C?H Amination via Nitrene Transfer Catalyzed by Mononuclear Non-Heme Iron-Dependent Enzymes

Article abstract of DOI:10.1002/cbic.201900783

Expanding the reaction scope of natural metalloenzymes can provide new opportunities for biocatalysis. Mononuclear non-heme iron-dependent enzymes represent a large class of biological catalysts involved in the biosynthesis of natural products and catabolism of xenobiotics, among other processes. Here, we report that several members of this enzyme family, including Rieske dioxygenases as well as α-ketoglutarate-dependent dioxygenases and halogenases, are able to catalyze the intramolecular C?H amination of a sulfonyl azide substrate, thereby exhibiting a promiscuous nitrene transfer reactivity. One of these enzymes, naphthalene dioxygenase (NDO), was further engineered resulting in several active site variants that function as C?H aminases. Furthermore, this enzyme could be applied to execute this non-native transformation on a gram scale in a bioreactor, thus demonstrating its potential for synthetic applications. These studies highlight the functional versatility of non-heme iron-dependent enzymes and pave the way to their further investigation and development as promising biocatalysts for non-native metal-catalyzed transformations.

Full text of DOI:10.1002/cbic.201900783

Accepted Article
Title: C—H Amination via Nitrene Transfer Catalyzed by Mononuclear
Non-Heme Iron-Dependent Enzymes
Authors: Maria Agustina Vila, Viktoria Steck, Sonia Rodriguez
Giordano, Ignacio Carrera, and Rudi Fasan
This manuscript has been accepted after peer review and appears as an
Accepted Article online prior to editing, proofing, and formal publication
of the final Version of Record (VoR). This work is currently citable by
using the Digital Object Identifier (DOI) given below. The VoR will be
published online in Early View as soon as possible and may be different
to this Accepted Article as a result of editing. Readers should obtain
the VoR from the journal website shown below when it is published
to ensure accuracy of information. The authors are responsible for the
content of this Accepted Article.
To be cited as: ChemBioChem 10.1002/cbic.201900783
Link to VoR: http://dx.doi.org/10.1002/cbic.201900783
A Journal of
www.chembiochem.org

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
C—H Amination via Nitrene Transfer Catalyzed by Mononuclear
Non-Heme Iron-Dependent Enzymes
Maria Agustina Vila[a],[c], Viktoria Steck[b],[c], Sonia Rodriguez Giordano[a], Ignacio Carrera[a],* and Rudi
Fasan[b],*
Abstract: Expanding the reaction scope of natural metalloenzymes
can provide new opportunities for biocatalysis. Mononuclear non-
heme iron-dependent enzymes represent a large class of biological
catalysts involved in the biosynthesis of natural products and catabo-
lism of xenobiotics, among other processes. Here, we report that sev-
eral members of this enzyme family, including Rieske dioxygenases
coordinated by a ‘2-His-1-carboxylate facial triad’ composed of
two His residues and a carboxylate group from an Asp or Glu
residue in the protein, along with one or two water molecules
(Figure 1 A).[4] The iron center is linked to a proximal [2Fe-2S]
cluster via a conserved aspartate hydrogen-bonded to the metal-
coordinating histidine ligands, which facilitates transfer of
as well as α-ketoglutarate dependent dioxygenases and halogenases, electrons during catalysis.[3] While featuring a similar 2-His-1-
are able to catalyze the intramolecular C—H amination of a sulfonyl
azide substrate, thereby exhibiting a promiscuous nitrene transfer re-
activity. One of these enzymes, naphthalene dioxygenase (NDO),
was further engineered resulting in several active site variants that are
functional as C—H aminases. Furthermore, this enzyme could be ap-
plied to execute this non-native transformation at a gram scale in a
bioreactor, thus demonstrating its potential for synthetic applications.
These studies highlight the functional versatility of non-heme iron-de-
pendent enzymes and pave the way to their further investigation and
development as promising biocatalysts for non-native metal-catalyzed
transformations.
carboxylate-bound iron center, members of the large subclass of
α-ketoglutarate (αKG)-dependent dioxygenases utilize αKG as
the source of electrons and co-substrate for oxidation (Figure 1
B).[1i, 5] In these enzymes, αKG participates in the coordination of
the Fe(II) atom prior to catalysis and hydroxylation of the substrate
is accompanied by oxidation of αKG to CO2 and succinate. In
αKG-dependent halogenases, the carboxylate ligand is missing,
resulting in a vacant coordination site that accommodates a halide
ligand (i.e., Cl- or Br-; Figure 1 C).[6] During catalysis, this allows
for the transfer of a halide radical to the carbon radical generated
upon H-atom abstraction in the substrate from the iron(IV)-oxo
intermediate, resulting in
a
C(sp3)—H chlorination (or
bromination) reaction. While these enzymes have been
extensively investigated in the context of their native reactions,[7]
their potential reactivity toward non-native transformations has
remained largely unexplored.[8]
Mononuclear non-heme iron (NHI) enzymes participate in a
broad range of oxidative processes implicated in the biosynthesis
of antibiotics and other natural products, DNA repair mechanisms,
and the metabolic degradation of xenobiotics.[1] As part of their
native function, these enzymes catalyze a variety of oxidation re-
actions mediated by high-valent iron-oxo intermediates, including
C(sp3)—H hydroxylation and halogenation via H-atom abstrac-
tion/radical rebound, desaturation, and arene hydroxylation and
dihydroxylation reactions.[2] The specific reactivity of these en-
zymes is influenced by the first and second sphere coordination
environment around the catalytic iron center in the active site (Fig-
ure 1). Rieske dioxygenases are multicomponent enzymes usu-
ally comprising two or three component systems: a reductase that
obtains electrons from NAD(P)H, often a Rieske ferredoxin com-
ponent that transfers the electrons, and an oxygenase component
that promotes the cis-1,2-dihydroxylation of arene substrates (e.g.
naphthalene, toluene), among other reactions.[1j, 3] The oxygen-
ase unit consist of a catalytic α subunit and a structural β subunit
organized into an α3β3 complex, where the active site iron atom is
Figure 1. Active sites of the different classes of mononuclear non-heme iron
(NHI) enzymes investigated in this work: (A) Rieske dioxygenases; (B) α-
ketoglutarate dependent dioxygenases; (C) α-ketoglutarate dependent
halogenases.
