Identification of DypB from rhodococcus jostii RHA1 as a lignin peroxidase

Article abstract of DOI:10.1021/bi101892z

Rhodococcus jostii RHA1, a polychlorinated biphenyl-degrading soil bacterium whose genome has been sequenced, shows lignin degrading activity in two recently developed spectrophotometric assays. Bioinformatic analysis reveals two unannotated peroxidase genes present in the genome of R. jostii RHA1 with sequence similarity to open reading frames in other lignin-degrading microbes. They are members of the Dyp peroxidase family and were annotated as DypA and DypB, on the basis of bioinformatic analysis. Assay of gene deletion mutants using a colorimetric lignin degradation assay reveals that a dypB mutant shows greatly reduced lignin degradation activity, consistent with a role in lignin breakdown. Recombinant DypB protein shows activity in the colorimetric assay and shows Michaelis-Menten kinetic behavior using Kraft lignin as a substrate. DypB is activated by Mn²⁺ by 5-23-fold using a range of assay substrates, and breakdown of wheat straw lignocellulose by recombinant DypB is observed over 24-48 h in the presence of 1 mM MnCl₂. Incubation of recombinant DypB with a β-aryl ether lignin model compound shows time-dependent turnover, giving vanillin as a product, indicating that C_α- C_β bond cleavage has taken place. This reaction is inhibited by addition of diaphorase, consistent with a radical mechanism for C-C bond cleavage. Stopped-flow kinetic analysis of the DypB-catalyzed reaction shows reaction between the intermediate compound I (397 nm) and either Mn^II (k_obs = 2.35 s^-1) or the β-aryl ether (k _obs = 3.10 s^-1), in the latter case also showing a transient at 417 nm, consistent with a compound II intermediate. These results indicate that DypB has a significant role in lignin degradation in R. jostii RHA1, is able to oxidize both polymeric lignin and a lignin model compound, and appears to have both Mn^II and lignin oxidation sites. This is the first detailed characterization of a recombinant bacterial lignin peroxidase.

Full text of DOI:10.1021/bi101892z

ARTICLE
pubs.acs.org/biochemistry
Identification of DypB from Rhodococcus jostii RHA1 as a
Lignin Peroxidase
†
‡
†
‡
,‡
Mark Ahmad, Joseph N. Roberts, Elizabeth M. Hardiman, Rahul Singh, Lindsay D. Eltis,* and
,†
Timothy D. H. Bugg*
†
Department of Chemistry, University of Warwick, Coventry CV4 7AL, U.K.
‡
Department of Microbiology and Immunology, University of British Columbia, 2350 Health Sciences Mall, Vancouver,
British Columbia V6T 1Z3, Canada
S Supporting Information
b
ABSTRACT: Rhodococcus jostii RHA1, a polychlorinated biphenyl-degrading soil bacter-
ium whose genome has been sequenced, shows lignin degrading activity in two recently
developed spectrophotometric assays. Bioinformatic analysis reveals two unannotated
peroxidase genes present in the genome of R. jostii RHA1 with sequence similarity to open
reading frames in other lignin-degrading microbes. They are members of the Dyp
peroxidase family and were annotated as DypA and DypB, on the basis of bioinformatic
analysis. Assay of gene deletion mutants using a colorimetric lignin degradation assay reveals
that a ΔdypB mutant shows greatly reduced lignin degradation activity, consistent with a
role in lignin breakdown. Recombinant DypB protein shows activity in the colorimetric
assay and shows MichaelisꢀMenten kinetic behavior using Kraft lignin as a substrate. DypB
2
þ
is activated by Mn by 5ꢀ23-fold using a range of assay substrates, and breakdown of
wheat straw lignocellulose by recombinant DypB is observed over 24ꢀ48 h in the presence
of 1 mM MnCl . Incubation of recombinant DypB with a β-aryl ether lignin model
2
compound shows time-dependent turnover, giving vanillin as a product, indicating that
C ꢀC bond cleavage has taken place. This reaction is inhibited by addition of diaphorase,
R
β
consistent with a radical mechanism for CꢀC bond cleavage. Stopped-flow kinetic analysis
of the DypB-catalyzed reaction shows reaction between the intermediate compound I
II
ꢀ1
ꢀ1
(
397 nm) and either Mn (k = 2.35 s ) or the β-aryl ether (kobs = 3.10 s ), in the latter case also showing a transient at 417 nm,
obs
consistent with a compound II intermediate. These results indicate that DypB has a significant role in lignin degradation in R. jostii
II
RHA1, is able to oxidize both polymeric lignin and a lignin model compound, and appears to have both Mn and lignin oxidation
sites. This is the first detailed characterization of a recombinant bacterial lignin peroxidase.
9
,10
ignin is a heterogeneous aromatic polymer found as a major
component of lignocellulose in plant cell walls and is
mediators.
However, despite the study of fungal lignin-
L
degrading enzymes for 25 years, these discoveries have not
translated to a commercial process for lignin breakdown, in part
because of the inherent difficulties in fungal genetics and protein
expression. There are several literature reports of soil bacteria
that are able to metabolize lignin;
extremely resistant to chemical and biochemical breakdown.
The recalcitrance of the lignin component of lignocellulose is a
limitation in the utilization of plant biomass for cellulosic bio-
1
11,12
ethanol generation. Consequently, there is considerable interest
however, the enzymology
in the identification of novel organisms and catalysts for lignin
breakdown.
Lignin degradation has been studied primarily in white-rot and
of bacterial lignin degradation is not well characterized. Strepto-
myces viridosporus has been reported to produce extracellular
peroxidases, which can metabolize lignin model compounds.
13
14
brown-rot fungi, which produce extracellular peroxidases and
Using a radiochemical assay for lignin breakdown, lignin degra-
2
laccases that are able to metabolize lignin. The white-rot fungus
dation activity has been reported in strains of Nocardia auto-
12
Phanaerochaete chrysosporium produces a heme-containing lignin
trophica and Rhodococcus sp. However, no bacterial lignin-
degrading genes have been identified to date, and no recombi-
nant lignin-degrading enzymes have been characterized.
3
peroxidase (LiP) that is able to catalyze the cleavage of lignin
4
,5
model compounds and a heme-dependent manganese perox-
II
idase enzyme (MnP) that catalyzes the rapid oxidation of Mn to
III 6,7
III
Mn . Mn is then able to oxidize a range of lignin model
8
compounds. P. chrysosporium and a range of other fungi
Received: November 28, 2010
also produce extracellular copper-dependent laccase enzymes,
which are able to oxidize lignin model compounds, using redox
Revised:
Published: May 02, 2011
April 28, 2011
r 2011 American Chemical Society
5096
dx.doi.org/10.1021/bi101892z
|Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
and 23% for TyrA, 13 and 31% for btDyP, and 9 and 9% for
DyPDEC1, respectively. The resulting alignment was used as a
profile in CLUSTALX 2.0 to align DyP sequences. Initially, all
sequences deposited in the Peroxibase database and DyP family
sequences found in the literature were considered for this
analysis. Sequences were parsed such that none in the final data
set were more than 40% identical. Signal sequences, as predicted
Figure 1. Nitrated lignin assay for monitoring lignin degradation, as
described in ref 15.
23
with SignalP 3.0, were removed from the input sequences.
Sequences lacking any one of the conserved residues, Asp153,
His226, Arg244, and Asp288 (DypB numbering), or a glutamate
in place of an aspartate, were removed from the analysis. The final
data set had 39 sequences (Figure S1 of the Supporting Informa-
tion). For the phylogenetic analyses, the alignment was edited so
that it contained only equivalent residues. Accordingly, columns
were deleted if they contained insertions, deletions, or residues
that were not within 3.0 Å (root-mean-square deviation) of the
equivalent residues in each of the superimposed DyP structures.
This alignment (Figure S1 of the Supporting Information)
served as the input for a maximum-likelihood tree built using
We have recently reported two spectrophotometric assays for
lignin breakdown: an assay involving fluorescently modified
lignin and a UVꢀvis assay involving chemically nitrated lignin
(Figure 1), which gives changes in A4 upon lignin breakdown,
30
15
due to production of nitrated phenol products. Using this assay,
we have identified several bacterial strains that show lignin
degradation activity in their extracellular fraction, including two
known aromatic degraders, Rhodococcus jostii RHA1 and Pseudo-
15
monas putida mt-2. These two strains were shown to break
24
down lignocellulose to a low-molecular weight phenolic bypro-
PHYLIP version 3.69. Bootstrapping was performed using 100
data sets by jumbling the sequence input order 21 times for each
bootstrap and the best tree generated separately by jumbling
1
5
duct. R. jostii RHA1 is a degrader of polychlorinated biphenyls
1
6
(
PCBs). The availability of a genome sequence for this
17
organism and the ability to analyze gene function in detail in
this organism have prompted us to seek to elucidate the identity
of extracellular lignin-degrading enzymes in this organism. Here
we report the existence of DyP-type peroxidases DypA and DypB
in R. jostii RHA1, and the identification of DypB as a lignin
peroxidase, the first characterization of a recombinant bacterial
lignin peroxidase.
2
1 times.
