. J., Tang, K., Turner, L. A., Price, M. E. and Naylor, A. M.
(1998), Effects of sildenafil on the relaxation of human corpus cavernosum tissue in
vitro and on the activities of cyclic nucleotide phosphodiesterase isozymes, J Urol
159, 2164-71.
Banerjee, N. C., Miller, G. E. and Stowe, C. M. (1967), Excretion of aminopyrine and its
metabolites into cows' milk, Toxicol. Appl. Pharmacol. 10, 604-12.
125

126
REFERENCES
Barková, K., Kinne, M., Ullrich, R., Hennig, L., Fuchs, A. and Hofrichter, M. (2011),
Regioselective hydroxylation of diverse flavonoids by an aromatic peroxygenase,
Tetrahedron 67, 4874-78.
Baum, H., Dodgson, K. and Spencer, B. (1959), The assay of arylsulphatases A and B in
human urine, Clin. Chim. Acta. 4, 453-55.
Beausse, J. (2004), Selected drugs in solid matrices: a review of environmental
determination, occurrence and properties of principal substances, TrAC-Trend
Anal Chem 23, 753-61.
Bergé, J., Feyereisen, R. and Amichot, M. (1998), Cytochrome P450 monooxygenases and
insecticide resistance in insects, Philos T Roy Soc B 353, 1701-05.
Bernhardt, R. (2006), Cytochromes P450 as versatile biocatalysts, J BIOTECHNOL 124,
128-45.
Bezalel, L., Hadar, Y., Fu, P. P., Freeman, J. P. and Cerniglia, C. E. (1996), Initial
oxidation products in the metabolism of pyrene, anthracene, fluorene, and
dibenzothiophene by the white rot fungus Pleurotus ostreatus, Appl Environ
Microbiol 62, 2554-9.
Bisswanger, H. (2000), Enzymkinetik Theorie und Methoden, Wiley-VCH Verlag,
Weinheim.
Bland, T. M., Haining, R. L., Tracy, T. S. and Callery, P. S. (2005), CYP2C-catalyzed
delta(9)-tetrahydrocannabinol metabolism: Kinetics, pharmacogenetics and
interaction with phenytoin, Biochem Pharmacol 70, 1096-103.
Blánquez, P., Casas, N., Font, X., Gabarrell, X., Sarrá , M., Caminal, G. and Vicent, T.
(2004), Mechanism of textile metal dye biotransformation by Trametes versicolor,
Water Res 38, 2166-72.
Bolwell, G. P., Bozak, K. and Zimmerlin, A. (1994), Plant cytochrome p450,
Phytochemistry 37, 1491-506.
Boolell, M., Allen, M. J., Ballard, S. A., Gepi-Attee, S., Muirhead, G. J., Naylor, A. M.,
Osterloh, I. H. and Gingell, C. (1996), Sildenafil: an orally active type 5 cyclic
GMP-specific phosphodiesterase inhibitor for the treatment of penile erectile
dysfunction, Internat Journal Impot Res 8, 47-52.
Booth, K. S., Kimura, S., Lee, H. C., Ikeda-Saito, M. and Caughey, W. S. (1989), Bovine
myeloperoxidase and lactoperoxidase each contain a high affinity site for calcium,
Biochem Biophys Res Co 160, 897-902.
Bort, R., Macé, K., Boobis, A., Gómez-Lechón, M.-J., Pfeifer, A. and Castell, J. (1999),
Hepatic metabolism of diclofenac: role of human CYP in the minor oxidative
pathways, Biochem. Pharmacol. 58, 787-96.
Bossche, H. V. and Koymans, L. (1998), Review Article Cytochromes P450 in fungi,
Mycoses 41, 32-38.
Boxall, A. B. A., Sinclair, C. J., Fenner, K., Kolpin, D. and Maund, S. J. (2004), Peer
reviewed: when synthetic chemicals degrade in the environment, Environ Sci
Technol 38, 368A-75A.
Breskvar, K., Ferenčak, Z. and Hudnik-Plevnik, T. (1995), The role of cytochrome
P45011α in detoxification of steroids in the filamentous fungus Rhizopus nigricans,
J Steroid Biochem Mol Bio 52, 271-75.
Brodie, B. B. and Axelrod, J. (1949), The fate of acetophenetidin (phenacetin) in man and
methods for the estimation of acetophenetidin and its metabolites in biological
material, J. Pharmacol. Exp. Ther. 97, 58-67.
Brooks, B. W., Chambliss, C. K., Stanley, J. K., Ramirez, A., Banks, K. E., Johnson, R. D.
and Lewis, R. J. (2005), Determination of select antidepressants in fish from an
effluent-dominated stream, Environ Toxicol Chem 24, 464-69.
Caputi, L., Di Tullio, A., Di Leandro, L., De Angelis, F. and Malatesta, F. (2005), A new
microperoxidase from Marinobacter hydrocarbonoclasticus, Biochim. Biophys.
Acta 1725, 71-80.

REFERENCES
Cardini, G. and Jurtshuk, P. (1968), Cytochrome P-450 involvement in the oxidation of n-
octane by cell-free extracts of corynebacterium sp. strain 7E1C, J BIOL CHEM
243, 6070-72.
Carrara, C., Ptacek, C. J., Robertson, W. D., Blowes, D. W., Moncur, M. C., Sverko, E.
and Backus, S. (2008), Fate of Pharmaceutical and Trace Organic Compounds in
Three Septic System Plumes, Ontario, Canada, Environ Sci Technol 42, 2805-11.
Casida, J. E., Engel, J. L., Essac, E. G., Kamienski, F. X. and Kuwatsuka, S. (1966),
Methylene-C 14 -Dioxyphenyl Compounds: Metabolism in Relation to Their
Synergistic Action, Science 153, 1130-33.
Celiz, M. D., Tso, J. and Aga, D. S. (2009), Pharmaceutical metabolites in the
environment: Analytical challenges and ecological risks, Environ Toxicol Chem
28, 2473-84.
Center for Drug Evaluation and Research (HFD-130) and Food and Drug Administration
(2008), Guidance for Industry on Safety Testing of Drug Metabolites; Availability,
Food Drug Admin 1-11.
Chapple, C. (1998), Molecular-genetic analysis of plant cytochrome P450-dependent
monooxygenases, Annu Rev Plant Phys 49, 311-43.
Chen, Z. R., Somogyi, A. A., Reynolds, G. and Bochner, F. (1991), Disposition and
metabolism of codeine after single and chronic doses in one poor and seven
extensive metabolisers., Br J Clin Pharmacol 31, 381–90.
Cirino, P. C. and Arnold, F. H. (2003), Exploring the diversity of heme enzymes through
directed evolution, in: Directed molecular evolution of proteins, Vol. 215-43 (Dr.
Susanne Brakmann, P. D. K. J., Ed.)
Clara, M., Strenn, B., Gans, O., Martinez, E., Kreuzinger, N. and Kroiss, H. (2005),
Removal of selected pharmaceuticals, fragrances and endocrine disrupting
compounds in a membrane bioreactor and conventional wastewater treatment
plants, Water Res 39, 4797-807.
Cleland, W. W. (1963), The kinetics of enzyme-catalyzed reactions with two or more
substrates or products: III. Prediction of initial velocity and inhibition patterns by
inspection, Biochim Biophys A Spec Sec Enzymol Sub 67, 188-96.
Clouse, S. D. and Sasse, J. M. (1998), Brassinosteroids: essential regulators of plant
growth and development, Annu Rev Plant Phys 49, 427-51.
Colonna, S., Gaggero, N., Manfredi, A., Casella, L., Gullotti, M., Carrea, G. and Pasta, P.
(1990), Enantioselective oxidations of sulfides catalyzed by chloroperoxidase,
Biochemistry 29, 10465-68.
Cooper, S. R. and Tulane, V. J. (1936a), Action of 4-Nitrocatechol as a Titration Indicator,
Ind Eng Chem Anal Ed 8, 210-11.
Cooper, S. R. and Tulane, V. J. (1936b), Action of 4-Nitrocatechol as a Titration Indicator,
Industrial & Engineering Chemistry Analytical Edition 8, 210-11.
Correia, M., Ortiz de Montellano, P. and Ortiz de Montellano, P. R. (2005), Inhibition of
Cytochrome P450 Enzymes
Cytochrome P450, 247-322 (Springer US,
Crameri, A., Raillard, S.-A., Bermudez, E. and Stemmer, W. P. C. (1998), DNA shuffling
of a family of genes from diverse species accelerates directed evolution, Nature
391, 288-91.
Črešnar, B. and Petrič, Š. Cytochrome P450 enzymes in the fungal kingdom, BBA-Protein
Struct M 1814, 29-35.
Cruz-Morató, C.et al. (2012), Biodegradation of pharmaceuticals by fungi and metabolites
identification
1-49 (Springer Berlin / Heidelberg,
Dallacosta, C., Monzani, E. and Casella, L. (2003), Reactivity study on microperoxidase-8,
J. Biol. Inorg. Chem. 8, 770-76.
127