[a]
Dr. María Agustina Vila, Prof. Dr. Sonia Rodríguez, Prof. Dr. Ignacio
Carrera. Laboratorio de Biocatálisis y Biotransformaciones, Departa-
mento de Química Orgánica y Departamento de Biociencias. Facul-
tad de Química, Universidad de la República, Montevideo – Uru-
guay, Av General Flores 2124, CP 11800.
E-mail: icarrera@fq.edu.uy
[b]
[c]
Dr. Viktoria Steck, Prof. Dr. R. Fasan
Department of Chemistry, RC Box 270216, University of Rochester
Rochester, NY 14627 (USA)
E-mail: rfasan@ur.rochester.edu
Equal contribution.
Supporting information for this article is given via a link at the end of
the document.
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
dioxygenase TauD, which catalyzes the oxidation of 2-
aminoethanesulfonic acid (taurine) under sulfur starvation
conditions[17] and tolerates non-native substrates;[17b] E. coli AlkB,
which is involved in the repair of alkylated DNA;[1a, 18] E. coli Gab
dioxygenase, a clavaminate synthase-like enzyme putatively
involved in the biosynthesis of β-lactam secondary metabolite;[19]
the multifunctional fungal dioxygenase AsqJ
(Aspergillus
nidulans), which catalyzes a tandem desaturation/epoxidation
reaction during the biosynthesis of the alkaloid 4′‐
20]
methoxyviridicatin;[1d,
hydroxylase
and the plant hyoscyamine 6β-
which catalyzes
(H6H),
a
hydroxylation/dehydrogenation/epoxidation sequence in the
biosynthesis of the anticholinergic alkaloid scopolamine in the
Solanaceae family of plants.[21] As a representative member of
NHI halogenases, we included in the panel the cyanobacterial
enzyme WelO5, which catalyzes the chlorination of 12-epi-
fischerindole as part of the biosynthesis of the welwitindolinone
natural products.[22] Compared to other well characterized NHI
halogenases such as SyrB2,[23] WelO5 was chosen because it
acts on a free-standing small-molecule substrate as opposed to a
carrier protein-linked substrate. In addition, its structure was
recently elucidated by X-ray crystallography.[22c]
Figure 2. Enzyme-catalyzed transformations involving iron nitrene species: (A)
TDO-catalyzed conversion of benzyl azide to benzonitrile and subsequent
dihydroxylation; (B) Mb/P450-catalyzed oxidative deamination of benzyl azides;
(C) P450/Mb-catalyzed C—H amination of sulfonyl azides.
Recently, the Rieske toluene dioxygenase (TDO) from
Pseudomonas putida F1 was found to catalyze an unexpected
transformation resulting in the conversion of benzyl azide to
benzonitrile, in addition to the expected cis-dihydroxylation
product (Figure 2A). Based on computational (DFT) and
mechanistic studies, we proposed a plausible mechanism for this
reaction that involves the formation of an iron(IV)–nitrene
intermediate, which, upon rearrangement of the hydroxyl group
and dehydration of the resulting benzaldoxime, leads to the
observed benzonitrile product.[9]
A
similar activity was
subsequently observed for Fe/αKG dependent dioxygenases.[10]
Interestingly, the proposed mechanism for these reactions share
similarities with that reported for the P450- and Mb-catalyzed
oxidative deamination of benzyl azides to yield benzaldehydes
(Figure 2B).[11] In addition, the putative iron–nitrene intermediate
invoked in the context of the TDO reaction is analogous to that
previously proposed by us to mediate C—H amination reactions
via nitrene transfer supported by engineered P450s[12] or other
hemoproteins (Figure 2C).[13] Based on these considerations, we
surmised that Rieske dioxygenases and potentially other non-
heme iron enzymes could be able to exhibit non-native nitrene
transfer reactivity.
Figure 3. C—H amination of 2,4,6-triisopropylbenzenesulfonyl azide 1a with
Rieske dioxygenases (A). (B) Biotransformations with 1a (5 mM) using E. coli
whole cells expressing NDO (JM109(DE3) (pDTG141)); TDO (JM109
(pDTG601)); CBDO (JM109 (pKK-CBDO)) at OD600
= 60 in potassium
phosphate buffer (50 mM, pH 7.2), anaerobic conditions. Control 1: E. coli
JM109 (DE3). Control 2: E. coli JM109 (pKK223-3). In each case, sulfonamide
(1c) amounted to <5%. (C) Same reaction of NDO and control as in (B) but with
10 mM 1a.
To test this hypothesis, we surveyed a diverse panel of
functionally and structurally different non-heme iron dependent
oxidases. This enzyme selection included the Rieske toluene
(TDO), chlorobenzene (CBDO) and naphthalene (NDO)
dioxygenases, which catalyze the cis-dihydroxylation of the
corresponding aromatic compound. TDO, which accepts a wide
variety of mono and disubstituted arenes as substrates, has been
extensively used in the last decades as a whole cell biocatalyst
for the preparation of enantiopure cis-cyclohexadienediols for
enantioselective synthesis.[14] In contrast, NDO engages larger
substrates such as polycyclic arenes,[15] while the substrate scope
of CBDO has been much less explored.[16] Additionally, the
enzyme panel included representative members of the subclass
of Fe/αKG dependent dioxygenases, namely E. coli taurine
Starting with the Rieske dioxygenases, we tested these
enzymes for their C—H amination activity using as a model
reaction the conversion of 2,4,6-triisopropylbenzenesulfonyl azide
1a to the sultam product 1b (Figure 3A). Due to the
multicomponent nature of Rieske dioxygenases, NDO, TDO and
CBDO were expressed in E. coli strains and tested in whole-cell
biotransformations.[24] Initial tests revealed that all of these Rieske
enzymes biocatalysts exhibited detectable C—H amination
activity (see SI Table S1). Based on these promising results, the
reaction conditions for these biotransformations were further
optimized (see SI Figure S1-S2). Under optimized conditions,
both NDO and CBDO were found to provide a significantly higher
conversion of sulfonyl azide 1a to the C—H amination product 1b
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
compared to the control strains (37-38% vs. 20-22%; Figure 3B).