DNA Manipulation and Plasmid Construction. The dyp
genes were amplified from fosmid clones RF0013J14 and
RF0013A20 containing appropriate fragments of RHA1 genomic
17
DNA (http://www.rhodococcus.ca ). The polymerase chain
reactions (PCRs) were conducted using Expand High-Fidelity
Polymerase (Roche), a Veriti 96-well Thermal Cycler (Applied
Biosystems), and the following primers: dypAFor, dypARev,
dypBFor, and dypBRev (Table S1 of the Supporting Informa-
tion). The amplicons were cloned into pET28a(þ) (Novagen)
using NdeI and HindIII, yielding pETDYPA1 and pETDYPB1 in
which dypA and dypB, respectively, are under the control of the
T7 promoter. The resulting genes encoded recombinant proteins
with a cleavable N-terminal poly-His tag (Ht-) and corresponded
to residues 50ꢀ429 of DypA (i.e., without the TAT signal
sequence) and residues 1ꢀ350 of DypB.
’
MATERIALS AND METHODS
Materials. The β-aryl ether lignin dimers 1ꢀ3 were synthe-
18
sized by the method of Nakatsubo et al., with the exception that
methyl chloroacetate was used instead of ethyl chloroacetate in
the first step. (þ)-Pinoresinol 4 was purchased from Arbo Nova
and used without further purification. O-(2-Hydroxyethyl)guaiacol
5
was synthesized by treatment of methyl carboxymethylguaiacol
0.25 g), an intermediate on the synthetic route mentioned
(
Construction of the mutagenic plasmid for ΔdypA and ΔdypB
genetic knockout strains was designed as described previously
1
8
above, with LiAlH (0.156 g) in tetrahydrofuran (10 mL) at
4
2
5
5
0 °C for 3 h, followed by addition of wet THF (5 mL), filtration,
using 800ꢀ1000 kb amplicons flanking the target gene. The
dypA and dypB upstream amplicons were prepared from RHA1
genomic DNA and the following primers: dAUF, dAUR, dBUF,
and dBUR (Table S1 of the Supporting Information). These
and evaporation of solvent, to give 5 as a yellow oil (0.225 g,
1
9
8%): H NMR (300 MHz, CDCl ) δ 6.9ꢀ7.0 (4H, m), 4.12
3
1
3
(2H, t, J = 5 Hz), 3.91 (2H, t, J = 5 Hz), 3.85 (3H, s); C NMR
0
(
7
75.5 MHz, CDCl ) δ 149.5, 148.0, 121.8, 121.1, 114.2, 111.8,
yielded a 790 bp amplicon that included the 5 90 bp of dypA and
3
þ
0
1.4, 61.2, 55.8; ESI-MS 191.0 (M Na ). Milled wood lignin was
1000 bp amplicon that included the 5 99 bp of dypB. The
19
prepared using a literature procedure. Kraft lignin, Clostridium
kluyveri diaphorase, and glucose oxidase were purchased from
Sigma-Aldrich. HPLC-grade DMSO was from Alfa Aesar. All other
reagents and chemicals were purchased from Sigma-Aldrich,
ACROS, MP Biomedicals, or Fisher and were used without further
purification. Enzymes for molecular cloning were purchased from
New England Biolabs.
downstream amplicons were similarly generated using dADF,
dADR, dBDF, and dBDR (Table S1 of the Supporting Informa-
tion). These yielded amplicons of 999 and 799 bp, respectively,
0
which included the 3 99 bp of the respective genes. For dypA or
dypB, a three-fragment ligation was performed to clone the
flanking amplicons into pK18mobsacB, yielding pK18dpA or
pK18dpB, respectively. Primer synthesis and nucleotide sequen-
cing of the final clones were performed at the Nucleic Acid-
Protein Services Unit (University of British Columbia).
Phylogenetic Analyses. A structure-based alignment of DyPs
2
0,21
22
was generated using STRAP
and CLUSTALX2.0. Briefly,
20,21
the STRAP editor and the TM-Align algorithm
were used to
Construction of Mutants. The ΔdypA and ΔdypB deletion
mutants of RHA1 were constructed essentially as described
align the four DyPs with published structures: EfeB of Escherichia
coli [Protein Data Bank (PDB) entry 2WX6], BtDyp of Bacter-
oides thetaiotaomicron VPI-5482 (PDB entry 2GVK), TyrA of
Shewanella oneidensis (PDB entry 2HAG), and DyPDec1 (PDB
entry 2D3Q). The sequence identities of the DypB homologues
to R. jostii RHA1 DypA and DypB are 27 and 13% for EfeB, 14
25
previously. Wild-type RHA1 was grown in Lysogeny Broth
Peptone medium (LBP) containing 30 μg/mL nalidixic acid at
30 °C for 72 h with shaking. The cultures were subsequently
plated on solid LBP agar and incubated for 72 h at 30 °C followed
by incubation for 24 h at room temperature. pK18dpA and pK18dpB
5
097
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
were transformed into E. coli S17-1 and grown for 24 h on LBP
plates supplemented with 25 μg/mL kanamycin at 37 °C. Cells
from both the RHA1 and transformed E. coli S17-1 plates were
resuspended in 2 mL of LBP, and 750 μL aliquots of both strains
were mixed carefully. The cell mixture was harvested by cen-
trifugation (9300g for 1 min) and resuspended in 1 mL of LBP,
and 100 μL aliquots were spread on LBP agar and incubated at
buffer A containing 1 mM EDTA and were loaded onto 8 mL of
Source 15Q anion exchange resin (GE Healthcare) packed in an
AP-1 column (Waters Corp., Milford, MA) that had been
equilibrated with buffer A and was operated at a flow rate of
3 mL/min. DyPs were eluted using a linear gradient from 80 to
500 mM NaCl in 20 column volumes. The purified protein,
which was >95% apparently homogeneous as determined by
sodium dodecyl sulfateꢀpolyacrylamide gel electrophoresis, was
exchanged into buffer A, concentrated to 20 mg/mL, frozen as
beads in liquid nitrogen, and stored at ꢀ80 °C until it was used.
UVꢀVisible Assays. A stock solution of nitrated MWL here-
ward wheat lignin (0.015 mM) was prepared in 750 mM Tris
buffer (pH 7.4) containing 50 mM NaCl. Assays (total volume of
200 μL) were conducted in 96-well Falcon Microtest clear plates,
using a TECAN GENios plate reader. To each well were added
30 μL of extracellular cell extract or recombinant protein solution
(0.1 mg/mL), 160 μL of nitrated lignin, 10 μL of 40 mM H O .
3
0 °C for 24 h to facilitate conjugation. Biomass was harvested
from the plates by resuspension in 2 mL of LBP, and 25 μL
aliquots were spread on LBP agar supplemented with 50 μg/mL
kanamycin and 30 μg/mL nalidixic acid and incubated at 30 °C
to initiate the integration of the pK18mobsacB plasmid via a
single homologous recombination event. After 72 h, kanamycin
r
resistant colonies (Kan ) were replica plated on LBP supplemen-
ted with 10% (w/v) sucrose and LBP supplemented with
5
0 μg/mL kanamycin. Both plates contained 30 μg/mL nalidixic
r
r
s
acid. Kan and sucrose sensitive (Kan /Suc ) colonies were
cultured in 25 mL of LBP overnight at 30 °C, and 25 μL aliquots
were plated on LBP agar supplemented with 10% (w/v) sucrose.
Plates were incubated for 72 h at 30 °C; sucrose resistant (Suc )
colonies were grown for 24 h at 30 °C following replica plating
on LBP agar and LBP agar supplemented with 50 μg/mL
2
2
The absorbance at 430 nm was measured at 1 min intervals for
20 min. Each assay was conducted in duplicate, with controls in
which a nitrated lignin or protein solution was replaced with
750 mM Tris (pH 7.4) containing 50 mM NaCl. Assays of
recombinant DypA and DypB were also conducted in the presence
r
s
kanamycin. Kanamycin sensitive (Kan ) colonies were screened
of 30 μL of 50 mM MnCl . The whole plate was repeated without
2
by colony PCR for truncated dypA and dypB gene products.
Bacterial Strains and Growth. Ht-DypA and DypB were
produced in E. coli BL21(DE3). Cells containing pETDYPA1 or
addition of H O . All readings were taken in duplicate.
Kinetic Analysis of DypB with Kraft Lignin. In a 1 mL
cuvette, Kraft lignin (500 μL, 0.5 mg/mL), succinate buffer
2
2
26
pETDYPB1 were grown in lysogeny broth (LB) containing
5 μg/mL kanamycin at 37 °C with shaking. Cultures were grown
to an OD of 0.5, at which point isopropyl β-D-thiogalactopyr-
(175 μL, pH 5.5, 50 mM), MnCl (200 μL, 50 mM), DypB
2
2
(25 μL, 1 mg/mL), and hydrogen peroxide (100 μL, 40 mM)
were mixed in the order stated, and the absorbance was mon-
itored for 5ꢀ10 min at 465 nm. Substrate concentrations were
6
00
anoside (IPTG) was added to a final concentration of 0.5 mM to
induce dyp expression. Cells were incubated for a further 12 h at
then varied for MnCl (final concentration of 1.25ꢀ10 mM) and
2
2
0 °C while being shaken before being harvested by centrifuga-
Kraft lignin (final concentration of 0.05ꢀ0.25 mg/mL), with the
total volume kept at 1.0 mL. The molar concentration of Kraft
lignin was calculated using an average molecular mass of 10000 Da.