128
REFERENCES
Damsten, M. C., van Vugt-Lussenburg, B. M. A., Zeldenthuis, T., de Vlieger, J. S. B.,
Commandeur, J. N. M. and Vermeulen, N. P. E. (2008), Application of drug
metabolising mutants of cytochrome P450 BM3 (CYP102A1) as biocatalysts for the
generation of reactive metabolites, Chem-Biol Interact 171, 96-107.
Daughton, C. G. and Ternes, T. A. (1999), Pharmaceuticals and personal care products in
the environment: agents of subtle change?, Environ Health Perspectiv 107, 907-38.
Davey, C. A. and Fenna, R. E. (1996), 2.3 Å resolution X-ray crystal structure of the
bisubstrate analogue inhibitor salicylhydroxamic acid bound to human
myeloperoxidase: a model for a prereaction complex with hydrogen peroxide,
Biochemistry 35, 10967-73.
Davies, M. B. (1984), The copper(II) catalyzed reaction of L-ascorbinc acid with
tris(oxalato)cobaltate(III) ion in aqueous solution, Inorg Chim Acta 92, 141-46.
Dawson, J. H. and Sono, M. (1987), Cytochrome P-450 and chloroperoxidase: thiolateligated
heme enzymes. Spectroscopic determination of their active-site structures
and mechanistic implications of thiolate ligation, Chemical Rev 87, 1255-76.
Degtyarenko, K. (2004), EC numbers for P450 enzymes, website,
Desjardins, A. E. and VanEtten, H. D. (1986), Partial purification of pisatin demethylase,
a cytochrome P-450 from the pathogenic fungus Nectria haematococca, Arch
Microbiol 144, 84-90.
Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH, L.-I. D. (2012),
Coprinus radians (Desmazieres) Fries
Ding, X. and Kaminsky, L. S. (2003), Human extrahepatic cytochromes P450: function in
xenobiotic metabolism and tissue-selective chemical toxicity in the respiratory and
gastrointestinal tracts, Annu Rev Pharmacol 43, 149-73.
Doddapaneni, H., Chakraborty, R. and Yadav, J. S. (2005), Genome-wide structural and
evolutionary analysis of the P450 monooxygenase genes (P450ome) in the white rot
fungus Phanerochaete chrysosporium : Evidence for gene duplications and
extensive gene clustering, BMC Genomics 6
Dorovska-Taran, V., Posthumus, M. A., Boeren, S., Boersma, M. G., Teunis, C. J.,
Rietjens, I. M. and Veeger, C. (1998), Oxygen exchange with water in heme-oxo
intermediates during H 2 O 2 -driven oxygen incorporation in aromatic hydrocarbons
catalyzed by microperoxidase-8., Eur. J. Biochem. 253, 659-568.
Dzialoszyński , L. (1955), 4-Nitrocatechol sulfate as a substrate for the assay of aryl
sulfatase, Acta. Biochim. Pol. 2, 421-28.
Eiben, S., Kaysser, L., Maurer, S., Kuhnel, K., Urlacher, V. B. and Schmid, R. D. (2006),
Preparative use of isolated CYP102 monooxygenases-A critical appraisal, J
BIOTECHNOL 124, 662-9.
ElSohly, M. A. and Feng, S. (1998), Δ9-THC Metabolites in Meconium: Identification of
11-OH-Δ9-THC, 8β,11-diOH-Δ9-THC, and 11-nor-Δ9-THC-9-COOH as Major
Metabolites of Δ9-THC J Anal Toxicol 22, 329-35.
Ferris, J. P., MacDonald, L. H., Patrie, M. A. and Martin, M. A. (1976), Aryl hydrocarbon
hydroxylase activity in the fungus Cunninghamella bainieri: Evidence for the
presence of cytochrome P-450, Arch Biochem Biophys 175, 443-52.
Feyereisen, R. (1999), Insect P450 enzymes, Annu Rev Entomol 44, 507-33.
Feyereisen, R. (2006), Evolution of insect P450, Biochem Soc Trans 34, 1252-5.
Fiedler, T. J., Davey, C. A. and Fenna, R. E. (2000), X-ray crystal structure and
characterization of halide-binding sites of human myeloperoxidase at 1.8 Ǻ
resolution, J BIOL CHEM 275, 11964-71.
Fishbein, L. and Falk, H. L. (1969), Environmental aspects of methylenedioxyphenyl and
related derivatives, Environ Res 2, 297-320.
Frear, D. S. (1995), Wheat microsomal cytochrome P450 monooxygenases:
characterization and importance in the metabolic detoxification and selectivity of
wheat herbicides, Drug metab drug interact 12, 329-57.

REFERENCES
Fricke, U., Zawinell, A., Zeidan, R. and Steden, M. (2012), Anatomisch-therapeutischchemische-Klassifikation
mit tagesdosen. Amtliche fassung des ATC-Index mit
DDD-Angabe für Deutschland im Jahre 2012, Deutsches Institut für Medizinische
Dokumentation und Information 1-220.
Fulco, A. J. (1991), P450BM-3 and other inducible bacterial P450 cytochromes:
biochemistry and regulation, Annu Rev Pharmacol 31, 177-203.
Garland, W. A., Hsiao, K. C., Pantuck, E. J. and Conney, A. H. (1977), Quantitative
determination of phenacetin and its metabolite acetaminophen by GLC-chemical
ionization mass spectrometry, J Pharm Sci 66, 340-4.
Garland, W. A., Nelson, S. D. and Sasame, H. A. (1976), Primary and secondary
deuterium isotope effects in the O-deethylation of phenacetin, Biochem Biophys
Res Co 72, 539-45.
German Association for Pharmaceutical Industry (2011), Pharma-Data 2011, German
Association for Pharmaceutical Industry (BPI e.V.),
Gibson, G. G. and Skett, P. (2001), Introduction to drug metabolism, viii, 256 p. (Nelson
Thornes Publishers, Cheltenham, UK.
Gibson, G. G. and Skett, P. (2008), Introduction to Drug Metabolism, 256 (Nelson
Thornes, London.
Gibson, M., Soper, C. J., Parfitt, R. T. and Sewell, G. J. (1984), Studies on the mechanism
of microbial N-demethylation of codeine by cell-free extracts of Cunninghamella
bainieri, Enzyme Microb Tech 6, 471-75.
Gillam, E. M. (2008), Engineering cytochrome P450 enzymes, CHEM RES TOXICOL 21,
220-31.
Gorsky, L. D., Koop, D. R. and Coon, M. J. (1984), On the stoichiometry of the oxidase
and monooxygenase reactions catalyzed by liver microsomal cytochrome P-450.
Products of oxygen reduction, J BIOL CHEM 259, 6812-17.
Greenberg, L. A. and Lester, D. (1946), The metabolic fate of acetanilid and other aniline
derivates, J. Pharmacol. Exp. Ther. 88, 87-98.
Gröbe, G., Ullrich, R., Pecyna, M., Kapturska, D., Friedrich, S., Hofrichter, M. and
Scheibner, K. (2012), High-yield production of aromatic peroxygenase by the
agaric fungus Marasmius rotula, AMB Express 1, 1-11.
Groves, J. T. (2006), High-valent iron in chemical and biological oxidations, J Inorg
Biochem 100, 434-47.
Groves, J. T. and McClusky, G. A. (1978), Aliphatic hydroxylation by highly purified liver
microsomal cytochrome P-450. Evidence for a carbon radical intermediate,
Biochem Biophys Res Commun 81, 154-60.
Guengerich, F. P. (1987), Oxidative cleavage of carboxylic esters by cytochrome P-450, J.
Biol. Chem. 262, 8459-62.
Guengerich, F. P. (2001), Common and uncommon cytochrome P450 reactions related to
metabolism and chemical toxicity, CHEM RES TOXICOL 14, 611-50.
Guengerich, F. P., Hosea, N. A., Parikh, A., Bell-Parikh, L. C., Johnson, W. W., Gillam, E.
M. J. and Shimada, T. (1998), Twenty years of biochemistry of human P450s, Drug
Metab Dispos 26, 1175-78.
Guengerich, F. P. and Montellano, P. R. (2005), Human cytochrome P450 enzymes
Cytochrome P450, 377-530 (Springer US,
Hager, L. P., Morris, D. R., Brown, F. S. and Eberwein, H. (1966), Chloroperoxidase. II.
Utilization of halogen anions., J BIOL CHEM 241, 1769-77.
Hall, P. F. (1986), Cytochromes P-450 and the regulation of steroid synthesis, Steroids 48,
131-96.
Hamman, M. A., Thompson, G. A. and Hall, S. D. (1997), Regioselective and
stereoselective metabolism of ibuprofen by human cytochrome P450 2C, Biochem.
Pharmacol. 54, 33-41.
129

130
REFERENCES
Hanano, A., Burcklen, M., Flenet, M., Ivancich, A., Louwagie, M., Garin, J. and Blee, E.
(2006), Plant seed peroxygenase is an original heme-oxygenase with an EF-hand
calcium binding motif, J BIOL CHEM 281, 33140-51.
Hannemann, F., Bichet, A., Ewen, K. M. and Bernhardt, R. (2007), Cytochrome P450
systems - biological variations of electron transport chains, BIOCHIM BIOPHYS
ACTA 1770, 330-44.
Hansen, L. L. and Brøsen, K. (1999), Quantitative determination of tolbutamide and its
metabolites in human plasma and urine by high-performance liquid
chromatography and UV detection, Therap Drug Monitor 21, 664.
Harrison, J. E. and Schultz, J. (1976), Studies on the chlorinating activity of
myeloperoxidase, J BIOL CHEM 251, 1371-4.
Hass, U., Dünnbier, U. and Massmann, G. (2012), Occurrence of psychoactive compounds
and their metabolites in groundwater downgradient of a decommissioned sewage
farm in Berlin (Germany), Environ Sci Pollut R 1-11.
Hayhurst, G. P., Harlow, J., Chowdry, J., Gross, E., Hilton, E., Lennard, M. S., Tucker, G.
T. and Ellis, S. W. (2001), Influence of phenylalanine-481 substitutions on the
catalytic activity of cytochrome P450 2D6., Biochem J 355, 373-79.
Heberer, T. (2002), Occurrence, fate, and removal of pharmaceutical residues in the
aquatic environment: a review of recent research data, Toxicol Lett 131, 5-17.
Hennessy, D. J. (1965), Carbamate insecticides, hydride-transferring ability of
methylenedioxybenzenes as a basis of synergistic activity, J Agricul Food Chem 13,
218-20.
Hernandez, A. and Ruiz, M. T. (1998), An EXCEL template for calculation of enzyme
kinetic parameters by non-linear regression, Bioinformatics 14, 227-28.
Hjelmeland, L. M., Aronow, L. and Trudell, J. R. (1977), Intramolecular determination of
substituent effects in hydroxylations catalyzed by cytochrome P-450, Mol
Pharmacol 13, 634-9.
Hoffmann, K.-J., Gyllenhaal, O. and Vessman, J. (1987), Analysis of α-hydroxy
metabolites of metoprolol in human urine after phosgene/trimethylsilyl
derivatization, Biol. Mass Spectrom. 14, 543-48.
Hoffmann, K. J. (1986), Identification of the main urinary metabolites of omeprazole after
an oral dose to rats and dogs, Drug Metab Dispos 14, 341-48.
Hofrichter, M., Scheibner, K., Schneegaß, I. and Fritsche, W. (1998), Enzymatic
combustion of aromatic and aliphatic compounds by manganese peroxidase from
Nematoloma frowardii, Appl Environ Microbiol 64, 399-404.
Hofrichter, M. and Ullrich, R. (2006), Heme-thiolate haloperoxidases: versatile
biocatalysts with biotechnological and environmental significance, Appl Microbiol
Biot 71, 276-88.
Hofrichter, M. and Ullrich, R. (2010), New trends in fungal biooxidation. Industrial
Applications, in: Mycota X, Vol. 10, pp. 425-49 (Esser, K., Ed.) Springer Berlin
Heidelberg,
Hofrichter, M., Ullrich, R., Pecyna, M., Liers, C. and Lundell, T. (2010), New and classic
families of secreted fungal heme peroxidases, Appl Microbiol Biot 87, 871-97.
Holmes, E.et al. (2007), Detection of urinary drug metabolite (xenometabolome)
signatures in molecular epidemiology studies via statistical total correlation (NMR)
spectroscopy, Anal. Chem. 79, 2629-40.
Hrycay, E. G. and Bandiera, S. M. (2012), The monooxygenase, peroxidase, and
peroxygenase properties of cytochrome P450, Arch Biochem Biophys 522, 71-89.
Hu, S. and Hager, L. P. (1999), Asymmetric epoxidation of functionalized cis-olefins
catalyzed by chloroperoxidase, Tetrahedron Lett 40, 1641-44.
Hyland, R., Roe, E. G. H., Jones, B. C. and Smith, D. A. (2001), Identification of the
cytochrome P450 enzymes involved in the N-demethylation of sildenafil, Br J Clin
Pharmacol 51, 239-48.