The observed background activity may stem from heme-
containing enzymes present in E. coli.[25] For the NDO-expressing
cells, further improved conversion of 1a into 1b over the
background levels was achieved at higher substrate loading (i.e.
10 mM vs. 5 mM, Figure 3C). In contrast to NDO and CBDO, the
cells expressing TDO showed no significant activity over
background levels (Figure 3B). Overall, these results
demonstrate for the first time that the NDO and CBDO Rieske
dioxygenases possess promiscuous nitrene transferase activity
that can be applied for catalyzing C—H amination reactions.
Along with the C—H amination product, both biocatalysts were
found to generate a small amount of sulfonamide 1c (approx. 4%,
see Supporting Information). This by-product was previously
observed also in the context of hemoprotein-catalyzed nitrene
Figure 4. NDO structure (pdb 1NDO). (A) Quaternary structure (α3β3) of NDO
showing alpha subunits in green and beta subunits in orange. (B) View of the
active site (boxed panel (A)) showing the iron atom (red, sphere), Fe-
coordinating residues (purple, stick) and active site residues targeted for
mutagenesis (yellow, stick model).
Based on these results, the aforementioned mutations were
recombined to generate a double and triple mutant library. From
this library, NDO variants carrying a F224V mutation along with
H295I, H295T, or H295V, were found to be functional C—H
aminases (Figure S3). Although none of these NDO variants
showed improved C—H amination activity compared to the wild-
type enzyme, these results showed that mutation of various active
site positions, and in particular at the level of residues 224, 295,
307, and 358, are compatible with this non-native reactivity, and
expected to facilitate further protein engineering efforts.
In order to examine the synthetic potential of NDO in the
context of this non-native reaction, we performed a large-scale
biotransformation of 1a (1 g, 3 mM) with NDO-expressing E. coli
JM109 (DE3) cells in a bioreactor (5 L). For this purpose, we used
our previously optimized fed-batch protocol to express the
dioxygenase in a high cell density culture (approx. 50 g/L cdw),
where the substrate is added after reaching stationary phase.[30]
As shown in Figure 5, this experiment showed progressive
consumption of the azide substrate 1a and concomitant
accumulation of the desired sultam product 1b within three hours.
transfer reactions with sulfonyl azides and other azide-based
26]
nitrene precursors.[12,
Nevertheless, both Rieske enzymes
showed a high selectivity toward the formation of the desired C—
amination product 1b (>90 %) over this side-reaction.
H
Interestingly, the NDO-dependent C—H amination activity was
found to be reasonably oxygen tolerant, maintaining in the
presence of air approximately 60% of the activity levels observed
under anaerobic conditions (see Supporting Information). This is
contrast to P450-catalyzed nitrene transfer reactions, which
require strictly anaerobic conditions both with purified proteins
and in whole cells.[12, 26]
Encouraged by these findings, we selected NDO as the
target for mutagenesis studies in order to gain insights into the
effect of active site mutations on the non-native C—H amination
activity of this enzyme. To this end, we generated a series of
libraries sampling 12 representative amino acids with different
side-chain groups—i.e., Phe, Ser, Tyr, Leu, Pro, His, Ile, Thr, Asn,
Val, Ala, Asp, using the degenerate NHT codon—[27] at positions
Asn201, Phe202, His295, Leu307, Phe352 and Trp358, which are
located within the substrate binding pocket of the enzyme[28]
(Figure 4). Since Phe224 serves as a gating residue at the
entrance of the substrate channel leading to the active site,[29] this
position was replaced with eight smaller amino acids (i.e., Ile, Thr,
Asn, Ser, Val, Ala, Asp, Gly) using the RNC degenerate codon.
These libraries were expressed in 48-well plates and screened for
C—H amination activity toward substrate 1a by HPLC. The initial
screening experiments showed that mutagenesis of positions
N201, F202 and F352 had mostly deleterious effects on the C—
H amination activity of the enzyme. In contrast, various large-to-
small mutations at positions Phe224 (→ Ala, Ile, Val), His295 (→
Ile, Thr, Val), and Trp358 (→ Asn, along with Phe and Tyr) led to
wild type-like activity toward the non-native reaction (Figure S3).
At position 307, a Leu→His mutation was also tolerated.
Under these conditions, only a negligible fraction of the
sulfonamide byproduct 1c was generated in the course of the
reaction. Overall, this protocol enabled the conversion of 1 g 1a
into 1b in 45% yield and >90% selectivity, thus providing a proof-
of-concept of the scalability of the NDO-driven reaction.
Figure 5. Biotransformation profile for the C—H amination reaction of 1 gram of
2,4,6-triisopropylbenzensulfonyl azide 1a carried out in a 5 L bioreactor using E.
coli JM109 (DE3) (pDTG141-Kan).