Incubation of DypB with Lignocellulose and Lignin. Pow-
dered wheat straw lignocellulose or milled wood lignin (5 mg)
was added to succinate buffer (3 mL, 50 mM, pH 5.5), and
then DypB (100 μL, 1 mg/mL) was added, followed by H O
tion. Pellets were washed three times with 20 mM sodium
phosphate and 10% glycerol (pH 8.0) and frozen at ꢀ80 °C
until they were used.
Protein Purification. Unless otherwise stated, the purification
protocol for the DyPs was identical. Approximately 4.5 g of cells
(wet weight) was thawed in 20 mL of 20 mM sodium phosphate
2
2
and 0.1 mM EDTA (pH 8.0) and lysed at 4 °C using an Emulsi
Flex-C5 homogenizer (Avestin). Cell debris was removed by
centrifugation (140000g for 50 min). Ht-DyP was purified from
the supernatant using a column with 10 mL of Ni-NTA resin
(100 mL, 40 mM). The resulting solution was incubated at 30 °C
for 48 h. Aliquots (1 mL) were taken at 1, 3, 5, 24, and 48 h. These
fractions were analyzed by HPLC. Aliquots for HPLC (400 μL)
were mixed with CCl COOH (100%, w/v, 40 μL), and the solu-
3
(QIAgen) according to the manufacturer’s instructions in the
tion was then centrifuged for 4 min at 10000 rpm. The experi-
ment was also repeated with hydrogen peroxide replaced with
glucose (final concentration of 0.3 mM) and glucose oxidase
(final concentration of 0.5 mg/mL). Both experiments were
presence of 0.1 mM EDTA. DyP was eluted as the apoprotein
from the column and was exchanged into 20 mM 3-(N-morpho-
lino)propanesulfonic acid (MOPS) and 80 mM NaCl (pH 7.5)
(buffer A) using a 30K NMWL Amicon Ultra-15 Centrifugal
conducted with and without MnCl (1 mM). HPLC analysis was
2
Filter Device (Millipore). The affinity tag was removed by
incubating the protein with a 200:1 molar ratio of human
R-thrombin (Haematologic Technologies Inc.) for 12 h at room
temperature, followed by a second IMAC column. The DyPs
were eluted from the second IMAC column using buffer A
containing 20 mM imidazole and subsequently buffer exchanged
into buffer A. As purified, preparations of DypA and DypB had
Rz values (ASoret/A280) of <0.1. To reconstitute the proteins,
hemin chloride was dissolved in DMSO to a final concentration
of 20 mg/mL and added dropwise with gentle stirring to protein
in buffer A to a final 2:1 hemin:protein molar ratio. Excess hemin
was removed via centrifugation followed by gel filtration chro-
matography using Sephadex G-25 fine (GE Healthcare). Recon-
stituted DypA and DypB had Rz values of 2.7 and 2.9, respec-
tively. The reconstituted DyPs were dialyzed overnight at 4 °C in
conducted using a Phenomenex Luna 5 μm C18 reverse phase
column (100 Å, 50 mm ꢁ 4.6 mm) on a Hewlett-Packard Series
1100 analyzer, at a flow rate of 0.5 mL/min, with monitoring at
310 nm. The gradient was as follows: 20 to 30% MeOH/H O
2
over 5 min, 30 to 50% MeOH/H O from 5 to 12 min, and 50 to
2
80% MeOH/H O from 12 to 25 min.
2
Reaction of DypB with Lignin Model Compounds. Samples
of lignin model compounds (all three types of β-aryl ether,
pinoresinol) (2 mL, 5 mM solution in a 9:1 acetone/water
mixture) were diluted with succinate buffer (3 mL, 50 mM, pH
2
7
5.5), and then DypB (25 μL, 1 mg/mL) and MnCl (100 μL,
2
50 mM) were added, followed by H O (100 μL, 40 mM). The
2
2
resulting solution was incubated at 30 °C for 48 h. Aliquots
(1 mL) were taken at 1, 3, 5, 24, and 48 h. These fractions were
analyzed by HPLC, gas chromatography and mass spectroscopy
5
098
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
Figure 2. Operonic structure of dypA and dypB in R. jostii RHA1.
(
(
GCꢀMS), and liquid chromatography and mass spectroscopy
LCꢀMS). Aliquots for HPLC (400 μL) were mixed with
Table 1. Annotation of DyPs and Proteins Encoded by
Predicted Cotranscribed Genes in the RHA1 Genome
CCl COOH (100%, w/v, 40 μL) and centrifuged (13000 rpm
3
a
BeTs to characterized percent identity
for 5 min) prior to analysis. Samples for GCꢀMS (200 μL) were
extracted into HPLC-grade ethyl acetate (1 mL) and then N,O-
bis(trimethylsilyl)acetamide (200 μL) and (chlorotrimethlyl)-
silane (10 μL). The solution was left for 1 h and then diluted 100-
fold in the same solvent prior to injection. For each model
compound, a control in which no DypB was added was run. In
the case of β-aryl ether 2, the hydrogen peroxide was also
replaced with glucose oxidase (final concentration of 0.5 mg/mL)
and glucose (final concentration of 0.3 mM), and the reaction
b
gene ID
ro02407
annotation
DypB
homologues
(%)
BtDyp of Bacteroides
thetaiotaomicron VPI-5482
Encapsulin of Thermotoga
maritima
41
ro02408
encapsulin
DypA
34
ro05773
ro05774
EfeB of E. coli K-12
EfeO of E. coli K-12
EfeU of E. coli K-12
32
35
27
lipoprotein
II
II
ro05775 Fe /Pb permease
was repeated without MnCl . This reaction was repeated in the
2
a
b
Best hits from a local alignment of the NCBI database. Based on full-
presence of C. kluyveri diaphorase (0.5 mL, 2 mg/mL), with or
without NADH (2 mL, 5 mM stock). LCꢀMS analysis was
conducted using a Phenomenex Luna 5 μm C18 reverse phase
column (100 Å, 50 mm ꢁ 4.6 mm) on an Agilent 1200
instrument, at a flow rate of 0.5 mL/min, with a Bruker HCT
Ultra mass spectrometer. The LC gradient was as follows: 20 to
length alignment.
Zymomonas mobiliz (GenBank accession number Q5NM63,
5
5% identical).
The gene identified in R. jostii RHA1 (ro02407) encodes a
3
1
0% MeOH/H O over 5 min, 30 to 50% MeOH/H O from 5 to
2
2
350-amino acid peroxidase (M = 37222), a member of the DyP
r
2 min, and 50 to 80% MeOH/H O from 12 to 25 min.
2
peroxidase family, that shows high activity toward dye substrates
2
8ꢀ30
Stopped-Flow Kinetic Analysis. Kinetic measurements were
and contains a distal aspartic acid proton donor.
Two
taken with a MOS 450-Biologic spectrophotometer and an SFM
putative DyPs were encoded in the RHA1 genome. These were
named DypA and DypB on the basis of the phylogenetic
analysis below. To gain insight into the identity and function of
the predicted RHA1 DyPs, we employed local alignments to
explore the dyp genes and their genomic context (Figure 2). The
predicted DypA protein is 32% identical in amino acid sequence
to EfeB from E. coli K-12, also known as YcdB in strain O157:
H7 (Table 1). In E. coli, this gene belongs to the efeUOB
31
3
00 observation chamber equipped with a 1 cm observation cell
at 25 °C, monitoring at a fixed wavelength. Recombinant DypB
(
(
8 μM) was prepared in 30 mM citrate/60 mM phosphate buffer
pH 6.0) and mixed with freshly prepared solutions of hydrogen
peroxide, MnCl , and β-aryl ether 2 at 8 or 80 μM. All measure-
2
ments were taken in triplicate. The decay of DypB compound I
was observed at 397 nm (DOI 10.1021/bi200427h). The decay
of compound II was observed at 417 nm, which is the isosbestic
2
þ
operon involved in Fe uptake under acidic conditions.
Consistent with this finding, EfeB has a twin-arginine transport
TAT) signal sequence, EfeO possesses an N-terminal cupre-
doxin domain, and EfeU is a homologue of the high-affinity iron
7
point between compound II and the resting enzyme. Kinetic
transients were fitted to single- and double-exponential kinetic
models, and in most cases, good fits to single-exponential func-
tions were observed.
(
3
2
permease FTR1 from Saccharomyces cerevisiae. The efeUOB
operon is conserved in RHA1 (Figure 2 and Table 1), and DypA
is predicted to contain a TAT signal sequence, suggesting that it
is exported.