REFERENCES
Ichinose, H., Wariishi, H. and Tanaka, H. (2002), Identification and characterization of
novel cytochrome P450 genes from the white-rot basidiomycete Coriolus
versicolor, Appl Microbiol Biot 58, 97-105.
Iida, T., Sumita, T., Ohta, A. and Takagi, M. (2000), The cytochrome P450ALK multigene
family of an n-alkane-assimilating yeast, Yarrowia lipolytica: cloning and
characterization of genes coding for new CYP52 family members, Yeast 16, 1077-
87.
Imai, Y., Matsunaga, I., Kusunose, E. and Ichihara, K. (2000), Unique heme environment
at the putative distal region of hydrogen peroxide-dependent fatty acid -
hydroxylase from Sphingomonas paucimobilis (peroxygenase P450 SPα ), J Biochem
128, 189-94.
Isin, E. M. and Guengerich, F. P. (2007), Complex reactions catalyzed by cytochrome
P450 enzymes, BBA - Gen Subjects 1770, 314-29.
Jacqz-Aigrain, E., Funck-Brentano, C. and Cresteil, T. (1993), CYP2D6- and CYP3Adependent
metabolism of dextromethorphan in humans, Pharmacogenetics 3, 197-
204.
Jin, Y.et al. (2005), CYP2D6 Genotype, antidepressant use, and tamoxifen metabolism
during adjuvant breast cancer treatment, J Natl Cancer Inst 97, 30-39.
Johansson, T., Weidolf, L. and Jurva, U. (2007), Mimicry of phase I drug metabolism –
novel methods for metabolite characterization and synthesis, Rapid Commun Mass
Sp 21, 2323-31.
Joo, H., Lin, Z. and Arnold, F. H. (1999), Laboratory evolution of peroxide-mediated
cytochrome P450 hydroxylation, Nature 399, 670-73.
Joseph-Horne, T. and Hollomon, D. W. (1997), Molecular mechanisms of azole resistance
in fungi, FEMS Microbiol Lett 149, 141-49.
Julsing, M. K., Cornelissen, S., Bühler, B. and Schmid, A. (2008), Heme-iron oxygenases:
powerful industrial biocatalysts?, Curr Opin Chem Biol 12, 177-86.
Jung, S. T., Lauchli, R. and Arnold, F. H. Cytochrome P450: taming a wild type enzyme,
Curr Opin Biotech 22, 809-17.
Jux, U., Baginski, R. M., Arnold, H.-G., Krönke, M. and Seng, P. N. (2002), Detection of
pharmaceutical contaminations of river, pond, and tap water from Cologne
(Germany) and surroundings, Int J Hyg Envir Heal 205, 393-98.
Karlson, U., Dwyer, D. F., Hooper, S. W., Moore, E. R., Timmis, K. N. and Eltis, L. D.
(1993), Two independently regulated cytochromes P-450 in a Rhodococcus
rhodochrous strain that degrades 2-ethoxyphenol and 4-methoxybenzoate, J
Bacteriol 175, 1467-74.
Kasai, N.et al. Metabolism of mono- and dichloro-dibenzo-p-dioxins by Phanerochaete
chrysosporium cytochromes P450, Appl Microbiol Biot 86, 773-80.
Katagiri, M., Ganguli, B. N. and Gunsalus, I. C. (1968), A soluble cytochrome P-450
functional in methylene hydroxylation, J BIOL CHEM 243, 3543-46.
Kellermann, G. H. and Luyten-Kellermann, M. (1979), On the regulation of antipyrine and
oxazepam metabolism in man., Res Commun Chem Pathol Pharmacol 23, 287-96.
Kellner, D. G., Hung, S.-C., Weiss, K. E. and Sligar, S. G. (2002), Kinetic characterization
of compound I formation in the thermostable cytochrome P450 CYP119, J BIOL
CHEM 277, 9641-44.
Kelly, S., Kelly, D., Jackson, C., Warrilow, A., Lamb, D. and Ortiz de Montellano, P. R.
(2005), The Diversity and importance of microbial cytochromes P450, in:
Cytochrome P450: Structure, Mechanism, and Biochemistry, Vol. 585-617
(Springer US,
Khan, M. M. T. and Martell, A. E. (1967), Metal ion and metal chelate catalyzed oxidation
of ascorbic acid by molecular oxygen. I. Cupric and ferric ion catalyzed oxidation,
J Am Chem Soc 89, 4176-85.
131

REFERENCES
Kilpatrick, G. J. and Smith, T. W. (2005), Morphine-6-glucuronide: Actions and
mechanisms, Med Res Rev 25, 521-44.
Kinne, M. (2010), The extracellular peroxygenase of the agaric fungus Agrocybe aegerita:
catalytic properties and physiological background with particular emphasis on
ether cleavage, Qucosa, Zittau.
Kinne, M., Poraj-Kobielska, M., Aranda, E., Ullrich, R., Hammel, K. E., Scheibner, K. and
Hofrichter, M. (2009a), Regioselective preparation of 5-hydroxypropranolol and
4'-hydroxydiclofenac with a fungal peroxygenase, Bioorg Med Chem Lett 19,
3085-87.
Kinne, M., Poraj-Kobielska, M., Ralph, S. A., Ullrich, R., Hofrichter, M. and Hammel, K.
E. (2009b), Oxidative cleavage of diverse ethers by an extracellular fungal
peroxygenase, J BIOL CHEM 284, 29343-49.
Kinne, M., Ullrich, R., Hammel, K. E., Scheibner, K. and Hofrichter, M. (2008),
Regioselective preparation of (R)-2-(4-hydroxyphenoxy)propionic acid with a
fungal peroxygenase, Tetrahedron Lett 49, 5950-53.
Kinne, M., Zeisig, C., Ullrich, R., Kayser, G., Hammel, K. E. and Hofrichter, M. (2010),
Stepwise oxygenations of toluene and 4-nitrotoluene by a fungal peroxygenase,
Biochem. Biophys. Res. Commun. 397, 18-21.
Klebanoff, S. J. (1999), Myeloperoxidase, P Assoc Am Physician 111, 383-89.
Klebanoff, S. J. (2005), Myeloperoxidase: friend and foe, J Leukocyte Biol 77, 598-625.
Kluge, M., Ullrich, R., Dolge, C., Scheibner, K. and Hofrichter, M. (2009), Hydroxylation
of naphthalene by aromatic peroxygenase from Agrocybe aegerita proceeds via
oxygen transfer from H 2 O 2 and intermediary epoxidation, Appl. Microbiol.
Biotechnol. 81, 1071-76.
Kluge, M., Ullrich, R., Scheibner, K. and Hofrichter, M. (2007), Spectrophotometric assay
for detection of aromatic hydroxylation catalyzed by fungal haloperoxidase–
peroxygenase, Appl Microbiol Biot 75, 1473-78.
Kluge, M., Ullrich, R., Scheibner, K. and Hofrichter, M. (2012), Stereoselective benzylic
hydroxylation of alkylbenzenes and epoxidation of styrene derivatives catalyzed by
the peroxygenase of Agrocybe aegerita, Green Chem 14, 440-46.
Ku, H. Y.et al. (2008), The contributions of cytochromes P450 3A4 and 3A5 to the
metabolism of the phosphodiesterase type 5 inhibitors sildenafil, udenafil, and
vardenafil, Drug Metab Dispos 36, 986-90.
Lah, L.et al. The versatility of the fungal cytochrome P450 monooxygenase system is
instrumental in xenobiotic detoxification, Mol Microbiol 81, 1374-89.
Law, M. Y., Charles, S. A. and Halliwell, B. (1983), Glutathione and ascorbic acid in
spinach (Spinacia oleracea) chloroplasts. The effect of hydrogen peroxide and of
Paraquat., Biochem J 210 899–903.
Lee, D.-S.et al. (2003), Substrate recognition and molecular mechanism of fatty acid
hydroxylation by cytochrome P450 from Bacillus subtilis, J BIOL CHEM 278,
9761-67.
Lee, S. H.et al. (2010), Decreased detoxification genes and genome size make the human
body louse an efficient model to study xenobiotic metabolism, Insect Mol Biol 19,
599-615.
Leemann, T., Transon, C. and Dayer, P. (1993), Cytochrome P450TB (CYP2C): A major
monooxygenase catalyzing diclofenac 4'-hydroxylation in human liver, Life
Sciences 52, 29-34.
Lemberger, L. (1973), Tetrahydrocannabinol metabolism in man, Drug Metab Dispos 1,
461-68.
Lequeu, J., Fauconnier, M.-L., Chammaï, A., Bronner, R. and Blée, E. (2003), Formation
of plant cuticle: evidence for the occurrence of the peroxygenase pathway, Plant J
36, 155-64.
132