After the promising results obtained for the Rieske
biocatalysts, we investigated of the C—H amination activity of
other NIH enzymes such as Fe/αKG dependent dioxygenases
and halogenases. As summarized in Table 1, all of the tested
Fe/αKG dependent dioxygenases and halogenases, with the only
exception of AlkB, were found to display promiscuous C—H
amination activity in the reaction with 1a. Among them, TauD and
AsqJ showed the highest activity, resulting in the generation of
the C—H amination product in 25-27% yield as determined by
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
HPLC. Since these enzymes rely on αKG as co-substrate for
metal coordination in the context of their native dioxygenase /
halogenase activity, these reactions were also carried out in the
presence of media supplemented with αKG along with iron and
ascorbic acid. Interestingly, a differential effect of the added αKG
was observed for the different enzymes. Specifically, no change
in C—H amination activity was observed for Gab and WelO5
under these conditions, whereas an approximately 2-fold
reduction in yield was noted for TauD, AsqJ and H6H. These
results suggest that at least for the latter enzymes, αKG may not
be required and possibly have an inhibitory effect in the context
of the non-native reaction. For most of the active enzymes, the
C—H amination was found to proceed with relatively good
selectivity, ranging from 53 to 67%. As an exception, H6H
exhibited modest chemoselectivity (33%), producing sulfonamide
1c as the major product of the reaction.
dioxygenases did not show detectable activity toward this
substrate (data not shown), some of the Fe/αKG dependent
enzymes exhibited appreciable activity toward this nitrene
transfer reaction. In particular, the dioxygenases AlkB and H6H
and chlorinase WelO5 were found able to covert 2a into 2b in 2-
6% yield in the whole-cell reactions. Furthermore, while AlkB and
H6H were found to produce 2-phenylpropyl carbamate (2c) as
well as 2-phenylpropan-1-ol (2d) as byproducts, the latter being
observed also in the context of the P450-catalyzed reaction,[12b]
WelO5 exhibited a significantly higher selectivity for the desired
C—H amination reaction, producing 2b with 80% selectivity and
no alcohol byproduct (Table 1, Entry 12; SI Figure S6). Altogether,
these experiments revealed that NHI enzymes are able to engage
different types of nitrene donors in intramolecular C—H amination
reactions, with the individual members of this group displaying
differential selectivity and reactivity toward the two different azide
substrates. While the mechanism of these reactions awaits further
11]
elucidation, our previous studies[9,
suggest a mechanism
Table 1. Activity and selectivity of Fe/αKG dependent dioxygenases and
halogenases toward the intramolecular C—H amination of 2,4,6-
triisopropylbenzensulfonyl azide 1a (reaction (1)) and 2-phenylpropyl
carbonazidate 2a (reaction (2)).[a]
analogous to that of hemoprotein-catalyzed C—H amination,[12-13]
in which binding of the azide substrate to the iron center leads to
an iron-nitrene intermediate via nitrogen extrusion, followed by a
nitrene C—H insertion process.
In summary, this work demonstrates that mononuclear non-
heme iron dependent oxidases are capable of catalyzing
intramolecular C—H amination reactions involving sulfonyl azides
and carbonyl azides, thereby exhibiting a promiscuous nitrene
transferase activity. While this work was being finalized for
submission, the Arnold group reported a similar type of reactivity
for the Fe/αKG-dependent Pseudomonas savastanoi ethylene-
forming enzyme (PsEFE).[31] Complementing and expanding
upon these findings, the present studies show that other types of
αKG-dependent dioxygenases as well as Rieske dioxygenases
and αKG-dependent halogenases exhibit such non-native
reactivity. Through mutagenesis studies, we gained initial insights
into the tolerance of active site mutations within the Rieske
naphthalene dioxygenase NDO toward this catalytic activity.
Notably, this enzyme could be applied for a gram-scale C—H
amination reaction in a bioreactor, which provides a proof-of-
principle demonstration of its synthetic potential in the context of
this non-native transformation. Overall, our studies illustrate the
functional versatility of non-heme iron-dependent enzymes and
open the way to their further investigation and development as
biocatalysts for other non-native metal-catalyzed transformations,
which are the subject of ongoing studies in our laboratory.
O
O
S
NH
N3
SO2
SO2NH2
Biocat.
+
(1)
1c
1a
1b
O
O
O
O
O
Biocat.
N
+
O
(2)
H
N3
NH2
2c
2a
2b
1b
2b
Conditions
Entry
Enzyme
[b]
Yield[c]
Yield[c]
(Selectivity)[d]
(Selectivity)[d]
1
2
3
4
5
6
7
8
9
TauD
TauD
AlkB
AlkB
Gab
A
B
A
B
A
B
A
B
A
B
A
B
27% (63%)
13% (62%)
n.a.
n.a.
n.a.
2% (30%)
n.a.
n.a.
n.a.
n.a.
n.a.
6% (27%)
n.a.
n.a.
n.a.
14% (57%)
13% (53%)
24% (67%)
12% (59%)
13% (33%)
n.a.
Gab
AsqJ
AsqJ
H6H
H6H
WelO5
WelO5
10
11
12
15% (58%)
13% (61%)
Experimental Section
3% (80%)
[a] E. coli cells expressing the indicated enzyme at OD600 = 40 in 50 mM kPi buffer (pH
7.0), 3 mM substrate. Condition A: no additives; Condition B: containing aKG, ascorbate,
and iron salt (see SI Table S2). [c] HPLC yield as calculated based on calibration curve
with authentic standards, and subtracted by the background activity of BL21 cells. N.a. =
not active. [d] Selectivity for formation of C—H amination product over reduction (and
alcohol) byproduct, i.e., 1b/(1b+1c) or 2b/(2b+2c+2d). See also SI Figure S5 and S6.
Reagents and Analytical Methods. Chemical reagents and
solvents were purchased from Sigma-Aldrich unless otherwise
stated. Compound 1a, 1b, 2a, and 2b were prepared as reported
previously.[12] HPLC analyses were performed using a Shimadzu
LC-2010A-HT equipped with a VisionHT-C18 reverse-phase
column and a multi-diode UV-Vis detector, or on a Shimadzu
Prominence Liquid Chromatograph LC-20AT equipped with an
Eclipse XDB-C18 Zorbax reverse-phase column (250 mm length,
4.6 mm diameter, 5 µm particle size) and a diode-array detector.