’
RESULTS
Identification of dyp Genes by Bioinformatic Analysis. As a
Among characterized homologues, DypB is 41% identical in
starting point for bioinformatic analysis, we examined the
genome of Nocardia farcinica, closely related to Nocardia auto-
trophica, which has been found to show activity for lignin degra-
amino acid sequence to BtDyP, whose crystal structure has been
30
determined. The dypB gene is predicted to be operonically
coupled with ro02408, annotated here as enc as its gene product is
34% identical to encapsulin from T. maritima, which forms a
15,16
dation,
for peroxidase genes of unknown function. N. farcinica
11
contains three unannotated peroxidase genes (GenBank accession
numbers Q5YV75, Q5YZF4, and Q5YPL4). Searches using the
BLAST algorithm revealed that Q5YPL4 shared homologues in
Streptomyces coelicolor (GenBank accession number Q9FBY9, 66%
identical), Mycobacterium tuberculosis (GenBank accession number
A0R4G9, 63% identical), Pseudomonas fluorescens (GenBank
accession number Q4KA97, 56% identical), R. jostii RHA1
cellular nanocompartment (Figure 2). Moreover, the RHA1
DypB is predicted to contain the C-terminal sequence respon-
33
sible for targeting proteins for encapsulation.
Phylogenetic Analysis. The four DyP structures available in
the PDB were superimposed using STRAP to guide the sequence
alignment. The associated TM-score for the four aligned
sequences exceeded 0.75, with overall root-mean-square devia-
tion (rmsd) values ranging from 3.1 to 3.2 Å, consistent with the
(
GenBank accession number Q0SE24, 52% identical), and
5
099
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
Figure 3. Radial phylogram of DyPs. The names of bacterial or fungal strains are indicated in subscript with the protein or abbreviated organism names.
Structure-based sequence alignments were performed using EfeBK-12 from E. coli K-12 (2WX6), TyrA from S. oneidensis (2HAG), BtDyPVPI-5482 from B.
thetaiotaomicron VPI-5482 (2GVK), DyPDEC1 from Thanatephorus cucumeris DEC1 (2D3Q), DypARHA1 and DypBRHA1 from R. jostii RHA1
(
ABG97551.1 and ABG94212.1, respectively), YfeXK-12 from E. coli K-12 (BAE76711.1), AnaPXPCC 7120 from Anabaena sp. PCC 7120 (BAB77951.1),
TfuDypYX from Thermobifida fusca YX, MsP1 from Marasmius scorodonius, TalTAP from Termitomyces albuminosus, Bsu168 from Bacillus subtilis 168
CAB15852.1), MvaPYR-1 from Mycobacterium vanbaalenii PYR-1 (ABM12972.1), PdePD1222 from Paracoccus denitrificans PD1222 (ABL69832.1),
(
RpABisB18 from Rhodopseudomonas palustris BisB18 (ABD87513.1), CteKF-1 from Comamonas testosteroni KF-1 (EED66859.1), DdiAX4 from
Dictyostelium discoideum AX4 (EAL70759.1), MtbH37Rv from M. tuberculosis H37Rv (CAB09574.1), Cco13826 from Campylobacter concisus 13826
(EAT98288.1), AcspADP1 from Acinetobacter sp. ADP1 (CAG67144.1), PpaSIR-1 from Plesiocystis pacifica SIR-1 (EDM76509.1), CyspPCC 7424 from
Cyanothece sp. PCC 7424 (ACK71272.1), CviATCC 12472 from Chromobacterium violaceum ATCC 12472 (AAQ59612.1), MxaDK 1622 from Myxococcus
xanthus DK 1622 (ABF90727.1), OanATCC 49185 from Ochrobactrum anthropi ATCC 49185 (ABS17389.1), PsspPRwf-1 from Psychrobacter sp. PRwf-1
(
ABQ94167.1), ChuATCC 33406 from Cytophaga hutchinsonii ATCC 49185 (ABG59511.1), Pos from Pleurotus ostreatus (CAK55151.1), PgrCRL from
Puccinia graminis CRL (AAWC01001299.1), Ppa from Phakopsora pachyrhizi, Lbi1S238N-H82 and Lbi2S238N-H82 from Laccaria bicolor S238N-H82
ABFE01001782.1 and EDR12662.1, respectively), BfuB05.10 from Botryotinia fuckeliana B05.10 (EDN26366.1), PchWisconson 54ꢀ1255 from Penicillium
chrysogenum Wisconson 54-1255 (CAP99029.1), AfuAf293 from Aspergillus fumigatus Af293 (EAL86784.2), PplMad-698-R from Postia placenta Mad-698-R
EED79944.1), and Amsp1 and Amsp2 from Amycolatopsis sp. (personal communication with M. Chang, University of California, Berkeley, CA). As
(
(
described in Materials and Methods, the structure-based alignment included only equivalent residues. The tree was calculated using protein maximum
likelihood. Bootstrap values of critical nodes are indicated.
shared ferredoxin-like structural fold of these proteins. The TM-
align superposition further indicated that the four DyPs have 234
equivalent residue positions. After the alignment of the parsed set
of collected DyP sequences with the structural alignment profile,
a total of 39 sequences were retained in the final alignment
Using a maximum likelihood analysis, the DyP sequences were
grouped into four discrete subfamilies (Figure 3). These were
3
5
identified as AꢀD to be consistent with a previous analysis.
The four DyPs for which structural data are available belong to
subfamilies A, B, and D. Subfamilies A and B, which are grouped
within the phylogram, comprise bacterial DyPs. Subfamilies C
and D, which are also grouped together, comprise bacterial and
fungal sequences, respectively. The most divergent of the ana-
lyzed DyPs are 6% identical in sequence. Within subfamilies, this
value is 15%. RHA1 DypA and DypB were named according to
the subfamilies in which they were grouped.
(
Figure S1 of the Supporting Information). For the phylogenetic
analyses, only the structurally equivalent residues were main-
tained in the alignment, corresponding to an average of 55% of
each sequence (Figure S1 of the Supporting Information).
Chlorite dismutase, a heme-containing enzyme with the same
structural fold as DyP, was excluded from the alignment as
the TM-scores were below 0.6, the overall rmsd values ranged
from 3.5 to 3.8 Å, and the enzyme lacks the distal aspartate of
3
0
Analysis of Gene Deletion Mutants. Deletion mutants of
R. jostii RHA1 in genes dypA and dypB were prepared, using the
3
4
25
DyPs.
method of van der Geize et al. The culture supernatant from
5
100
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
Methods. Both proteins were found to exhibit peroxidase activity
using ABTS as a substrate (DOI 10.1021/bi200427h). Using
ABTS as a substrate, the optimal activity for DypB was observed
at pH 3.6, but activity was retained at pH 5ꢀ6. Each protein was
15
tested in the nitrated lignin UVꢀvis assay, in the presence and
absence of 2 mM hydrogen peroxide. As shown in Figure 5,
DypB exhibited activity in the presence of hydrogen peroxide,
but not in the absence of hydrogen peroxide. Under the same
conditions, DypA exhibited no activity, in the presence or
absence of hydrogen peroxide. These data therefore support
the conclusions of the earlier gene knockout experiments,
implying that DypB shows activity as a lignin peroxidase but
DypA does not. DypB was found to show 5.4-fold greater activity
in this assay in the presence of 1 mM MnCl (see Table 2),
2
2
þ
suggesting that Mn acts as a cofactor or mediator for DypB.
Using the ABTS assay, DypB showed 8.5-fold higher activity in
the presence of 1 mM MnCl (see Table 2), and a specific activity
2
ꢀ
1
ꢀ1
of 4.8 μmol min (mg of protein) was calculated using ABTS
as a substrate.
To evaluate further its lignin-degrading activity, recombinant
DypB was then assayed against samples of lignin and lignocellu-
lose. When 25 μg of recombinant DypB was incubated at 30 °C
with 5 mg of wheat straw lignocellulose in 50 mM succinate
buffer (pH 5.5) for 48 h, in the presence of either 2 mM hydrogen
peroxide or 0.3 mM glucose and 1.5 mg of glucose oxidase, no
changes were observed in the reverse phase HPLC chromato-
grams of the culture supernatant. However, incubation of DypB
in the presence of 1 mM MnCl with wheat straw lignocellulose,
2
0
.3 mM glucose, and 1.5 mg of glucose oxidase led to time-
dependent changes in the reverse phase HPLC chromatograms
of the aliquots that had been removed. As shown in Figure 5, a
peak at 17 min disappeared in 1ꢀ3 h, while peaks at 19.8 and 21.8
min disappeared over 24ꢀ48 h, consistent with breakdown.
Although the identity of these peaks is not known, the time-
Figure 4. Activities of R. jostii RHA1 gene deletion strains and DypA
and DypB recombinant enzymes using the nitrated lignin UVꢀvis assay.
dependent changes were not observed in the absence of MnCl or
2
DypB; moreover, similar time-dependent changes are observed
(
A) Activity (change in absorbance at 430 nm) in the nitrated lignin
upon incubation of R. jostii RHA1 with lignocellulose, and no
UVꢀvis assay of the culture supernatant obtained from wild-type RHA1
and ΔdypB and ΔdypA gene deletion mutants, in the presence (white)
and absence (gray) of 2 mM hydrogen peroxide. (B) Activity (change in
absorbance over a 20 min assay) of 25 μg of recombinant DypA and
DypB protein in the nitrated lignin UVꢀvis assay, in the presence
15
changes are observed in the presence of nondegradingBa. subtilis.
A similar incubation experiment conducted using wheat straw
milled wood lignin as a substrate also resulted in time-dependent
changes by reverse phase HPLC over 24ꢀ48 h (data not shown).