REFERENCES
Lewis, D. F. V. (1996), Cytochromes P450: structure, function and mechanism., Taylor &
Francis, London.
Lewis, D. F. V. (2001), Guide to Cytochromes P450. Structure and Function., Taylor &
Francis, London and New York.
Lewis, D. F. V. and Wiseman, A. (2005), A selective review of bacterial forms of
cytochrome P450 enzymes, Enzyme Microb Tech 36, 377-84.
Li, A. P., Lu, C., Brent, J. A., Pham, C., Fackett, A., Ruegg, C. E. and Silber, P. M. (1999),
Cryopreserved human hepatocytes: characterization of drug-metabolizing activities
and applications in higher throughput screening assays for hepatotoxicity,
metabolic stability, and drug–drug interaction potential, Chem-Biol Interact 121,
17-35.
Li, X., Schuler, M. A. and Berenbaum, M. R. (2007), Molecular Mechanisms of Metabolic
Resistance to Synthetic and Natural Xenobiotics, Annu Rev Entomol 52, 231-53.
Lienert, J., Güdel, K. and Escher, B. I. (2007), Screening Method for Ecotoxicological
Hazard Assessment of 42 Pharmaceuticals Considering Human Metabolism and
Excretory Routes, Environ Sci Technol 41, 4471-78.
Lindegardh, N., Davies, G. R., Hien, T. T., Farrar, J., Singhasivanon, P., Day, N. P. J. and
White, N. J. (2006), Rapid degradation of oseltamivir phosphate in clinical samples
by plasma esterases, Antimicrob Agents Ch 50, 3197-99.
Liu, G., Gelboin, H. V. and Myers, M. J. (1991), Role of cytochrome P450 IA2 in
acetanilide 4-hydroxylation as determined with cDNA expression and monoclonal
antibodies, Arch Biochem Biophys 284, 400-6.
Lu, A. Y. H. and Coon, M. J. (1968), Role of Hemoprotein P-450 in Fatty Acid ω-
Hydroxylation in a Soluble Enzyme System from Liver Microsomes J BIOL CHEM
243, 1331-32.
MacKeith, R. A., McCague, R., Olivo, H. F., Roberts, S. M., Taylor, S. J. C. and Xiong, H.
(1994), Enzyme-catalysed kinetic resolution of 4-endo-hydroxy-2-
oxabicyclo[3.3.0]oct-7-en-3-one and employment of the pure enantiomers for the
synthesis of anti-viral and hypocholestemic agents, Bioorg Med Chem 2, 387-94.
Mahato, S. B. and Majumdar, I. (1993), Current trends in microbial steroid
biotransformation, Phytochemistry 34, 883-98.
Malonek, S., Rojas, M. C., Hedden, P., Gaskin, P., Hopkins, P. and Tudzynski, B. (2005),
Functional Characterization of Two Cytochrome P450 Monooxygenase Genes,
P450-1 and P450-4, of the Gibberellic Acid Gene Cluster in Fusarium proliferatum
(Gibberella fujikuroi MP-D), Appl Environ Microbiol 71, 1462-72.
Maloney, A. P. and VanEtten, H. D. (1994), A gene from the fungal plant pathogen Nectria
haematococca that encodes the phytoalexin-detoxifying enzyme pisatin demethylase
defines a new cytochrome P450 family, Mol Gen Genet 243, 506-14.
Mansuy, D. (1998), The great diversity of reactions catalyzed by cytochromes P450, Comp
Biochem Phys C 121, 5-14.
Marco-Urrea, E., Perez-Trujillo, M., Cruz-Morato, C., Caminal, G. and Vicent, T. (2010a),
Degradation of the drug sodium diclofenac by Trametes versicolor pellets and
identification of some intermediates by NMR, J Hazard Mater 176, 836-42.
Marco-Urrea, E., Pérez-Trujillo, M., Cruz-Morató, C., Caminal, G. and Vicent, T. (2010b),
White-rot fungus-mediated degradation of the analgesic ketoprofen and
identification of intermediates by HPLC–DAD–MS and NMR, Chemosphere 78,
474-81.
Marco-Urrea, E., Perez-Trujillo, M., Vicent, T. and Caminal, G. (2009), Ability of whiterot
fungi to remove selected pharmaceuticals and identification of degradation
products of ibuprofen by Trametes versicolor, Chemosphere 74, 765-72.
Marco-Urrea, E., Radjenovic, J., Caminal, G., Petrovic, M., Vicent, T. and Barcelo, D.
(2010c), Oxidation of atenolol, propranolol, carbamazepine and clofibric acid by a
133

REFERENCES
biological Fenton-like system mediated by the white-rot fungus Trametes
versicolor, Water Res 44, 521-32.
Martinez Bueno, M. J., Agüera, A., Gómez, M. J., Hernando, M. D., García-Reyes, J. F.
and Fernández-Alba, A. R. (2007), Application of Liquid
Chromatography/Quadrupole-Linear Ion Trap Mass Spectrometry and Time-of-
Flight Mass Spectrometry to the Determination of Pharmaceuticals and Related
Contaminants in Wastewater, Anal Chem 79, 9372-84.
Martinović, D., Hogarth, W. T., Jones, R. E. and Sorensen, P. W. (2007), Environmental
estrogens suppress hormones, behavior, and reproductive fitness in male fathead
minnows, Environ Toxicol Chem 26, 271-78.
Masaphy, S., Levanon, D., Henis, Y., Venkateswarlu, K. and Kelly, S. L. (1995),
Microsomal and cytosolic cytochrome P450 mediated benzo(a)pyrene
hydroxylation in Pleurotus pulmonarius, Biotechnol Lett 17, 969-74.
Maspahy, S., Lamb, D. C. and Kelly, S. L. (1999), Purification and Characterization of a
Benzo[a]pyrene Hydroxylase from Pleurotus pulmonarius, Biochem Biophys Res
Co 266, 326-29.
Masubuchi, Y., Hosokawa, S., Horie, T., Suzuki, T., Ohmori, S., Kitada, M. and
Narimatsu, S. (1994), Cytochrome P450 isozymes involved in propranolol
metabolism in human liver microsomes. The role of CYP2D6 as ring-hydroxylase
and CYP1A2 as N-desisopropylase, Drug Metab Dispos 22, 909-15.
Matsuzaki, F. and Wariishi, H. (2004), Functional diversity of cytochrome P450s of the
white-rot fungus Phanerochaete chrysosporium, Biochem Bioph Res Co 324, 387-
93.
Matsuzaki, F. and Wariishi, H. (2005), Molecular characterization of cytochrome P450
catalyzing hydroxylation of benzoates from the white-rot fungus Phanerochaete
chrysosporium, Biochem Bioph Res Co 334, 1184-90.
Maurel, P. (1996), The use of adult human hepatocytes in primary culture and other in
vitro systems to investigate drug metabolism in man, Adv Drug Deliv Rev 22, 105-
32.
McClellan, K. and Perry, C. M. (2001), Oseltamivir: A review of its use in influenza, Drugs
61, 263-83.
Meunier, B., de Visser, S. P. and Shaik, S. (2004), Mechanism of Oxidation Reactions
Catalyzed by Cytochrome P450 Enzymes, Chem Rev 104, 3947-80.
Meyer, U. A. and Zanger, U. M. (1997), Molecular mechanisms of genetic polymorphisms
of drug metabolism, Ann Rev Pharmacol 37, 269-96.
Miners, J. O. and Birkett, D. J. (1996), Use of tolbutamide as a substrate probe for human
hepatic cytochrome P450 2C9, Methods Enzymol 272, 139-45.
Miners, J. O., Coulter, S., Tukey, R. H., Veronese, M. E. and Birkett, D. J. (1996),
Cytochromes P450, 1A2, and 2C9 are responsible for the human hepatic O-
demethylation of R- and S-naproxen, Biochem Pharmacol 51, 1003-8.
Miyamoto, G., Zahid, N. and Uetrecht, J. P. (1997), Oxidation of Diclofenac to Reactive
Intermediates by Neutrophils, Myeloperoxidase, and Hypochlorous Acid, CHEM
RES TOXICOL 10, 414-19.
Mompelat, S., Le Bot, B. and Thomas, O. (2009), Occurrence and fate of pharmaceutical
products and by-products, from resource to drinking water, Environ Int 35, 803-14.
Muirhead, G. J., Rance, D. J., Walker, D. K. and Wastall, P. (2002), Comparative human
pharmacokinetics and metabolism of single-dose oral and intravenous sildenafil,
Brit J Clin Pharmacol 53, 13S-20S.
Munro, A. W., Girvan, H. M. and McLean, K. J. (2007), Cytochrome P450 - redox partner
fusion enzymes, BBA - General Subjects 1770, 345-59.
Munro, A. W.et al. (2002), P450 BM3: the very model of a modern flavocytochrome,
Trends Biochem Sci 27, 250-57.
134