These experiments thus demonstrated that both Rieske and
Fe/αKG
dependent
dioxygenases/halogenases
possess
promiscuous nitrene transferase activity. On the basis of these
results, the entire enzyme panel was further tested for their ability
to catalyze another type of intramolecular C—H amination
reaction, namely the cyclization of carbonazidate 2a to give the
oxazolidinone product 2b (Table 1).[12b] While Rieske
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
Strains and plasmids. E. coli JM109 were obtained from New
England Biolabs and E. coli JM109 (DE3) from Promega. E. coli
JM109 (pDTG601) expressing Toluene Dioxygenase (TDO) was
kindly donated by Prof. David T. Gibson, E. coli JM109 (DE3)
(pDTG141) expressing Naphthalene Dioxygenase (NDO) was
obtained from Prof. Rebecca Parales (University of California,
USA). E. coli BL21 (pET28b_SUMO) containing the AsqJ gene
was kindly donated by Profs. Lukas Hintermann and Michael Groll
(Technical University of Munich). E. coli BL21 (pET22b_H6H)
containing the H6H gene was kindly provided by Alejandra B.
Cardillo (Universidad de Buenos Aires).
S2 and cloned into the Xho I/Nde I cassette of pET22b (Novagen)
to give the expression vector pET22b_WelO5. The NDO-derived
single-site saturation libraries were prepared by a QuickChange
PCR protocol using the corresponding primers in Table S2 and
pDTG141 as template. Double and triple mutants were
constructed by the same procedure using primers 23-40 and the
first-generation mutants as template..
Protein Expression. For the Fe/α-KG dependent oxidases the
corresponding recombinant strains of E. coli BL21 were grown in
LB media (AlkB, Gab, AsqJ, H6H and WelO5) or LB low salt
media (TauD) supplemented with ampicillin (100 mg L-1) at 37 °C,
200 rpm. Enzymatic expression was induced with β-D-1-
Culture media. Luria-Bertani (LB) medium used for cell growth
contained: Bacto Tryptone (10 g/L), Bacto Yeast Extract (5 g/L),
and sodium chloride (10 g/L). For LB low salt medium, 5 g/L
sodium chloride was used. Agar (15 g/L) was added for solid
media. When needed the medium was supplemented with sterile
ampicillin sodium salt (0.1 g/L) or kanamycin (0.05 g/L). Mineral
Salts Broth (MSB) used for the precultures for the bioreactor
cultures contained: K2HPO4 (16 g/L), KH2PO4 (14 g/L), (NH4)2SO4
(5 g/L), Bacto Yeast Extract (15 g/L); after sterilization the medium
was supplemented with sterile glucose (30 g/L), MgSO4•7H2O (2
g/L) and Kanamycin (0.05 g/L). Bioreactor defined mineral salt
medium consisted of: KH2PO4 (7.5 g/L), citric acid (2.0 g/L),
MgSO4•7H2O (5.0 g/L), ferric ammonium citrate (0.3 g/L), 98%
H2SO4 (1.4 mL/L) and trace metal solution (1.5 mL/L). After
sterilization, pH is regulated to 6.8 by addition of conc. ammonium
hydroxide, followed by supplementation with sterile thiamine
hydrochloride (0.3 g/L) and ampicillin sodium salt (0.1 g/L). Trace
metal solution contained: citric acid (40 g/L), MnSO4•2H2O (30
g/L), NaCl (10 g/L), FeSO4•7H2O (1 g/L), CoCl2•6H2O (1 g/L),
ZnSO4•7H2O (1 g/L), CuSO4•5H2O (0.1 g/L), H3BO3 (0.1 g/L),
NaMoO4•2H2O (0.1 g/L); pH was adjusted to 3.0 with ammonium
hydroxide.
thiogalactopyranoside (IPTG) when the cultures reached OD600
=
0.6, and incubated under the desired conditions: TauD (0.5 mM
IPTG, 30 °C, 5 h),[17b] AlkB (1 mM IPTG, 37 °C, 3 h),[33] Gab (0.5
mM IPTG, 30 °C, 5 h), AsqJ (0.1 mM IPTG, 30 °C, 4 h),[20] H6H
(1 mM IPTG, 30 °C, 4 h),[34] WelO5 (1 mM IPTG, 16 °C, 16 h).[22a]
Cells were harvested at 4000 rpm for 20 min, resuspended in
potassium phosphate buffer (50 mM, pH 7.2) and stored briefly at
-4 °C. For the Rieske dioxygenases, the corresponding
recombinant strains of E. coli JM109 or JM109 (DE3) were grown
in LB media supplemented with ampicillin (100 mg L-1) at 37 °C,
200 rpm. Enzymatic expression was induced with 0.25 mM β-D-
1-thiogalactopyranoside (IPTG) when the cultures reached OD600
= 0.5, and incubated overnight at 27 °C, 200 rpm. Cells were
harvested at 4,000 rpm for 15 min and washed once with
potassium phosphate buffer (50 mM, pH 7.2). The cell pellet was
resuspended in potassium phosphate buffer (50 mM, pH 7.2)
containing 5% v/v glycerol and 5% w/v glucose, frozen and stored
at -78 °C. The engineered NDO libraries were expressed in 48-
deep-well plates by inoculating single colonies into 4 mL of LB
media supplemented with ampicillin (100 mg L-1) in each well.
Plates were incubated overnight at 37 °C, 180 rpm. 40 μL of these
overnight cultures were transferred to a new 48-well plate
containing 4 mL fresh LB media containing ampicillin (100 mg L-
1) and incubated under the same conditions until OD600 = 0.6.
Enzyme expression was induced by adding 0.25 mM IPTG,
following by incubation overnight at 27 °C, 180 rpm. The plates
were centrifuged at 4,000 rpm for 30 minutes and the cell pellets
were resuspended in potassium phosphate buffer (50 mM, pH
7.2) containing 5% v/v glycerol and 5% w/v glucose to reach an
OD600 = 30. The cell suspensions were subjected to a freeze/thaw
cycle prior to the reactions.