These experiments indicate that recombinant DypB can, under
certain conditions, react with a polymeric lignin substrate.
(white) or absence (gray) of 2 mM hydrogen peroxide.
the mutant strains was examined for lignin degradation activity
DypB was also incubated with samples of commercially
available Kraft lignin, an industrially produced byproduct of
paper manufacturing. When Kraft lignin was incubated in the
15
using the previously described nitrated lignin UVꢀvis assay.
Although this assay shows some experimental scatter using cell
supernatant, we have previously found that lignin-degrading
strains show an increase in absorbance at 430 nm, relative to
presence of 1ꢀ10 mM MnCl
, monitoring of the reaction by
2
UVꢀvis spectroscopy versus time revealed a time-dependent
change in the UVꢀvis spectrum at 465 nm in a 10 min assay.
Control experiments in the absence of lignin gave a <10% change
in absorbance, indicating that the observed change was due to
15
controls, whereas nondegraders show a zero or negative value.
As shown in Figure 4, wild-type R. jostii RHA1 shows hydrogen
15
peroxide-dependent activity, as observed previously, whereas
the ΔdypB strain shows no observable activity in the presence or
absence of hydrogen peroxide and a significant decrease in
activity compared to that of the wild-type strain. The ΔdypA
strain exhibited a slightly lower activity than the wild-type strain.
These data provide some evidence of a role for the dypB gene in
the ability of R. jostii RHA1 to break down lignin but are incon-
clusive with regard to the role of the dypA gene.
II
III
reaction with lignin, rather than oxidation of Mn to Mn . Rate
measurements at this wavelength at a range of MnCl and Kraft
2
lignin concentrations were found to follow saturation kinetics
(Figure S2 of the Supporting Information). The deduced appar-
ent K
10000 for commercial Kraft lignin, an apparent K
value for MnCl is 8.4 mM. Using the average M
value of
value of
m
2
r
m
20 μM can be calculated for Kraft lignin; however, this value may
not be meaningful, because the substrate is heterogeneous, and
the enzyme may interact with only a part of the lignin substrate.
To examine the reaction of DypB with different possible lignin
substructures, we then assayed recombinant DypB against a
Kinetic Evaluation of Recombinant DypA and DypB.
Recombinant DypA and DypB proteins were expressed in
E. coli as His fusion proteins, purified by affinity chromatogra-
6
phy, and reconstituted with heme, as described in Materials and
5
101
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
Figure 5. Changes vs time in the reverse phase HPLC chromatogram of an incubation of recombinant DypB with wheat straw lignocellulose. Twenty-
five micrograms of recombinant DypB was incubated at 30 °C with 5 mg of wheat straw lignocellulose in 50 mM succinate buffer (pH 5.5) for 48 h, in the
presence of 0.3 mM glucose and 1.5 mg of glucose oxidase, and 1 mM MnCl
and Methods, with UV detection at 310 nm.
2
; aliquots were analyzed by reverse phase HPLC, as described in Materials
Table 2. Activation of DypB by MnCl , Using Different Steady-State Assay Methods, with Polymeric or Small Molecule Substrates
2
without MnCl
2
with MnCl
2
substrate
assay method
with substrate
without substrate
<0.001
with substrate
without substrate
x-fold activation
a
nitrated lignin
Kraft lignin
ABTS
UVꢀvis
0.0015
0.039
0.0081
0.24
0.0025
0.010
5.4
6.2
8.5
23
b
f
UVꢀvis
ND
c
UVꢀvis
0.0065
0.0023
<0.0005
0.0549
0.053
<0.002
d
f
f
β-aryl ether 2
lignocellulose
HPLC
ND
ND
e
HPLC
0% after 24 h
100% after 24 h
. Change in absorbance at 465 nm in a 10 min assay, with 3 μg
. Change in absorbance per minute at 430 nm, with 1 μg of protein, 0.5 mM MnCl , and 10 mM ABTS in 50 mM sodium
acetate buffer (pH 5.0). Disappearance of the HPLC peak vs time (millimoles per cubic decimeter per hour), with 2 μg of protein and 1 mM MnCl .
a
b
Change in absorbance at 430 nm in a 20 min assay, with 1 μg of protein and 1 mM MnCl
2
c
of protein and 1.5 mM MnCl
2
2
d
2
e
2
Percent disappearance of the 17 min HPLC peak after incubation for 24 h, with 100 μg of protein in 50 mM succinate buffer (pH 5.5) and 1 mM MnCl .
Not determined.
f
series of lignin model compounds, synthesized according to
incubation of β-aryl ether 2 with DypB were analyzed by
established literature methods (see Materials and Methods), in
the presence of 2 mM hydrogen peroxide. Three different β-aryl
ether model compounds were tested at a concentration of 2 mM
LCꢀMS. A broad envelope of product peaks was observed,
found by LCꢀMS to give peaks in the range of m/z 400ꢀ800,
higher than the molecular weight of the starting material 2,
therefore indicating the likelihood that recombination of radical
products was taking place, to give higher-molecular weight
species. The expected C ꢀC bond cleavage reaction, shown
(
the p-hydroxy analogue 1, the guaicyl analogue 2, and the
syringyl analogue 3), as well as pinoresinol 4. Each substrate
was incubated with DypB at a concentration of 2 mM, and the
reaction was analyzed by reverse phase HPLC over 1ꢀ5 h.
Gradual consumption of guaicyl β-aryl ether 2 was observed, to
the extent of 20ꢀ30% peak height over 5 h (Figure S3 of the
Supporting Information), but no turnover was detected with the
other substrates. By monitoring the rate of disappearance of 2 by
HPLC versus time, we determined specific activities for the
R
β
in Figure 6, should release guaiacol as a product: no guaiacol was
observed directly; however, a peak at a retention time of 27 min
showed an ion at m/z 407, consistent with a guaiacol trimer
þ
(MK 407), and the same species was formed upon incubation
of guaiacol with DypB.
To probe the existence of radical intermediates in the mechan-
ism of this transformation, the reaction of DypB with 2 was
repeated in the presence of C. kluyveri diaphorase, known to be
ꢀ
ꢀ1
1
erythro and threo isomers of 2. Values of 0.017 μmol min (mg
of protein) for the erythro isomer and 0.005 μmol min (mg
of protein) for the threo isomer were calculated.
ꢀ
1
ꢀ
1
36
involved in free radical defense. In the presence of 0.4 mg/mL
Investigation of Reaction Products of Recombinant DypB
with β-Aryl Ether 2. The turnover of β-aryl ether 2 by DypB
allowed a more detailed study of the molecular site of action and
catalytic mechanism of DypB. The reaction products from
diaphorase and 0.8 mM NADH, no reaction products were
observed, implying that the cleavage reaction had been completely
inhibited. In the presence of 0.4 mg/mL diaphorase but with-
out added NADH, several additional products were observed.
5
102
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
Figure 6. A. Structures of lignin model compounds tested as substrates for DypB. B. Expected and observed products from C
β-aryl ether 2 by DypB. HPLC and LCꢀMS data for reaction of DypB with β-aryl ether 2 are shown in Figures S3ꢀS7 of the Supporting Information.
R
ꢀC
β
bond cleavage of
Selected ion monitoring at m/z 153 for product vanillin revealed
a small peak with a retention time of 28.4 min, with the same
þ
retention time and molecular ion (m/z 153.2 MH ) as an
authentic sample of vanillin. The observed peak for vanillin
was small; however, separate studies have shown that vanillin is
itself oxidized by peroxidase DypB (see below). A peak was
observed with a retention time of 28.6 min at m/z 191.0, which
matched the expected molecular weight of the reduced C ꢀC
R
β
bond fragment 5, whose structure is shown in Figure 7. An
authentic standard for this alcohol was synthesized by LiAlH4
reduction of methyl carboxymethyl guaiacol (see Materials and
MethodsExperimental Procedures), and the standard gave a
retention time and a mass spectrum identical to those of the
peak formed by DypB and 2 (Figure S4 of the Supporting
Information). The guaiacol trimer was again observed in this
experiment. Therefore, LCꢀMS data have been obtained to
support each of the expected cleavage products (see Figure 7)
formed by C ꢀC bond cleavage of 2 by DypB.
R
β
II
Role of Mn in DypB Catalysis. We have observed using
2
þ
several different assays that DypB activity is enhanced by Mn
see Table 2). By analogy with the fungal lignin-degrading
peroxidases, DypB might therefore be a manganese peroxidase,
(
II
III 6,7
whose primary role is to oxidize Mn to Mn . A third class of
peroxidase, typified by Pleurotus eryngii versatile peroxidase,
II
37
oxidizes Mn and lignin at comparable rates. To distinguish
between these possibilities, the reaction rates of DypB-catalyzed
II
oxidation of Mn versus lignin must be compared directly, which
can be achieved using stopped-flow UVꢀvis spectrophotometry.
In the following paper (DOI 10.1021/bi200427h), a stable
compound I-like intermediate species has been characterized
for DypB, with a λmax of 397 nm. Therefore, the rates of oxidation
Figure 7. Catalytic cycle of DypB, showing pathways and rate constants
for turnover of β-aryl ether 2, to generate vanillin, or reaction with Mn .