REFERENCES
Munro, A. W. and Lindsay, J. G. (1996), Bacterial cytochromes P-450, Mol Microbiol 20,
1115-25.
Murai, T., Iwabuchi, H. and Ikeda, T. (2004), Identification of Gemfibrozil Metabolites,
Produced as Positional Isomers in Human Liver Microsomes, by On-line Analyses
Using Liquid Chromatography/Mass Spectrometry and Liquid
Chromatography/Nuclear Magnetic Resonance Spectroscopy, J Mass Spec Soc Jpn
52, 277-83.
Murray, M. (2000), Mechanisms of Inhibitory and Regulatory Effects of
Methylenedioxyphenyl Compounds on Cytochrome P450-Dependent Drug
Oxidation, Curr Drug Metab 1, 67-84.
Nagababu, E. and Rifkind, J. M. (2004), Heme Degradation by Reactive Oxygen Species
Antioxid Redox Sign 6, 967-78.
Nakayama, N., Takemae, A. and Shoun, H. (1996), Cytochrome P450foxy, a Catalytically
Self-Sufficient Fatty Acid Hydroxylase of the Fungus Fusarium oxysporum, J
Biochem 119, 435-40.
Narhi, L. O. and Fulco, A. J. (1987), Identification and characterization of two functional
domains in cytochrome P-450BM-3, a catalytically self-sufficient monooxygenase
induced by barbiturates in Bacillus megaterium, J BIOL CHEM 262, 6683-90.
Nebert, D. W. and Gonzalez, F. J. (1987), P450 Genes: Structure, Evolution, and
Regulation, Ann Rev Biochem 56, 945-93.
Nelson, D. (2009), The Cytochrome P450 Homepage, Human Genomics 4 59-65.
Nelson, D. R. (2011), Progress in tracing the evolutionary paths of cytochrome P450,
BBA - Proteins Proteomics 1814, 14-18.
Nelson, D. R.et al. The P450 superfamily: update on new sequences, gene mapping,
accession numbers, early trivial names of enzymes, and nomenclature, DNA Cell
Biol 12, 1-51.
Nelson, D. R.et al. (1996), P450 superfamily: update on new sequences, gene mapping,
accession numbers and nomenclature, Pharmacogenetics 6, 1-42.
Nelson, D. R. and Nebert, D. W. (2011), Cytochrome P450 (CYP) gene superfamily, in:
eLS, Vol. John Wiley & Sons, Ltd,
Nelson, S. D. (1982), Metabolic activation and drug toxicity, Journal of Medicinal
Chemistry 25, 753-65.
Nelson, S. D. and Trager, W. F. (2003), The use of deuterium isotope effects to probe the
active site properties, mechanism of cytochrome P450-catalyzed reactions, and
mechanisms of metabolically dependent toxicity, Drug Metab. Dispos. 31, 1481-98.
Nielsen, K., Møller, B. and Ortiz de Montellano, P. R. (2005), Cytochrome P450s in Plants
Cytochrome P450, 553-83 (Springer US,
Nomenclature Committee of the International Union of Biochemistry and Molecular
Biology (NC-IUBMB) (2012), Enzyme Nomenclature,
http://www.chem.qmul.ac.uk/iubmb/enzyme/
Nottebaum, L. J. and McClure, T. D. (2010), Methods, systems, and computer program
products for producing theoretical mass spectral fragmentation patterns of
chemical structures, in: ip.com, Vol. Syngenta Participations AG, USA.
Oaks, J. L.et al. (2004), Diclofenac residues as the cause of vulture population decline in
Pakistan, Nature 427, 630-33.
Oatis, J. E., Jr., Russell, M. P., Knapp, D. R. and Walle, T. (1981), Ring-hydroxylated
propranolol: synthesis and beta-receptor antagonist and vasodilating activities of
the seven isomers, J Med Chem 24, 309-14.
Ogliaro, F., de Visser, S. P., Cohen, S., Sharma, P. K. and Shaik, S. (2002), Searching for
the second oxidant in the catalytic cycle of cytochrome P450: a theoretical
investigation of the iron(III)-hydroperoxo species and its epoxidation pathways, J
Am Chem Soc 124, 2806-17.
135

136
REFERENCES
Ohkuma, M., Zimmer, T., Iida, T., Schunck, W.-H., Ohta, A. and Takagi, M. (1998),
Isozyme Function of n-Alkane-inducible Cytochromes P450 in Candida maltosa
Revealed by Sequential Gene Disruption, J BIOL CHEM 273, 3948-53.
Omura, T. (2005), Heme-thiolate proteins, Biochem. Biophys. Res. Commun. 338, 404-09.
Omura, T., Sanders, E., Estabrook, R. W., Cooper, D. Y. and Rosenthal, O. (1966),
Isolation from adrenal cortex of a nonheme iron protein and a flavoprotein
functional as a reduced triphosphopyridine nucleotide-cytochrome P-450
reductase, Arch Biochem Biophys 117, 660-73.
Omura, T. and Sato, R. (1964), The Carbon Monoxide-binding Pigment of Liver
Microsomes, J BIOL CHEM 239, 2370-78.
Ophardt, E. C. (2003), Amino acids, in: Virtual ChemBook, Vol. Elmhurst College.
Orlando, R., Piccoli, P., De Martin, S., Padrini, R., Floreani, M. and Palatini, P. (2004),
Cytochrome P450 1A2 is a major determinant of lidocaine metabolism in vivo:
effects of liver function, Clin Pharmacol Ther 75, 80-88.
Ortiz de Montellano, P. (2005), Cytochrome P450 - Structure, Mechanism and
Biochemistry, Ortiz de Montellano, P., Ed.) Kluwer Academic/Plenum Publishers,
New York.
Ortiz de Montellano, P., Voss, J. and Ortiz de Montellano, P. R. (2005), Substrate
Oxidation by Cytochrome P450 Enzymes
Cytochrome P450, 183-245 (Springer US,
Ortiz de Montellano, P. R. and de Voss, J. J. (2005), Substrate oxidation by cytochrome
P450 enzymes, in: Cytochrome P450 - Structure, Mechanism and Biochemistry,
Vol. 183-245 (Ortiz De Montellano, P. R., Ed.) Kluwer Academic/Plenum
Publishers, New York.
Osman, A. M., Koerts, J., Boersma, M. G., Boeren, S., Veeger, C. and Rietjens, I. M.
(1996), Microperoxidase/H2O2-catalyzed aromatic hydroxylation proceeds by a
cytochrome-P-450-type oxygen-transfer reaction mechanism, Eur J Biochem 240,
232-8.
Otey, C. R., Bandara, G., Lalonde, J., Takahashi, K. and Arnold, F. H. (2006), Preparation
of human metabolites of propranolol using laboratory-evolved bacterial
cytochromes P450, Biotechnol Bioeng 93, 494-9.
Paice, M. G., Reid, I. D., Bourbonnais, R., Archibald, F. S. and Jurasek, L. (1993),
Manganese peroxidase, produced by Trametes versicolor during pulp bleaching,
demethylates and delignifies kraft pulp, Appl Environ Microbiol 59, 260-65.
Parliment, E., The European Parliment and Council, E., The Council Of The European
Union (2004), Directive 2004/27/EC of the European Parliament and of the
Council of 31 March 2004 amending directive 2001/83/EC on the community code
relating to medicinal products for human use, 136, pp. 0034 - 57 (Official J,
Pearce, R. E., Vakkalagadda, G. R. and Leeder, J. S. (2002), Pathways of carbamazepine
bioactivation in vitro I. Characterization of human cytochromes P450 responsible
for the formation of 2- and 3-hydroxylated metabolites, Drug Metab Dispos 30,
1170-9.
Pecyna, M. (2012), Gen sequences of fungal peroxygenases, oral communication,
International Graduate School Zittau, oral communication,
Pecyna, M. J., Ullrich, R., Bittner, B., Clemens, A., Scheibner, K., Schubert, R. and
Hofrichter, M. (2009), Molecular characterization of aromatic peroxygenase from
Agrocybe aegerita, Appl Microbiol Biotechnol 84, 885-97.
Perry, T. L., Culling, C. F. A., Berry, K. and Hansen, S. (1964), 7-
Hydroxychlorpromazine: Potential Toxic Drug Metabolite in Psychiatric Patients,
Science 146, 81-83.
Peter, S., Kinne, M., Wang, X., Ullrich, R., Kayser, G., Groves, J. T. and Hofrichter, M.
(2011), Selective hydroxylation of alkanes by an extracellular fungal peroxygenase,
FEBS J 278, 3667-75.

REFERENCES
Peters, F. T., Dragan, C. A., Wilde, D. R., Meyer, M. R., Zapp, J., Bureik, M. and Maurer,
H. H. (2007), Biotechnological synthesis of drug metabolites using human
cytochrome P450 2D6 heterologously expressed in fission yeast exemplified for the
designer drug metabolite 4'-hydroxymethyl-alpha-pyrrolidinobutyrophenone,
Biochem Pharmacol 74, 511-20.
Petrović, M., Gonzalez, S. and Barceló, D. (2003), Analysis and removal of emerging
contaminants in wastewater and drinking water, TrAC-Trend Anal Chem 22, 685-
96.
Philpot, R. M. and Hodgson, E. (1972), The production and modification of cytochrome P-
450 difference spectra by in vivo administration of methylenedioxyphenyl
compounds, Chem-Biol Interact 4, 185-94.
Piontek, K., Ullrich, R., Liers, C., Diederichs, K., Plattner, D. A. and Hofrichter, M.
(2010), Crystallization of a 45 kDa peroxygenase/peroxidase from the mushroom
Agrocybe aegerita and structure determination by SAD utilizing only the haem
iron, Acta Crystallogr Sect F Struct Biol Cryst Commun 66, 693-8.
Piontek, K.et al. (2012), Crystal structures of aromatic peroxygenase: substrate-binding
mode in a new class of heme redox enzymes,
Poraj-Kobielska, M., Kinne, M., Ullrich, R., Scheibner, K. and Hofrichter, M. (2011a), A
spectrophotometric assay for the detection of fungal peroxygenases, Anal Bioch
421, 327-29.
Poraj-Kobielska, M., Kinne, M., Ullrich, R., Scheibner, K., Kayser, G., Hammel, K. E. and
Hofrichter, M. (2011b), Preparation of human drug metabolites using fungal
peroxygenases, Biochem Pharmacol 82, 789-96.
Prieto, T., Marcon, R. O., Prado, F. M., Caires, A. C., Di Mascio, P., Brochsztain, S.,
Nascimento, O. R. and Nantes, I. L. (2006), Reaction route control by
microperoxidase-9/CTAB micelle ratios, Phys. Chem. Chem. Phys. 8, 1963-73.
Quertemont, E. (2004), Genetic polymorphism in ethanol metabolism: acetaldehyde
contribution to alcohol abuse and alcoholism, Mol Psychiatry 9, 570-81.
Quinn, B., Gagne, F. and Blaise, C. (2008), An investigation into the acute and chronic
toxicity of eleven pharmaceuticals (and their solvents) found in wastewater effluent
on the cnidarian, Hydra attenuata, Sci Total Environ 389, 306-14.
Rittle, J. and Green, M. T. (2010), Cytochrome P450 compound I: capture,
characterization, and C-H bond activation kinetics, Science 330, 933-7.
Rodarte-Morales, A., Feijoo, G., Moreira, M. and Lema, J. (2012), Biotransformation of
three pharmaceutical active compounds by the fungus Phanerochaete
chrysosporium in a fed batch stirred reactor under air and oxygen supply,
Biodegradation 23, 145-56.
Roy, A., Kucukural, A. and Zhang, Y. (2010), I-TASSER: a unified platform for automated
protein structure and function prediction, Nat Protocol 5, 725-38.
Ruiz-Garcia, A., Bermejo, M., Moss, A. and Casabo, V. G. (2008), Pharmacokinetics in
drug discovery, J Pharm Sci 97, 654-90.
Sachverständigen Rat für Umweltfragen (2007), Arzneimittel in der Umwelt, 1-95.
Saikia, S., Parker, E. J., Koulman, A. and Scott, B. (2007), Defining Paxilline Biosynthesis
in Penicillium paxilli, J BIOL CHEM 282, 16829-37.
Sawayama, A. M., Chen, M. M., Kulanthaivel, P., Kuo, M. S., Hemmerle, H. and Arnold,
F. H. (2009), A panel of cytochrome P450 BM3 variants to produce drug
metabolites and diversify lead compounds, Chem Eur J 15, 11723-9.
Scheibner, K. and Hofrichter, M. (1998), Conversion of aminonitrotoluenes by fungal
manganese peroxidase, Journal of Basic Microbiology 38, 51-59.
Scheller, U., Zimmer, T., Kärgel, E. and Schunck, W.-H. (1996), Characterization of then-
Alkane and Fatty Acid Hydroxylating Cytochrome P450 Forms 52A3 and 52A4,
Arch Biochem Biophys 328, 245-54.
137