Cloning and vector construction. The pDTG141-Kan
expression vector was obtained by replacing the β-lactamase
cassette with a kanamycin cassette in pDTG141. The kanamycin
resistance gene was amplified from pET28b using the
corresponding primers in Table S2 and the resulting amplicon was
used in a RFC reaction[32] to switch ampicillin resistance for
kanamycin resistance in pDTG141 to give the expression vector
pDTG141-Kan. The pKK_CBDO expression vector was
constructed by subcloning the CBDO operon from vector
pTCB144 into pKK223-3. The CBDO operon was amplified using
the corresponding primers in Table S2. The resulting amplicon
was digested with Hind III and cloned into Hind III-digested and
CIAP dephosphorylated pKK223-3 to yield pKK_CBDO. The
genes encoding for TauD, AlkB and Gab were amplified from E.
coli genome (DH5α strain) using the corresponding primers in
Table S2 followed by digestion with Xho I and Nde I restriction
enzymes and ligation into the Xho I/Nde I cassette of pET22b
(Novagen) to give the expression vectors pET22b_TauD,
pET22b_AlkB and pET22b_Gab, respectively. The vector
encoding for WelO5 was prepared using a synthetic DNA
template (Genscript) with optimized codons for expression in E.
coli. The WelO5 gene was amplified using the primers in Table
Whole-cell C—H amination reactions. Reactions using E. coli
whole cells expressing the non-heme iron oxidases were carried
out at 400 µL-scale using OD600 = 30-60 and 2.5-10 mM substrate
concentration in potassium phosphate buffer (50 mM, pH 7.2). In
a typical procedure, the whole cell suspensions were transferred
to an anaerobic chamber. Additives (αKG, ascorbate, iron salt)
were added in the anaerobic chamber from stock solutions in
argon purged MilliQ water. Reactions were initiated by addition of
substrate (400 mM stock solution in EtOH) and the reaction
mixtures were stirred or shaken in the chamber for 12 hours at
room temperature. After that, the reactions were analyzed by
adding 8 µL of internal standard (fluorenone, 50 mM in DMSO),
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
followed by extraction with 400 µL of dichloromethane. The
organic layer was removed via evaporation and the residue was
dissolved in 300 μL methanol, filtered through 0.22 μm syringe
filters, and analyzed by HPLC. Separation method: Injection
volume: 20 µL. Flow rate: 1 mL/min. Gradient: 30% acetonitrile in
water (0.1% TFA) for 3 min, then increased to 90% over 18 min.
Detector: 254 nm. Calibration curves were constructed using
synthetically produced products. All measurements were
performed at least in duplicate.
thank Juan Mauricio Minoia from Universidad de Buenos Aires for
preparing bacterial cultures expressing H6H.
Keywords: non-heme iron-dependent oxidases • Rieske
diooxygenases • nitrene transfer • C—H amination • protein
engineering
References
Large scale biotransformation. Scale-up biotransformation was
performed in a 5-liter bioreactor (Sartorius Biostat A plus) using a
modification of a previously published procedure.[30] Fresh plates
of E. coli JM109 DE3 (pDTG141-Kan) were used to inoculate 5
mL LB-Kan which was incubated in orbital shaker overnight (150
rpm, 37 °C). Two 500 mL Erlenmeyer flasks containing 150 mL of
MSB medium were inoculated with 1.5 mL of the overnight culture
and grown for 12 hours (37 °C, 150 rpm). Both 500 mL cultures
were used to inoculate the bioreactor (Sartorius Biostat A plus)
charged with an initial volume of 2.5 L, and set to 500 rpm, 30 °C,
and air flow rate of 4 L/min, and a pulse of antifoam agents
(Aldrich’s Antifoam 201: Silicone dispersion in water 1:1) was
added at the beginning of the run. The pH value was adjusted
automatically to 6.8 by addition of ammonium hydroxide
throughout the reaction. A glucose fed-batch started 6 hours after
inoculation by adding glucose (0.7 g/mL solution) from an initial
rate of 0.08 mL/min to 0.54 mL/min in a 20 h period. Enzyme
expression was induced by adding IPTG to a final concentration
of 10 mg/L after 12 hours from inoculation, and stirrer speed was
set to 900 rpm. After the culture reached the stationary phase
(~26 h, ~30 g/L cdw), 1.5 L of media was removed from the
bioreactor and 1 g of 2,4,6-triisopropylbenzenesulfonyl azide 1a
in MeOH was added to a final concentration of 3 mM. Samples
were withdrawn at different time points, extracted with DCM, and
analyzed by HPLC as described above.
[1]
a) B. I. Fedeles, V. Singh, J. C. Delaney, D. Li, J. M. Essigmann, J. Biol.
Chem. 2015, 290, 20734-20742; b) A.-X. Cheng, X.-J. Han, Y.-F. Wu,
H.-X. Lou, Int. J. Mol. Sci. 2004, 15, 1080-1095; c) S.-S. Gao, N.
Naowarojna, R. Cheng, X. Liu, P. Liu, Nat. Prod. Rep. 2018, 35, 792-
837; d) H. Nakamura, Y. Matsuda, I. Abe, Nat. Prod. Rep. 2018, 35, 633-
645; e) C. Perry, Emmanuel L. C. de los Santos, L. M. Alkhalaf, G. L.
Challis, Nat. Prod. Rep. 2018, 35, 622-632; f) L. Shen, C.-X. Song, C.
He, Y. Zhang, Annu. Rev. Biochem. 2014, 83, 585-614; g) S. C. Farrow,
P. J. Facchini, Front. Plant Sci. 2014, 5; h) M. D. White, E. Flashman,
Curr. Opin. Chem. Biol. 2016, 31, 126-135; i) M. S. Islam, T. M. Leissing,
R. Chowdhury, R. J. Hopkinson, C. J. Schofield, Annu. Rev. Biochem.
2018, 87, 585-620; j) S. M. Barry, G. L. Challis, ACS Catal. 2013, 3, 2362-
2370.
[2]
a) A. L. Feig, S. J. Lippard, Chem. Rev. 1994, 94, 759-805; b) L. Que, R.