II
5
103
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
a
Table 3. Stopped-Flow Kinetic Data for DypB, Recorded at 397 nm (compound I) or 417 nm (compound II)
397 nm
417 nm
ꢀ
1
2
ꢀ1
2
conditions
amplitude
k
obs (s ) (χ )
amplitude
kobs (s ) (χ )
3
ꢀ4
ꢀ4
ꢀ4
ꢀ4
ꢀ4
ꢀ4
ꢀ3
1.01 (4.3 ꢁ 10ꢀ4
1.18 (5.7 ꢁ 10ꢀ4
1
1
1
1
1
1
:1 DypB:H
2
O
2
7.4 ꢁ 10ꢀ
0.84 (5.0 ꢁ 10
2.35 (4.2 ꢁ 10
3.10 (5.9 ꢁ 10
2.81 (5.2 ꢁ 10
1.99 (4.9 ꢁ 10
2.11 (6.2 ꢁ 10
0.38
)
)
)
)
)
)
2.3 ꢁ 10
b
)
ꢀ3
ꢀ3
ꢀ3
ꢀ3
ꢀ3
:1:1 DypB:H
2
O
2
:MnCl
:β-aryl ether
:MnCl :β-aryl ether
:MnCl
:β-aryl ether
2
6.3 ꢁ 10
6.1 ꢁ 10
7.6 ꢁ 10
8.3 ꢁ 10
6.3 ꢁ 10
:1:1 DypB:H
2
O
2
7.5 ꢁ 10ꢀ3
)
:1:1:1 DypB:H
2
O
2
2
:1:10 DypB:H
2
O
2
2
:1:10 DypB:H
2
O
2
ꢀ
3
ꢀ
1.8 ꢁ 10
a
Experiments were conducted as described in Meterials and Methods, using 8 μM DypB and 1 or 10 equiv of hydrogen peroxide, MnCl , or β-aryl ether
2
2
, at pH 6.0 and 25 °C. In each case, the transient kinetic data were fit to a single exponential (except the final row) where the data gave a better fit to a
b
two-step process. The individual data sets are shown in Figures S10ꢀS15 of the Supporting Information. Transient not observed.
II
of β-aryl ether 2 and Mn by DypB were measured via stopped-
flow UVꢀvis spectrophotometry.
ponding enzymes, and possible protein engineering and pathway
engineering approaches for lignin breakdown; hence, we have
undertaken the identification of lignin-degrading enzymes in
R. jostii RHA1.
A series of stopped-flow experiments were conducted with
recombinant DypB, mixing 8 μM enzyme in citrate/phosphate
buffer (pH 6.0) with 1 equiv of hydrogen peroxide and 1 or 10
We have used a bioinformatics approach to identify two DyP-
type peroxidases in R. jostii RHA1 that have homologues in other
lignin-degrading bacteria, for which spectroscopic and kinetic
characterization is presented in the following paper (DOI
10.1021/bi200427h). The phylogenetic analyses of DyP perox-
idases confirm and extend previous studies. Using only a
sequence-based alignment and neighbor joining methods, DyPs
equiv of MnCl and/or β-aryl ether 2, and monitoring the
2
reaction either at 397 nm (for compound I) or 417 nm (for
compound II), and the data are listed in Table 3. When the
sample was mixed with only hydrogen peroxide, a first-order
decay of compound I (kobs = 0.84 s ) was observed at 397 nm,
because of the breakdown of compound I. Addition of 1 equiv of
either MnCl (k = 2.35 s ) or β-aryl ether 2 (kobs = 3.10 s
gave first-order transients with higher kobs values, consistent with
a reaction of DypB compound I with either Mn or lignin model
compound 2. No significant change in kobs was found with an
increase in the ratio of MnCl or β-aryl ether 2 to DypB from 1:1
to 10:1 (see Table 3). Single-electron oxidation of Mn or β-aryl
ether by compound I should generate a compound II intermedi-
ate, observed in other peroxidases at 417 nm. Observation at
17 nm revealed a first-order transient when DypB was mixed
with 1 equiv of hydrogen peroxide and β-aryl ether 2 (k = 1.18
ꢀ
1
ꢀ
1
ꢀ1
35
)
had been classified into four discrete subfamilies, AꢀD. As in
2
obs
our analysis, subfamilies AꢀC were classified as bacterial while
subfamily D was classified fungal, consistent with the Peroxibase
II
38
database. However, our analysis reclassifies some DyPs. For
35
2
example, the cyanobacterial AnaPX appears to be a C-type
enzyme. The similarity between A- and B-type DyPs on one hand
with C and D types on the other has been disputed because of
II
7
39
their low levels of sequence identity. However, the current TM-
align superposition establishes the homology of the DyPs
through their common structural fold and conserved active site
residues.
4
obs
ꢀ1
s
), but no change at 417 nm was detected upon mixing with
MnCl (see Table 3).
2
Gene deletion studies show that a ΔdypB deletion mutant of
R. jostii RHA1 has greatly impaired activity for lignin breakdown,
whereas a ΔdypA deletion mutant is unimpaired, compared with
wild-type RHA1. Expression of recombinant DypA and DypB
proteins and the nitrated lignin UVꢀvis assay have confirmed
that DypB shows activity in the presence of hydrogen peroxide,
As noted above, conversion of β-aryl ether 2 by DypB gave rise
to little or no vanillin product; therefore, we suspected that
vanillin could be oxidized by DypB, which was verified by pre-
steady-state kinetics. Incubation of 8 μM DypB with 1 mM
vanillin was found to give a 4-fold acceleration in the kobs for the
reaction of compound I at 397 nm and pH 5.5 and 7.5 (Figure S2
of the Supporting Information). In this case, we did not observe a
compound II-like intermediate during the reaction of compound I.
which is enhanced in the presence of MnCl , whereas DypA does
2
not. DypB also shows MichaelisꢀMenten kinetic behavior with
samples of Kraft lignin, shows effects upon lignocellulose break-
down in the presence of MnCl , and accepts a β-aryl ether lignin
2
’
DISCUSSION
model compound as a synthetic substrate. Taken together, these
data indicate that peroxidase DypB is involved in lignin break-
down by R. jostii RHA1. The turnover of only the disubstituted
β-aryl ether 2, and not the other lignin model compounds,
suggests that DypB is quite selective in its lignin degrading
activity. The cellular role of the DypA enzyme may be iron uptake
or utilization, because molecular genetic evidence has recently
been published for the deferrochelation of heme iron by the DyP
The microbial degradation of lignin has been studied mainly in
white-rot and brown-rot fungi, with the assumption that bacteria
are unable or slow to degrade lignin. However, there are several
reports of bacteria that have lignin degrading ability,
have previously identified a group of lignin-degrading bacteria,
including R. jostii RHA1, that have activity comparable to that of
some lignin-degrading fungi. There are reports of extracellular
peroxidases in St. viridosporus that are able to metabolize lignin
model compounds, but no protein sequence data are available for
these enzymes. The identification of bacterial lignin-degrading
genes would allow the high-level overexpression of the corres-
11,12
and we
15
40
homologues in E. coli. Indeed, of 111 unique A-type DyP genes
41
in the BioCyc database, 86% are predicted to have a TAT signal
sequence and 90% are encoded by genes that are clustered with
those predicted to encode iron uptake proteins.
1
4
5
104
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
II
The activation of DypB by Mn ions is reminiscent of the
with the encapsulin gene. The operonic coupling of the DyP and
encapsulin genes appears to occur only in burkholderials and
actinomycetales. It therefore seems probable that the targeting of
RHA1 DypB is somehow achieved with the aid of the adjacent
encapsulin gene, though the precise mechanism of this process
remains to be determined. Interestingly, members of the encap-
sulin family have been identified in the supernatants of myco-
II
fungal manganese peroxidase, whose rate of oxidation of Mn to
Mn is much faster than its rate of oxidation of lignin model
compounds. Activation of DypB by Mn was observed using
several different assays and substrates (see Table 2), and the
presence of Mn was found to be necessary to metabolize ligno-
cellulose, implying that Mn can act as an oxidative mediator for
DypB. Nevertheless, Mn is a relatively poor substrate for DypB-
III
6
,7
II
II
II
II
42
bacterial cell cultures.
4
5
catalyzed oxidation, with a k /K value 10 ꢀ10 -fold lower
In conclusion, we have identified a novel manganese-depen-
dent lignin peroxidase DypB, which plays a significant role in
lignin degradation by R. jostii RHA1. Considering that the in vitro
activity of recombinant DypB with lignocellulose is quite modest,
we suspect that there is a group of oxidative enzymes used by the
host for lignin metabolism, of which DypB is only one member.
Nevertheless, it is the first recombinant bacterial lignin peroxidase
to be kinetically characterized. The identification of DypB and
further bacterial lignin-degrading enzymes, and their availability in
recombinant form, will provide valuable new reagents for the
conversion of lignocellulose to biofuels and renewable chemicals.
cat
m
than that observed with fungal Mn peroxidases (DOI 10.1021/
bi200427h). Moreover, the observation of MichaelisꢀMenten
kinetic behavior of Kraft lignin suggests that DypB may be able to
bind lignin (or some part of the lignin structure) as part of its
a
II
catalytic reaction. To examine whether DypB oxidizes Mn in
preference to β-aryl ether 2, we have studied these reactions
using pre-steady-state kinetics. The kinetic data in Table 3
II
indicate a faster reaction of compound I in the presence of Mn
or β-aryl ether 2, but only modest increases in kobs were found;
only in the case of β-aryl ether 2 was a compound II species
observed at 417 nm. These data are not consistent with DypB
acting as a Mn peroxidase, in which case much higher rates would
’
ASSOCIATED CONTENT
II
be expected in the presence of Mn . There is some evidence of
turnover of β-aryl ether 2 via a one-electron oxidation mechan-
ism, consistent with the inhibition of turnover by diaphorase.