138
REFERENCES
Schenkman, J. B., Cinti, D. L., Orrenius, S., Moldeus, P. and Kraschnitz, R. (1972), The
nature of the reverse type I (modified type II) spectral change in liver microsomes,
Biochemistry 11, 4243-51.
Schenkman, J. B., Sligar, S. G. and Cinti, D. L. (1981), Substrate interaction with
cytochrome P-450, Pharmacol Therapeut 12, 43-71.
Schmidt, H. W. H., Haemmerli, S. D., Schoemaker, H. E. and Leisola, M. S. A. (1989),
Oxidative degradation of 3,4-dimethoxybenzyl alcohol and its methyl ether by the
lignin peroxidase of Phanerochaete chrysosporium, Biochemistry 28, 1776-83.
Schroer, K., Kittelmann, M. and Lütz, S. (2010), Recombinant human cytochrome P450
monooxygenases for drug metabolite synthesis, Biotechnol Bioeng 106, 699-706.
Schuler, M. A. (1996a), Plant Cytochrome P450 Monooxygenases, CRC Cr Rev Plant Sci
15, 235-84.
Schuler, M. A. (1996b), The Role of Cytochrome P450 Monooxygenases in Plant-Insect
Interactions, Plant Physiol 112, 1411-19.
Schwarz, H. J. (1979), Pharmacokinetics and metabolism of temazepam in man and
several animal species, Br J Clin Pharmacol 8, 23S–29S.
Scott, J. G. (2008), Insect cytochrome P450s: Thinking beyond detoxification, Recent
Advances in Insect Physiology, Toxicology and Molecular Biology ISBN: 978-81-
308-0242-8, 117-24.
Segel, H. I. (1994), Enzyme Kinetics: Behavior and Analysis of Rapid Equilibrium and
Steady-State Enzyme Systems, John Wiley Sons, Inc., New York
Segre, E. J. (1975), Naproxen metabolism in man, J Clin Pharmacol 15, 316-23.
Shoji, O., Fujishiro, T., Nakajima, H., Kim, M., Nagano, S., Shiro, Y. and Watanabe, Y.
(2007), Hydrogen peroxide dependent monooxygenations by tricking the substrate
recognition of cytochrome P450BSβ, Angew Chem Int Ed 46, 3656-59.
Shoji, O., Wiese, C., Fujishiro, T., Shirataki, C., Wünsch, B. and Watanabe, Y. (2010),
Aromatic C–H bond hydroxylation by P450 peroxygenases: a facile colorimetric
assay for monooxygenation activities of enzymes based on Russig’s blue formation,
J Biol Inorg Chem 15, 1109-15.
Shoun, H., Fushinobu, S., Jiang, L., Kim, S.-W. and Wakagi, T. Fungal denitrification and
nitric oxide reductase cytochrome P450nor, Philos T Roy Soc B 367, 1186-94.
Shoun, H. and Tanimoto, T. (1991), Denitrification by the fungus Fusarium oxysporum
and involvement of cytochrome P-450 in the respiratory nitrite reduction, J BIOL
CHEM 266, 11078-82.
Simonneaux, G., Le Maux, P. and Simonneaux, G. (2006), Carbene Complexes of Heme
Proteins and Iron Porphyrin Models
Bioorganometallic Chemistry, 17, pp. 83-122 (Springer Berlin / Heidelberg,
Skordi, E., Wilson, I. D., Lindon, J. C. and Nicholson, J. K. (2005), Kinetic studies on the
intramolecular acyl migration of β-1-O-acyl glucuronides: Application to the
glucuronides of (R)- and (S)-ketoprofen, (R)- and (S)-hydroxy-ketoprofen
metabolites, and tolmetin by 1H-NMR spectroscopy, Xenobiotica 35, 715-25.
Sligar, S. G. and Gunsalus, I. C. (1976), A thermodynamic model of regulation: modulation
of redox equilibria in camphor monoxygenase, P Natl Acad Sci USA 73, 1078-82.
Sono, M., Roach, M. P., Coulter, E. D. and Dawson, J. H. (1996), Heme-containing
oxygenases, Chem Rev 96, 2841-88.
Souček, P. and Gut, I. (1992), Cytochromes P-450 in rats: structures, functions, properties
and relevant human forms, Xenobiotica 22, 83-103.
Stamets, P. (1993), Growing gourmet and medicinal mushrooms: a companion guide to
The mushroom cultivator, Ten Speed Press (Berkeley, CA) Berleöey.
Stemmer, W. P. C. (1994), Rapid evolution of a protein in vitro by DNA shuffling, Nature
370, 389-91.
Strode, C.et al. (2008), Genomic analysis of detoxification genes in the mosquito Aedes
aegypti, Insect Biochem Molec 38, 113-23.

REFERENCES
Stundl, U. M., Patzak, D. and Schauer, F. (2000), Purification of a soluble cytochrome
P450 from Trichosporon montevideense, J Basic Microbiol 40, 289-92.
Subramanian, V., Doddapaneni, H., Syed, K. and Yadav, J. (2010), P450 Redox Enzymes
in the White Rot Fungus Phanerochaete chrysosporium: Gene Transcription,
Heterologous Expression, and Activity Analysis on the Purified Proteins, Curr
Microbiol 61, 306-14.
Sugano, Y., Muramatsu, R., Ichiyanagi, A., Sato, T. and Shoda, M. (2007), DyP, a unique
dye-decolorizing peroxidase, represents a novel heme peroxidase family: ASP171
replaces the distal histidine of classical peroxidases, J BIOL CHEM 282, 36652-8.
Sundaramoorthy, M., Terner, J. and Poulos, T. L. (1995), The crystal structure of
chloroperoxidase: a heme peroxidase--cytochrome P450 functional hybrid,
Structure 3, 1367-77.
Sundaramoorthy, M. T., J. Poulos, T.L. (1995), The crystal structure of chloroperoxidase:
a heme peroxidase--cytochrome P450 functional hybrid., Structure 3, 1367-77.
Suzuki, K., Sanga, K.-i., Chikaoka, Y. and Itagaki, E. (1993), Purification and properties
of cytochrome P-450 (P-450lun) catalyzing steroid 11β-hydroxylation in
Curvularia lunata, BBA - Protein Struct M 1203, 215-23.
Syed, K., Porollo, A., Lam, Y. W. and Yadav, J. S. (2011), A Fungal P450 (CYP5136A3)
Capable of Oxidizing Polycyclic Aromatic Hydrocarbons and Endocrine
Disrupting Alkylphenols: Role of Trp 129 and Leu 324 , PLoS ONE 6, e28286.
Syed, K. and Yadav, J. S. (2012), P450 monooxygenases (P450ome) of the model white rot
fungus Phanerochaete chrysosporium, Crc Cr Rev Microbiol 38, 339-63.
Sztal, T., Chung, H., Berger, S., Currie, P. D., Batterham, P. and Daborn, P. J. (2012), A
Cytochrome P450 Conserved in Insects Is Involved in Cuticle Formation, PLoS
ONE 7, e36544.
Tam, Y. K., Ke, J., Coutts, R. T., Wyse, D. G. and Gray, M. R. (1990), Quantification of
Three Lidocaine Metabolites and Their Conjugates, Pharm Res 7, 504-07.
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J., Gonzalez, F. J. and Miners, J. O.
(1993a), Validation of 4-nitrophenol as an in vitro substrate probe for human liver
CYP2E1 using cDNA expression and microsomal kinetic techniques, Biochem
Pharmacol 46, 1975-81.
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J. and Miners, J. O. (1993b), Highperformance
liquid chromatographic assay for 4-nitrophenol hydroxylation, a
putative cytochrome P-4502E1 activity, in human liver microsomes, J. Chromatogr.
616, 73-8.
Ternes, T. A., Joss, A. and Siegrist, H. (2004), Peer Reviewed: Scrutinizing
Pharmaceuticals and Personal Care Products in Wastewater Treatment, Environ
Sci Technol 38, 392A-99A.
Thomas, J. L., Duax, W. L., Addlagatta, A., Kacsoh, B., Brandt, S. E. and Norris, W. B.
(2004), Structure/function aspects of human 3beta-hydroxysteroid dehydrogenase,
Mol cell endocrinol 215, 73-82.
Tien, M. and Kirk, T. K. (1984), Lignin-degrading enzyme from Phanerochaete
chrysosporium: Purification, characterization, and catalytic properties of a unique
H 2 O 2 -requiring oxygenase, P Natl Acad Sci USA 81, 2280-84.
Tien, M., Kirk, T. K., Bull, C. and Fee, J. A. (1986), Steady-state and transient-state
kinetic studies on the oxidation of 3,4-dimethoxybenzyl alcohol catalyzed by the
ligninase of Phanerocheate chrysosporium Burds, J. Biol. Chem. 261, 1687-93.
Topham, J. C. (1970), Relationship between difference spectra and metabolism:
Barbiturates, drug interaction and species difference, Biochem Pharmacol 19,
1695-701.
Tucker, G. T.et al. (1994), The demethylenation of methylenedioxymethamphetamine
(“ecstasy”) by debrisoquine hydroxylase (CYP2D6), Biochem Pharmacol 47, 1151-
56.
139