Y. N. Ho, Chem. Rev. 1996, 96, 2607-2624; c) M. Costas, M. P. Mehn,
M. P. Jensen, L. Que, Chem. Rev. 2004, 104, 939-986; d) M. M. Abu-
Omar, A. Loaiza, N. Hontzeas, Chem. Rev. 2005, 105, 2227-2252; e) E.
G. Kovaleva, J. D. Lipscomb, Nat. Chem. Biol. 2008, 4, 186-193; f) S.
Martinez, R. P. Hausinger, J. Biol. Chem. 2015, 290, 20702-20711.
D. J. Ferraro, L. Gakhar, S. Ramaswamy, Biochem. Biophys. Res.
Commun. 2005, 338, 175-190.
[3]
[4]
a) R. E. Parales, J. Ind. Microbiol. Biotechnol. 2003, 30, 271-278; b) B.
Kauppi, K. Lee, E. Carredano, R. E. Parales, D. T. Gibson, H. Eklund, S.
Ramaswamy, Structure 1998, 6, 571-586.
[5]
a) C. Schofield, R. Hausinger, 2-Oxoglutarate-dependent oxygenases,
Royal Society of Chemistry, 2015; b) F. P. Guengerich, J. Biol. Chem.
2015, 290, 20700-20701; c) R. P. Hausinger, Crit. Rev. Biochem. Mol.
Biol. 2004, 39, 21-68.
[6]
[7]
A. Timmins, S. P. De Visser, Catalysts 2018, 8, 314.
a) W. Hüttel, Chem. Ing. Tech. 2013, 85, 809-817; b) C. Loenarz, C. J.
Schofield, Nat. Chem. Biol. 2008, 4, 152-156; c) M. Hibi, J. Ogawa, Appl.
Microbiol. Biotechnol. 2014, 98, 3869-3876; d) J. Latham, E.
Brandenburger, S. A. Shepherd, B. R. K. Menon, J. Micklefield, Chem.
Rev. 2018, 118, 232-269; e) A. J. Mitchell, J.-K. Weng, Plant Physiol.
2019, 179, 813-829; f) Y. Nakashima, T. Mori, H. Nakamura, T. Awakawa,
S. Hoshino, M. Senda, T. Senda, I. Abe, Nat. Commun. 2018, 9, 104; g)
C. Peters, R. M. Buller, Catalysts 2019, 9, 221-241.
Statistical Analyses. GraphPad Prism software 8 was used for
data analysis of Rieske dioxygenase catalyzed reactions. Data
are presented as mean ± SD values from five replicates. The
results were analyzed and compared by one-way ANOVA
followed by Sidak’s Multiple Comparison Test, and in all cases
statistical significance was set at P < 0.05.
[8]
[9]
M. Guo, T. Corona, K. Ray, W. Nam, ACS Cent. Sci. 2019, 5, 13-28.
a) N. Thevenet, V. de la Sovera, M. A. Vila, N. Veiga, D. Gonzalez, G.
Seoane, I. Carrera, Org. Lett. 2015, 17, 684-687; b) M. A. Vila, M. Pazos,
C. Iglesias, N. Veiga, G. Seoane, I. Carrera, Chembiochem 2016, 17,
291-295.
Acknowledgements
R.F. and V.S. acknowledge financial support from the U.S.
National Science Foundation grants CHE-1609550 and CHE-
1834390. MS instrumentation was supported by the U.S. National
Science Foundation grants CHE-0840410 and CHE-0946653.
A.V., S.R. and I.C. acknowledge financial support from
PEDECIBA, CSIC-UdelaR (Proyect INI-1828), Agencia Nacional
de Investigación e Innovación (ANII - Uruguay, Project FCE
103485) and the PhosAgro/ UNESCO / IUPAC Partnership in
Green Chemistry for Life for a research grant awarded to I.C. A.V.
acknowledges CAP-UdelaR for a PhD. Scholarship. We also
[10] M. Davidson, M. McNamee, R. Fan, Y. Guo, W.-c. Chang, J. Am. Chem.
Soc. 2019, 141, 3419-3423.
[11] a) S. Giovani, H. Alwaseem, R. Fasan, ChemCatChem 2016, 8, 2609-
2613; b) S. Giovani, R. Singh, R. Fasan, Chem. Sci. 2016, 7, 234-239.
[12] a) R. Singh, M. Bordeaux, R. Fasan, ACS Catal.
2014, 4, 546-552; b) R. Singh, J. N. Kolev, P. A. Sutera, R. Fasan, ACS Catal.
2015, 5, 1685-1691.
[13] M. Bordeaux, R. Singh, R. Fasan, Bioorg. Med. Chem. 2014, 22, 5697-
5704.
[14] a) T. Hudlicky, D. Gonzalez, D. Gibson, Aldrichimica acta 1999, 32, 35-
62; b) S. E. Lewis, Chem. Commun. 2014, 50, 2821-2830; c) T. Hudlicky,
ACS Omega 2018, 3, 17326-17340.
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
[15] a) S. Resnick, K. Lee, D. Gibson, J. Ind. Microbiol. 1996, 17, 438-457; b)
R. E. Parales, K. Lee, S. M. Resnick, H. Jiang, D. J. Lessner, D. T.
Gibson, J. Bacteriol. 2000, 182, 1641-1649; c) C. Chopard, R. Azerad, T.
Prangé, J. Mol. Catal. B: Enzym. 2008, 50, 53-60; d) D. J. Ferraro, A.
Okerlund, E. Browna, S. Ramaswamy, IUCrJ 2017, 4, 648-656.
[16] a) C. Werlen, H.-P. E. Kohler, J. R. van der Meer, J. Biol. Chem. 1996,
271, 4009-4016; b) S. Yildirim, J. Zezula, T. Hudlicky, B. Witholt, A.
Schmid, Adv. Synth. Catal. 2004, 346, 933-942; c) S. Yildirim, T. T.
Franko, R. Wohlgemuth, H.-P. E. Kohler, B. Witholt, A. Schmid, Adv.
Synth. Catal. 2005, 347, 1060-1072.