The increases in kobs in the presence of substrate are modest,
suggesting that DypB is not a highly active peroxidase, but it is
possible that β-aryl ether 2 may not be the optimal substrate. The
behavior of DypB is more reminiscent of that of the Pl. eryngii
versatile peroxidase, which has both manganese peroxidase and
S
b
Supporting Information. Sequences of oligonucleotide
primers (Table S1), alignment of Dyp protein sequences (Figure
S1), steady-state kinetic data for reaction of DypB with Kraft
lignin (Figure S2); HPLC and LCꢀMS data for reaction of DypB
with β-aryl ether 2 (Figures S3ꢀS7), and stopped-flow kinetic
data for reaction of DypB with hydrogen peroxide, MnCl , and
2
β-aryl ether 2 (Figures S8ꢀS15). This material is available free
37
lignin peroxidase interaction sites. In the following paper (DOI
0.1021/bi200427h), a putative manganese binding site is
of charge via the Internet at http://pubs.acs.org.
1
identified in the crystal structure of RHA1 DypB.
’
AUTHOR INFORMATION
We have identified using LCꢀMS the reaction products from
turnover of β-aryl ether 2 by DypB. The observation of a complex
mixture of higher-molecular weight reaction products, and the
inhibition of this reaction by diaphorase, are consistent with a
radical cleavage mechanism. In the presence of diaphorase, we
have identified vanillin and guaiacol derivative 5 as products,
indicating that C ꢀC bond cleavage is taking place, as found for
Corresponding Author
*
Department of Chemistry, University of Warwick, Coventry
CV4 7AL, U.K. Telephone: 44-(0)2476-573018. Fax: 44-(0)
476-524112. E-mail: T.D.Bugg@warwick.ac.uk.
2
Author Contributions
M.A. and J.N.R. contributed equally to the research.
R
β
8
the fungal Mn peroxidase. Because DypB also shows activity
toward guaiacol and vanillin as substrates (data not shown), the
mechanism is likely to proceed via one-electron oxidation of the
phenolic ring, followed by CꢀC bond cleavage, as shown in
Figure 7. We did not detect guaiacol itself in this experiment, but
we did observe a species corresponding to a guaiacol trimer, also
formed upon reaction of DypB with guaiacol as the substrate;
therefore, our interpretation is that guaiacol is formed from 2 but
is then rapidly oxidized by further reaction with DypB.
Funding Sources
This work was supported by research grants from the BBSRC
IBTI Biorefinery Club (Grant BB/H004270/1) and the Inno-
vative Materials Research Centre, University of Warwick, and a
Natural Sciences and Engineering Research Council (NSERC)
of Canada Discovery grant (to L.D.E.). J.N.R. was the recipient of
an NSERC studentship.
The cellular location of DypB must logically be extracellular,
because the sample used for our UVꢀvis assay is the extracellular
protein fraction after centrifugation. However, the protein se-
quence for R. jostii RHA1 DypB contains neither a Sec signal
sequence nor a TAT signal sequence. Interestingly, most DypA
homologues contain predicted TAT signal sequences, but most
DypB homologues do not. Immediately adjacent to the dypB
gene (ro02407) in the RHA1 genome is the ro02408 gene,
predicted to encode an encapsulin protein that has been shown
Notes
a
A reviewer has pointed out that the saturation kinetic behavior
observed with Kraft lignin, which is surprising, could perhaps be
caused by a physical phenomenon such as aggregation of the
substrate above a threshold level.
’ ACKNOWLEDGMENT
We thank Jie Liu for skilled technical assistance.
33
to form an icosahedral nanocompartment. Sutter et al. have
identified a C-terminal peptide tag that is found in proteins that
33
’ ABBREVIATIONS
associate with encapsulin, and this C-terminal tag is found in
41
0
RHA1 DypB. A survey of the BioCyc database revealed that
4% of the 150 B-type dyp genes are predicted to be co-operonic
PCB, polychlorinated biphenyl; ABTS, 2,2 -azinobis(3-ethyl-
benzthiazoline-6-sulfonic acid); LiP, lignin peroxidase; MnP,
1
5
105
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
manganese peroxidase; DyP, dye-decolorizing peroxidase; TAT,
twin-arginine transport; LBP, lysogeny broth peptone; MWL, milled
wood lignin; HPLC, high-performance liquid chromatography.
(18) Nakatsubo, F., Sato, K., and Higuchi, H. (1975) Synthesis of
guaiacyldlycerol-β-guaiacyl ether. Holzforschung 29, 165–168.
(19) Zimmermann, W., Paterson, A., and Broda, P. (1988) Conven-
tional and high-performance size-exclusion chromatography of graminac-
eous lignin carbohydrate complexes. Methods Enzymol. 161, 191–199.
(
20) Zhang, Y., and Skolnick, J. (2005) TM-align: A protein
’
REFERENCES
structure alignment algorithm based on the TM-score. Nucleic Acids
(
1) Sustainable biofuels: Prospects and challenges, Report by the
Royal Society, January 2008.
2) Wong, D. W. S. (2009) Structure and action mechanism of
lignolytic enzymes. Appl. Biochem. Biotechnol. 157, 174–209.
3) Tien, M., and Kirk, T. K. (1984) Lignin-degrading enzyme from
Phanerochaete chrysosporium: Purification, characterization, and catalytic
Res. 33, 2302–2309.
(21) Gille, C., and Frommel, C. (2001) STRAP: Editor for STRuc-
tural Alignments of Proteins. Bioinformatics 17, 377–378.
(22) Larkin, M. A., Blackshields, G., Brown, N. P., Chenna, R.,
McGettigan, P. A., McWilliam, H., Valentin, F., Wallace, I. M., Wilm, A.,
Lopez, R., Thompson, J. D., Gibson, T. J., and Higgins, D. G. (2007)
Clustal W and clustal X version 2.0. Bioinformatics 23, 2947–2948.
(23) Bendtsen, J. D., Nielsen, H., von Heijne, G., and Brunak, S.
(2004) Improved prediction of signal peptides: SignalP 3.0. J. Mol. Biol.
340, 783–795.
(
(
2 2
properties of a unique H O -requiring oxygenase. Proc. Natl. Acad. Sci.
U.S.A. 81, 2280–2284.
(
4) Hammel, K. E., Tien, M., Kalyanaraman, B., and Kirk, T. K.
(
1984) Mechanism of oxidative C -C cleavage of a lignin model dimer
by Phanerochaete chrysosporium ligninase: Stoichiometry and involve-
ment of free radicals. J. Biol. Chem. 260, 8348–8353.
R
β
(24) Felsenstein, J. (1989) Phylip: Phylogeny inference package
(version 3.2). Cladistics 5, 164–166.
(
5) Miki, K., Renganathan, V., and Gold, M. H. (1986) Mechanism
(25) van der Geize, R., Hessels, G. I., van Gerwen, R., van der
Meijden, P., and Dijkhuizen, L. (2001) Unmarked gene deletion
mutagenesis of kstD, encoding 3-ketosteroid Δ -dehydrogenase, in
of β-aryl ether dimeric lignin model compound oxidation by lignin
peroxidase of Phanerochaete chrysosporium. Biochemistry 25, 4790–4796.
1
(
6) Glenn, J. K., Akileswaran, L., and Gold, M. H. (1986) Mn(II)
Rhodococcus erythropolis SQ1 using sacB as counter-selectable marker.
FEMS Microbiol. Lett. 205, 197–202.
oxidation is the principal function of the extracellular Mn peroxidase
from Phanerochaete chrysosporium. Arch. Biochem. Biophys. 251, 688–696.
(26) Bertani, G. (2004) Lysogeny at mid-twentieth century: P1, P2,
and other experimental, systems. J. Bacteriol. 186, 595–600.
(7) Wariishi, H., Dunford, H. B., MacDonald, I. D., and Gold, M. H.
(
1989) Manganese peroxidase from the lignin-degrading basidomycete
(27) Falk, J. E. (1964) Porphyrins and metalloporphyrins; their general,
physical and coordination chemistry, and laboratory methods, Elsevier
Publishing Co., Amsterdam.
Phanerochaete chrysosporium: Transient state kinetics and reaction
mechanism. J. Biol. Chem. 264, 3335–3340.
(
8) Tuor, U., Wariishi, H., Schoemaker, H. E., and Gold, M. H.
(28) Sugano, Y., Muramatsu, R., Ichiyanagi, A., Sato, T., and Shoda,
M. (2007) DyP, a unique dye-decolorizing peroxidase, represents a
novel heme peroxidase family: Asp-171 replaces the distal histidine of
classical peroxidases. J. Biol. Chem. 282, 36652–36658.