140
REFERENCES
Tudzynski, B., Hedden, P., Carrera, E. and Gaskin, P. (2001), The P450-4 Gene of
Gibberella fujikuroi Encodes ent-Kaurene Oxidase in the Gibberellin Biosynthesis
Pathway, Appl Environ Microbiol 67, 3514-22.
Tuynman, A., Spelberg, J. L., Kooter, I. M., Schoemaker, H. E. and Wever, R. (2000),
Enantioselective Epoxidation and Carbon - Carbon Bond Cleavage Catalyzed by
Coprinus cinereus Peroxidase and Myeloperoxidase, J BIOL CHEM 275, 3025-30.
Udenfriend, S., Clark, C. T., Axelrod, J. and Brodie, B. B. (1954), Ascorbic Acid in
Aromatic Hydroxylation, J BIOL CHEM 208, 731-40.
Uljé, K. (2001), Coprinus radians Snowarski, M., Ed.)
Ullrich, R., Dolge, C., Kluge, M. and Hofrichter, M. (2008), Pyridine as novel substrate
for regioselective oxygenation with aromatic peroxygenase from Agrocybe
aegerita, FEBS Lett. 582, 4100-06.
Ullrich, R. and Hofrichter, M. (2005), The haloperoxidase of the agaric fungus Agrocybe
aegerita hydroxylates toluene and naphthalene, FEBS Lett. 579, 6247-50.
Ullrich, R. and Hofrichter, M. (2007), Enzymatic hydroxylation of aromatic compounds,
Cell Mol Life Sci 64, 271-93.
Ullrich, R., Liers, C., Schimpke, S. and Hofrichter, M. (2009), Purification of
homogeneous forms of fungal peroxygenase, Biotech J 4, 1619-26.
Ullrich, R., Nüske, J., Scheibner, K., Spantzel, J. and Hofrichter, M. (2004), Novel
haloperoxidase from the agaric basidiomycete Agrocybe aegerita oxidizes aryl
alcohols and aldehydes, Appl Environ Microbiol 70, 4575-81.
Ullrich, R., Pecyna, M., Kluge, M., Kinne, M., Liers, C., Hofrichter, M. (2008), Some
special reactions of Agrocybe aegerita peroxygenase (AaP). in: 8th International
Peroxidase Symposium, Vol. 48 (Tampere, Finland.
Upthagrove, A. L. and Nelson, W. L. (2001), Importance of amine pKa and distribution
coefficient in the metabolism of fluorinated propranolol derivatives. Preparation,
identification of metabolite regioisomers, and metabolism by CYP2D6, Drug Metab
Dispos 29, 1377-88.
Urlacher, V. B. and Girhard, M. Cytochrome P450 monooxygenases: an update on
perspectives for synthetic application, Trends Biotechnol 30, 26-36.
Urlacher, V. B., Lutz-Wahl, S. and Schmid, R. D. (2004), Microbial P450 enzymes in
biotechnology., Appl Microb Biotechnol 64, 317-25.
Vail, R. B., Homann, M. J., Hanna, I. and Zaks, A. (2005), Preparative synthesis of drug
metabolites using human cytochrome P450s 3A4, 2C9 and 1A2 with NADPH-P450
reductase expressed in &lt;i&gt;Escherichia coli&lt;/i&gt, J Ind Microbiol Biot
32, 67-74.
van den Brink, H. M., van Gorcom, R. F. M., van den Hondel, C. A. M. J. J. and Punt, P. J.
(1998), Cytochrome P450 Enzyme Systems in Fungi, Fungal Genet Biol 23, 1-17.
Veeger, C. (2002), Does P450-type catalysis proceed through a peroxo-iron intermediate?
A review of studies with microperoxidase, J. Inorg. Biochem. 91, 35-45.
Venkatakrishnan, K., Von Moltke, L. L. and Greenblatt, D. J. (2001), Human drug
metabolism and the cytochromes P450: application and relevance of in vitro
models, J Clin Pharmacol 41, 1149-79.
Waldon, D. J., Teffera, Y., Colletti, A. E., Liu, J., Zurcher, D., Copeland, K. W. and Zhao,
Z. (2010), Identification of quinone imine containing glutathione conjugates of
diclofenac in rat bile, Chem ResToxicol 23, 1947-53.
Walker, D. K. (1999), Pharmacokinetics and metabolism of sildenafil in mouse, rat, rabbit,
dog and man, Xenobiotica 29, 297-310.
Walsky, R. L. and Obach, R. S. (2004), Validated Assays for Human Cytochrome P450
activities, Drug Metab Dispos 32, 647-60.
Wang, X., Peter, S., Kinne, M., Hofrichter, M. and Groves, J. T. (2012), Detection and
Kinetic Characterization of a Highly Reactive Heme - Thiolate Peroxygenase
Compound I, J Am Chem Soc 134, 12897-900.

REFERENCES
Warman, A. J.et al. (2005), Flavocytochrome P450 BM3: an update on structure and
mechanism of a biotechnologically important enzyme, Biochem Soc Trans 33, 747-
53.
Warrington, J. S., Shader, R. I., von Moltke, L. L. and Greenblatt, D. J. (2000), In vitro
biotransformation of sildenafil (Viagra): identification of human cytochromes and
potential drug interactions, Drug Metab Dispos 28, 392-97.
Watanabe, K., Yamaori, S., Funahashi, T., Kimura, T. and Yamamoto, I. (2007),
Cytochrome P450 enzymes involved in the metabolism of tetrahydrocannabinols
and cannabinol by human hepatic microsomes, Life Sci 80, 1415-19.
Weis, R., Winkler, M., Schittmayer, M., Kambourakis, S., Vink, M., Rozzell, J. D. and
Glieder, A. (2009), A Diversified Library of Bacterial and Fungal Bifunctional
Cytochrome P450 Enzymes for Drug Metabolite Synthesis, Adv Synth Catal 351,
2140-46.
Werck-Reichhart, D. and Feyereisen, R. (2000), Cytochromes P450: a success story,
Genome Biol 1, reviews3003.1 - reviews03.9.
Wester, M. R., Johnson, E. F., Marques-Soares, C., Dijols, S., Dansette, P. M., Mansuy, D.
and Stout, C. D. (2003), Structure of Mammalian Cytochrome P450 2C5
Complexed with Diclofenac at 2.1 Å Resolution: Evidence for an Induced Fit Model
of Substrate Binding, Biochemistry 42, 9335-45.
Whitacre, D. M., Monteiro, S. C. and Boxall, A. B. A. (2010), Occurrence and fate of
human pharmaceuticals in the environment
reviews of environmental contamination and toxicology, 202, pp. 53-154 (Springer New
York,
WHO Collaborating Centre for Drug Statistics Methodology (2012), The Anatomical
Therapeutic Chemical (ATC),
Wickramasinghe, R. H. and Villee, C. A. (1975), Early role during chemical evolution for
cytochrome P450 in oxygen detoxification, Nature 256, 509-11.
Wiegel, S.et al. (2004), Pharmaceuticals in the river Elbe and its tributaries, Chemosphere
57, 107-26.
Williams, D. G., Patel, A. and Howard, R. F. (2002), Pharmacogenetics of codeine
metabolism in an urban population of children and its implications for analgesic
reliability, Br J Anaesth 89, 839-45.
Williams, P. A., Cosme, J., Sridhar, V., Johnson, E. F. and McRee, D. E. (2000),
Mammalian Microsomal Cytochrome P450 Monooxygenase: Structural
Adaptations for Membrane Binding and Functional Diversity, Mol Cell 5, 121-31.
Winter, H. R. and Unadkat, J. D. (2005), IDENTIFICATION OF CYTOCHROME P450
AND ARYLAMINE N-ACETYLTRANSFERASE ISOFORMS INVOLVED IN
SULFADIAZINE METABOLISM, Drug Metab Dispos 33, 969-76.
Wolfenden, B. S. and Willson, R. L. (1982), Radical-cations as reference chromogens in
kinetic studies of ono-electron transfer reactions: pulse radiolysis studies of 2,2'-
azinobis-(3-ethylbenzthiazoline-6-sulphonate), J Chem Soc, Perkin Transactions 2
805-12.
World Health Organization (2011), WHO Model List of Essential Medicines, 17th list,
World Health Organization (2012), World Health Organization. WHO Model Lists of
Essential Medicines,
Wrighton, S. A. and Stevens, J. C. (1992), The Human Hepatic Cytochromes P450
Involved in Drug Metabolism, CRC Cr Rev Toxicol 22, 1-21.
Yarman, A.et al. (2012), The aromatic peroxygenase from Marasmius rotula—a new
enzyme for biosensor applications, Anal Bioanal Chem 402, 405-12.
Yu, J., Chang, P.-K., Ehrlich, K. C., Cary, J. W., Montalbano, B., Dyer, J. M., Bhatnagar,
D. and Cleveland, T. E. (1998), Characterization of the Critical Amino Acids of an
Aspergillus parasiticus Cytochrome P-450 Monooxygenase Encoded by ordA That
141

REFERENCES
Is Involved in the Biosynthesis of Aflatoxins B1, G1, B2, and G2, Appl Environ
Microbiol 64, 4834-41.
Yun, C. H., Miller, G. P. and Guengerich, F. P. (2000), Rate-determining steps in
phenacetin oxidations by human cytochrome P450 1A2 and selected mutants,
Biochemistry 39, 11319-29.
Zimatkin, S. M. and Deitrich, R. A. (1997), Ethanol metabolism in the brain, Addict Bio 2,
387-400.
142

APPENDIX
6 Appendix
6.1 Aromatic ring hydroxylation
Appendix Figure 1 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of propranolol (0.5 mM) by the aromatic peroxygenases (2 Units mL -1 ) of A.
aegerita (AaeAPO) and C.radians (CraAPO).
Appendix Figure 2 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of diclofenac (0.5 mM) by the aromatic peroxygenase (2 Units mL -1 ) of A.
aegerita. The insets show the UV-Vis-spectra of products and the known structures: (I) -
diclofenac; (II) – 4'-hydroxydiclofenac; (III) – diclofenac-1',4'-quinone imine.
143