Ruszczycky, W.-c. Chang, F. Yan, Y.-n. Liu, H.-w. Liu, J. Am. Chem. Soc.
2018, 140, 7433-7436.
[22] a) M. L. Hillwig, X. Liu, Nat. Chem. Biol. 2014, 10, 921-923; b) M. L.
Hillwig, H. A. Fuhrman, K. Ittiamornkul, T. J. Sevco, D. H. Kwak, X. Liu,
Chembiochem 2014, 15, 665-669; c) A. J. Mitchell, Q. Zhu, A. O.
Maggiolo, N. R. Ananth, M. L. Hillwig, X. Liu, A. K. Boal, Nat. Chem. Biol.
2016, 12, 636-640.
[23] F. H. Vaillancourt, J. Yin, C. T. Walsh, Proc. Natl. Acad. Sci. U. S. A.
2005, 102, 10111-10116.
[24] J. M. Halder, B. M. Nestl, B. Hauer, ChemCatChem 2018, 10, 178-182.
[25] M. J. Weissenborn, S. A. Löw, N. Borlinghaus, M. Kuhn, S. Kummer, F.
Rami, B. Plietker, B. Hauer, ChemCatChem 2016, 8, 1636-1640.
[26] a) C. C. Farwell, R. K. Zhang, J. A. McIntosh, T. K. Hyster, F. H. Arnold,
ACS Cent. Sci. 2015, 1, 89-93; b) C. K. Prier, R. J. K. Zhang, A. R. Buller,
S. Brinkmann-Chen, F. H. Arnold, Nat Chem 2017, 9, 629-634.
[27] M. T. Reetz, D. Kahakeaw, R. Lohmer, Chembiochem 2008, 9, 1797-
1804.
[17] a) E. Eichhorn, J. R. van der Ploeg, M. A. Kertesz, T. Leisinger, J. Biol.
Chem. 1997, 272, 23031-23036; b) D. Wetzl, J. Bolsinger, B. M. Nestl,
B. Hauer, ChemCatChem 2016, 8, 1361-1366.
[18] a) H. Kondo, Y. Nakabeppu, H. Kataoka, S. Kuhara, S. Kawabata, M.
Sekiguchi, J. Biol. Chem. 1986, 261, 15772-15777; b) P. Ø. Falnes, R.
F. Johansen, E. Seeberg, Nature 2002, 419, 178-182.
[19] M. R. Chance, A. R. Bresnick, S. K. Burley, J.-S. Jiang, C. D. Lima, A.
Sali, S. C. Almo, J. B. Bonanno, J. A. Buglino, S. Boulton, H. Chen, N.
Eswar, et al., Protein Sci. 2002, 11, 723-738.
[28] B. Kauppi, K. Lee, E. Carredano, R. E. Parales, D. T. Gibson, H. Eklund,
S. Ramaswamy, Structure 1998, 6, 571-586.
[20] A. Bräuer, P. Beck, L. Hintermann, M. Groll, Angew. Chem., Int. Ed. 2016,
55, 422-426.
[29] D. E. Escalante, K. G. Aukema, L. P. Wackett, A. Aksan, J. Chem. Inf.
Model. 2017, 57, 550-561.
[21] a) G. Fodor, A. Romeike, G. Janzsó, I. Koczor, Tetrahedron Lett. 1959,
1, 19-23; b) T. Hashimoto, Y. Yamada, Plant Physiol. 1986, 81, 619-625;
c) T. Hashimoto, Y. Yamada, Eur. J. Biochem. 1987, 164, 277-285; d) T.
Hashimoto, J. Kohno, Y. Yamada, Plant Physiol. 1987, 84, 144-147; e)
T. Hashimoto, J. Kohno, Y. Yamada, Phytochemistry 1989, 28, 1077-
1082; f) J. Matsuda, S. Okabe, T. Hashimoto, Y. Yamada, J. Biol. Chem.
1991, 266, 9460-9464; g) T. Hashimoto, J. Matsuda, Y. Yamada, FEBS
Lett. 1993, 329, 35-39; h) I. Klinkenberg, A. Blokland, Neurosci. Biobehav.
Rev. 2010, 34, 1307-1350; i) J. Li, M. J. van Belkum, J. C. Vederas,
Bioorg. Med. Chem. 2012, 20, 4356-4363; j) R. Ushimaru, M. W.
[30] M. A. Vila, M. Brovetto, D. Gamenara, P. Bracco, G. Zinola, G. Seoane,
S. Rodríguez, I. Carrera, J. Mol. Catal. B: Enzym. 2013, 96, 14-20.
[31] N. W. Goldberg, A. M. Knight, R. K. Zhang, F. H. Arnold, J. Am. Chem.
Soc. 2019, 141, 19585-19588.
[32] F. van den Ent, J. Löwe, J. Biochem. Biophys. Methods 2006, 67, 67-74.
[33] S. Dinglay, S. C. Trewick, T. Lindahl, B. Sedgwick, Genes Dev. 2000, 14,
2097-2105.
[34] A. B. Cardillo, M. Perassolo, M. Sartuqui, J. Rodríguez Talou, A. M.
Giulietti, Biochem. Eng. J. 2017, 125, 180-189.
This article is protected by copyright. All rights reserved.

10.1002/cbic.201900783
ChemBioChem
COMMUNICATION
Entry for the Table of Contents
COMMUNICATION
Maria Agustina Vila, Viktoria Steck,
Sonia Rodríguez, Ignacio Carrera* and
Rudi Fasan*
Page No. – Page No.
C—H Amination via Nitrene Transfer
Catalyzed by Mononuclear Non-Heme
Iron-Dependent Enzymes
New reactions for NHI enzymes: Various members of the non-heme iron
dependent oxidase family exhibit promiscous nitrene transferase activity
catalyzing the C—H amination of sulfonyl azides and carbonazidates. The Rieske
toluene dioxygenase was applied to carry out this non-native reactiton at a gram
scale in a bioreactor.
This article is protected by copyright. All rights reserved.