(
1992) Oxidation of phenolic arylglycerol β-aryl ether lignin model
compounds by manganese peroxidase from Phanerochaete chrysospor-
ium: Oxidative cleavage of an R-carbonyl model compound. Biochemistry
1, 4986–4995.
3
(29) Zubieta, C., Krishna, S. S., Kapoor, M., Kozbial, P., McMullan,
D., Axelrod, H. L., Miller, M. D., Abdubek, P., Ambing, E., Astakhova, T.,
Carlton, D., Chiu, H. J., Clayton, T., Deller, M. C., Duan, L., Elsliger,
M. A., Feuerhelm, J., Grzechnik, S. K., Hale, J., Hampton, E., Han, G. W.,
Jaroszewski, L., Jin, K. K., Klock, H. E., Knuth, M. W., Kumar, A.,
Marciano, D., Morse, A. T., Nigoghossian, E., Okach, L., Oommachen,
S., Reyes, R., Rife, C. L., Schimmel, P., van den Bedem, H., Weekes, D.,
White, A., Xu, Q., Hodgson, K. O., Wooley, J., Deacon, A. M., Godzik, A.,
Lesley, S. A., and Wilson, I. A. (2007) Crystal structures of two novel
dye-decolorizing peroxidases reveal a β-barrel fold with a conserved
heme-binding motif. Proteins 69, 223–233.
(30) Zubieta, C., Joseph, R., Krishna, S. S., McMullan, D., Kapoor,
M., Axelrod, H. L., Miller, M. D., Abdubek, P., Acosta, C., Astakhova, T.,
Carlton, D., Chiu, H. J., Clayton, T., Deller, M. C., Duan, L., Elias, Y.,
Elsliger, M. A., Feuerhelm, J., Grzechnik, S. K., Hale, J., Han, G. W.,
Jaroszewski, L., Jin, K. K., Klock, H. E., Knuth, M. W., Kozbial, P., Kumar,
A., Marciano, D., Morse, A. T., Murphy, K. D., Nigoghossian, E., Okach,
L., Oommachen, S., Reyes, R., Rife, C. L., Schimmel, P., Trout, C. V., van
den Bedem, H., Weekes, D., White, A., Xu, Q., Hodgson, K. O., Wooley,
J., Deacon, A. M., Godzik, A., Lesley, S. A., and Wilson, I. A. (2007)
Identification and structural characterization of heme binding in a novel
dye-decolorizing peroxidase, TyrA. Proteins 69, 234–243.
(
9) Kawai, S., Umezawa, T., and Higuchi, T. (1988) Degradation
mechanisms of phenolic β-1 lignin substructure model compounds by
laccase of Coriolus versicolor. Arch. Biochem. Biophys. 262, 99–110.
(
Borneman, S. (1997) Reactivities of various mediators and laccases with
Kraft pulp and lignin model compounds. Appl. Environ. Microbiol. 63,
10) Bourbonnais, R., Paice, M. G., Freiermuth, B., Bodie, E., and
4
627–4632.
11) Vicuna, R. (1988) Bacterial degradation of lignin. Enzyme
Microb. Technol. 10, 646–655.
12) Zimmermann, W. (1990) Degradation of lignin by bacteria.
J. Biotechnol. 13, 119–130.
13) Crawford, D., Pometto, A., III, and Crawford, R. (1983) Lignin
(
(
(
degradation by Streptomyces viridosporus: Isolation and charaterization of
a new polymeric lignin degradation intermediate. Appl. Environ. Micro-
biol. 45, 898–904.
(
14) Ramachandra, M., Crawford, D., and Hertel, G. (1988) Char-
acterization of an extracellular lignin peroxidase of the lignocellulolytic
actinomycete Streptomyces viridosporus. Appl. Environ. Microbiol. 54,
057–3063.
(
Bending, G. D., and Bugg, T. D. H. (2010) Development of novel assays
for lignin degradation: Comparative analysis of bacterial and fungal
lignin degraders. Mol. BioSyst. 6, 815–821.
3
15) Ahmad, M., Taylor, C. R., Pink, D., Burton, K., Eastwood, D.,
(31) Cao, J., Woodhall, M. R., Alvarez, J., Cartron, M. L., and Andrews,
S. C. (2007) EfeUOB (YcdNOB) is a tripartite, acid-induced and CpxAR-
2þ
(
16) Seto, M., Kimbara, K., Shimura, M., Hatta, T., Fukuda, M., and
regulated, low-pH Fe transporter that is cryptic in Escherichia coli K-12
but functional in E-coli O157:H7. Mol. Microbiol. 65, 857–875.
(32) Stearman, R., Yuan, D. S., Yamaguchi-Iwai, Y., Klausner, R. D.,
and Dancis, A. (1996) A permease-oxidase complex involved in high-
affinity iron uptake in yeast. Science 271, 1552–1557.
Yano, K. (1995) A novel transformation of polychlorinated biphenyls by
Rhodococcus sp. strain RHA1. Appl. Environ. Microbiol. 61, 3353–3358.
(
Myhre, M., Femandes, C., Miyazawa, D., Wong, W., Lillquist, A. L.,
Wang, D., Dosanjh, M., Hara, H., Petrescu, A., Morin, R. D., Yang, G.,
Stott, J. M., Schein, J. E., Shin, H., Smailus, D., Siddiqui, A. S., Marra,
M. A., Jones, S. J. M., Holt, R., Brinkman, F. S. L., Miyauchi, K., Fukuda,
M., Davies, J. E., Mohn, W. W., and Eltis, L. D. (2006) The complete
genome of Rhodococcus sp. RHA1 provides insights into a catabolic
powerhouse. Proc. Natl. Acad. Sci. U.S.A. 103, 15582–15587.
17) McLeod, M. P., Warren, R. L., Hsiao, W. W. L., Araki, N.,
(33) Sutter, M., Boehringer, D., Gutmann, S., G €u nther, S., Prangishvili,
D., Loessner, M. J., Stetter, K. O., Weber-Ban, E., and Ban, N. (2008)
Structural basis of enzyme encapsulation into a bacterial nanocompartment.
Nat. Struct. Mol. Biol. 15, 939–947.
(34) de Geus, D. C., Thomassen, E. A., Hagedoorn, P. L., Pannu,
N. S., van Duijn, E., and Abrahams, J. P. (2009) Crystal structure of
5
106
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107

Biochemistry
ARTICLE
chlorite dismutase, a detoxifying enzyme producing molecular oxygen.
J. Mol. Biol. 387, 192–206.
(35) Ogola, H. J. O., Kamiike, T., Hashimoto, N., Ashida, H.,
Ishikawa, T., Shibata, H., and Sawa, Y. (2009) Molecular characteriza-
tion of a novel peroxidase from the cyanobacterium Anabaena sp. strain
PCC 7120. Appl. Environ. Microbiol. 75, 7509–7518.
(
36) Beyer, R. E., Segura Aguilar, J., Di Bernardo, S., Cavazzoni, M.,
Fato, R., Fiorentini, D., Galli, M. C., Setti, M., Jandi, L., and Lenaz, G.
1996) The role of DT-diaphorase in the maintenance of the reduced
(
antioxidant form of coenzyme Q in membrane systems. Proc. Natl. Acad.
Sci. U.S.A. 93, 2528–2532.
(37) Camarero, S., Sarkar, S., Ruiz-Duenas, F. J., Martinez, M. J., and
Martinez, A. T. (1999) Description of a versatile peroxidase involved in
the natural degradation of lignin that has both manganese peroxidase
and lignin peroxidase substrate interaction sites. J. Biol. Chem.
274, 10324–10330.
(
38) Koua, D., Cerutti, L., Falquet, L., Sigrist, C. J., Theiler, G., Hulo,
N., and Dunand, C. (2009) PeroxiBase: A database with new tools for
peroxidase family classification. Nucleic Acids Res. 37, D261–D266.
(39) Hofrichter, M., Ullrich, R., Pecyna, M. J., Liers, C., and Lundell,
T. (2010) New and classic families of secreted fungal heme peroxidases.
Appl. Microbiol. Biotechnol. 87, 871–897.
(40) Letoffe, S., Heuck, G., Delepelaire, P., Lange, N., and
Wandersman, C. (2009) Bacteria capture iron from heme by keeping
tetrapyrrol skeleton intact. Proc. Natl. Acad. Sci. U.S.A. 106, 11719–
11724.
(
41) Karp, P. D., Ouzounis, C. A., Moore-Kochlacs, C., Goldovsky,
L., Kaipa, P., Ahren, D., Tsoka, S., Darzentas, N., Kunin, V., and Lopez-
Bigas, N. (2005) Expansion of the BioCyc collection of pathway/
genome databases to 160 genomes. Nucleic Acids Res. 33, 6083–6089.
(42) Rosenkrands, I., Rasmussen, P. B., Carnio, M., Jacobsen, S.,
Theisen, M., and Andersen, P. (1998) Identification and characteriza-
tion of a 29-kilodalton protein from Mycobacterium tuberculosis culture
filtrate recognized by mouse memory effector cells. Infect. Immun.
66, 2728–2735.
5
107
dx.doi.org/10.1021/bi101892z |Biochemistry 2011, 50, 5096–5107