APPENDIX
6.2 Oxidation of alkyl side chains
Appendix Figure 3 HPLC-elution profiles of reaction mixtures showing the formation of products
after the conversion of tolbutamide (0.5 mM) by the aromatic peroxygenase (2 U mL -1 ) of A.
aegerita (AaeAPO) with and without ascorbic acid. The insets show mass- and UV-Vis-spectra of
tolbutamide and its products with structures. I – tolbutamide; II – 4-ketotolbutamide; III – 4-
hydroxytolbutamide.
6.3 O-Dealkylation
Appendix Figure 4 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of naproxen (0.5 mM) by the aromatic peroxygenase (2 U mL -1 ) of A. aegerita
(AaeAPO) with ascorbic acid. The insets show the mass spectra of naproxen (II) and O-
desmethylnaproxen (I).
144

APPENDIX
6.4 N-dealkylation
Appendix Figure 5 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of sildenafil (0.5 mM) by AaeAPO, MroAPO and a recombinant peroxygenase
from Novozymes (2 U mL -1 ). The insets show the mass spectra of sildenafil (I), N-
desmethylsildenafil (II) and proposed other products (III) and (IV).
6.5 Oxidation of steroids
Appendix Figure 6 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of pregnenolone (0.5 mM) by the aromatic peroxygenase (2 U mL -1 ) of M. rotula.
The insets show the mass spectra of two products and known structures: (I)–pregnenolone; (II)–
progesterone; (III), (IV) & (V)–not identified products.
145

APPENDIX
6.6 O-Demethylenation
Appendix Figure 7 HPLC-elution
profiles of reaction mixtures showing
the formation of 4-nitrocatechol after
the oxidation of NBD (0.5 mM) by
the aromatic peroxygenases (2 Units
mL -1 ) of C. verticillata (I), C. radians
(II) and A. aegerita (III). The
oxidation of NBD by M. rotula
peroxygenase and recombinant C.
cinerea peroxygenase was also tested
and gave 4-nitrocatechol as major
metabolite as well (data not shown).
Appendix Figure 8 HPLC-elution profiles of reaction mixtures showing the formation of products
after conversion of bromobenzodioxole (0.5 mM) by the aromatic peroxygenase (2 U mL -1 ) of A.
aegerita. The oxidation of chlorobenzodioxole gave a similar elution profile and corresponding
products (data not shown). The insets show mass spectra of products and the known structures: (I)
– bromobenzodioxole; (II) – bromocatechol; (III) – not identified product; (IV) –
trihydroxybromobenzene (bromopyrrogallol).
146

APPENDIX
6.7 Spectrophotometric nitrobenzodioxole assay
Appendix Figure 9 Determination of the
extinction coefficient of 4-nitrocatechol at
pH >12. UV-Vis spectra of different
concentrations of 4-nitocatechol (12.5, 25,
50, 62.5, 100, 125 µM) dissolved in
potassium phosphate/NaOH (50 mM/100
mM) were recorded and the extinction
coefficient calculated for the maximum at
514 nm. The inset shows the linear
relationship between the concentration of 4-
nitrocatechol and the absorbance at 514 nm.
6.8 Stoichiometrical analyses
Appendix Table 1 Stoichiometry of nitrobenzodioxole oxidation by A. aegerita
peroxygenase. 1
H 2 O 2 added catechol produced catechol/H 2 O 2
23.44
46.88
70.3
93.75
187.5
µM
15.1
29.21
39.45
49.87
80.50
0.64 ± 0.01
0.62 ± 0.03
0.56 ± 0.04
0.53 ± 0.01
0.43 ± 0.03
1 The initial nitrobenzodioxole concentration was 0.25 mM.
147

APPENDIX
6.9 Experiment with propranolol at an environmentally relevant
concentrations
Ausgangslösung
1800 ml bi-destiliertes H 2 O
inclusive 2 ml 5 µM Propranolol
+ 900 ml Ausgangslösung
+ 100 ml bi-destiliertes H 2 O
= 1000 ml mit 5 nM Propranolol
1. Ansatz
+ 900 ml Ausgangslösung
+ 50 ml (inclusive 100 Units AaeApo)
+ 50 ml (inclusive 200 µl 30% H 2 O 2
= 1000 ml mit 5nM Propranolol
100 U/l AaeApo und 2 mM H 2 O 2
2. Umsatz
18 h
Inkubationszeit
zur Vergleichbarkeit mit der
Probelösung
3. Ultrafiltration
zur Enzymabtrennung
mit 10 molarer Natronlauge
auf pH 12
4. pH-Wert Einstellung
mit 10 molarer Natronlauge
auf pH 12
mit 25 ml Dichlormethan im
Scheidetrichter
5. Extraktion
mit 25 ml Dichlormethan
im Scheidetrichter
Dichlormethan mittels Turbo-Vap auf
1 ml aufkonzentrieren
6. Aufkonzentration
Dichlormethan mittels Turbo-Vap
auf 1 ml aufkonzentrieren
Dichlormethan mit 2 ml Acetonitril
mischen und auf 1 ml
aufkonzentrieren
7. Extraktion durch
Konzentration
Dichlormethan mit 2 ml Acetonitril
mischen und auf 1 ml
aufkonzentrieren
Acetonitril mit
2 ml bi-destiliertem H 2 O mischen und
auf 1 ml aufkonzentrieren
8. Extraktion durch
Konzentration
Acetonitril mit 2 ml bi-destiliertem
H 2 O mischen und auf 1 ml
aufkonzentrieren
HPLC
„Luna C18“-Säule
9. Analyse
HPLC
„Luna C18“-Säule
9. Vergleich
Appendix Figure 10 Process flow diagram of the experiment with AaeAPO and an
environmentally relevant concentration (5 nM) of propranolol (in German). Figure adapted
from Sylvia Gleissner (Master thesis supervised at the IHI Zittau)
148

APPENDIX
Appendix Figure 11 Examplary 3D-structures (stick models) of pharmaceuticals: sildenafil
(A); diclofenac (B) and dextromethorphan (C).
149

APPENDIX
6.10 List of SCI publications
1. Poraj-Kobielska, M., Kinne, M., Ullrich, R., Scheibner, K. & Hofrichter M. (2012).
A spectrophotometric assay for the detection of fungal peroxygenases. Analytical
Biochemistry, Vol. 421: 327-329. (doi: 10.1016/j.ab.2011.10.009) (impact factor: 3.0)
2. Poraj-Kobielska, M., Kinne, M., Ullrich, R., Scheibner, K., Kayser, G., Hammel, K.
E. and Hofrichter M. (2011). Preparation of human drug metabolites using fungal
peroxygenases. Biochemical Pharmacology, Vol. 82: 789-796 (doi:
10.1016/j.bcp.2011.06.020) (impact factor: 4.7)
3. Kinne, M., Poraj-Kobielska, M., Ullrich, R., Nousiainen, P.,Sipilä, J.,Scheibner, K.,
Hammel, K. E., Hofrichter, M. (2011). Oxidative cleavage of non-phenolic β -O-4
lignin model dimers by an extracellular aromatic peroxygenase. Holzforschung, Vol.
65: 673–679. (impact factor: 1.8)
4. Kinne, M., Poraj-Kobielska, M., Ralph, S. A., Ullrich, R., Hofrichter, M. and
Hammel, K. E. (2009). Oxidative cleavage of diverse ethers by an extracellular fungal
peroxygenase. The Journal of Biological Chemistry, Vol. 284: 29343-29349. (impact
factor: 5.3)
5. Kinne, M. Poraj-Kobielska, M., Aranda, E., Ullrich, R., Hammel, K. E., Scheibner,
K., & Hofrichter, M. (2009). Regioselective preparation of 5-Hydroxypropranolol and
4’-Hydroxydiclofenac with a fungal peroxygenase. Bioorganic & Medicinal
Chemistry Letters, Vol. 19: 3085–3087. (impact factor: 2.5)
150

ACKNOWLEDGEMENT
Acknowlegement
This dissertation could not have been written without the support of many people.
Foremost, I want to express my special gratitude to professor Martin Hofrichter who not
only served as my supervisor but also encouraged and challenged me throughout
this project, also for his persistence, and patience. He and my laboratory advisor
Dr. Matthias Kinne guided me through the dissertation process, never accepting
less than my best efforts. I thank them both.
I would like to thank professor Kenneth Hammel and professor Korneliusz Miksch for
willingness to be reviewer of my work.
Special thanks again to Dr. Matthias Kinne for his guidance, assistance, patience and
advice regarding the selection and operation of research methods and ideas.
I want to express my gratitude to Dr. René Ullrich, Martin Kluge and Marek Pecyna for
theirs assistance with the analytical instrumentation and fruitful discussions.
I would like to thank my laboratory colleagues Anna Karutz, Sylvia Gleißner, Stefanie
Hintersatz, Małgorzata Poraj-Kobielska, Natalia Lemańska-Malinowska and the
laboratory technicians Ulrike Schneider and Monika Brandt for their technical and
scientific assistance.
I want also express my gratitude to Dr. Gernot Kayser for fruitful discussions and help by
organize of financial support for this project.
Many thanks to whole Laboratory team for pleasant working environment and the nice
time together.
Lastly, I offer my regards and thanks to all of those who supported me in any respect
during the completion of the project, my family and friends for their
encouragement and understanding.

Versicherung an Eides statt
Hiermit versichere ich, dass ich die vorliegende Arbeit ohne unzulässige Hilfe Dritter und
ohne Benutzung anderer als der angegebenen Hilfsmittel angefertigt habe; die aus fremden
Quellen direkt oder indirekt übernommenen Gedanken sind als solche kenntlich gemacht.
Weitere Personen waren an der Abfassung der vorliegenden Arbeit nicht beteiligt. Die Hilfe
eines Promotionsberaters habe ich nicht in Anspruch genommen. Weitere Personen haben von
mir keine geldwerten Leistungen für Arbeit erhalten, die nicht als solche kenntlich gemacht
worden sind.
Die Arbeit wurde bisher weder im Inland noch im Ausland in gleicher oder ähnlicher Form
einer anderen Prüfungsbehörde vorgelegt.
Zittau, den 5. Oktober 2012
Marzena Poraj-Kobielska