Roles of small laccases from streptomyces in lignin degradation

Article abstract of DOI:10.1021/bi500285t

Laccases (EC 1.10.3.2) are multicopper oxidases that can oxidize a range of substrates, including phenols, aromatic amines, and nonphenolic substrates. To investigate the involvement of the small Streptomyces laccases in lignin degradation, we generated acid-precipitable polymeric lignin obtained in the presence of wild-type Streptomyces coelicolor A3(2) (SCWT) and its laccase-less mutant (SCΔLAC) in the presence of Miscanthus x giganteus lignocellulose. The results showed that strain SCΔLAC was inefficient in degrading lignin compared to strain SCWT, thereby supporting the importance of laccase for lignin degradation by S. coelicolor A3(2). We also studied the lignin degradation activity of laccases from S. coelicolor A3(2), Streptomyces lividans TK24, Streptomyces viridosporus T7A, and Amycolatopsis sp. 75iv2 using both lignin model compounds and ethanosolv lignin. All four laccases degraded a phenolic model compound (LM-OH) but were able to oxidize a nonphenolic model compound only in the presence of redox mediators. Their activities are highest at pH 8.0 with a low k_rel/K_app for LM-OH, suggesting that the enzymes' natural substrates must be different in shape or chemical nature. Crystal structures of the laccases from S. viridosporus T7A (SVLAC) and Amycolatopsis sp. 75iv2 were determined both with and without bound substrate. This is the first report of a crystal structure for any laccase bound to a nonphenolic β-O-4 lignin model compound. An additional zinc metal binding site in SVLAC was also identified. The ability to oxidize and/or rearrange ethanosolv lignin provides further evidence of the utility of laccase activity for lignin degradation and/or modification.

Full text of DOI:10.1021/bi500285t

Subscriber access provided by CLEMSON UNIV
Article
The roles of small laccases from Streptomyces in lignin degradation
Sudipta Majumdar, Tiit Lukk, Jose O. Solbiati, Stefan Bauer, Satish K Nair, John E. Cronan, and John A. Gerlt
Biochemistry, Just Accepted Manuscript • DOI: 10.1021/bi500285t • Publication Date (Web): 28 May 2014
Downloaded from http://pubs.acs.org on June 7, 2014
Just Accepted
“Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted
online prior to technical editing, formatting for publication and author proofing. The American Chemical
Society provides “Just Accepted” as a free service to the research community to expedite the
dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts
appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been
fully peer reviewed, but should not be considered the official version of record. They are accessible to all
readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered
to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published
in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just
Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor
changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers
and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors
or consequences arising from the use of information contained in these “Just Accepted” manuscripts.
Biochemistry is published by the American Chemical Society. 1155 Sixteenth Street
N.W., Washington, DC 20036
Published by American Chemical Society. Copyright © American Chemical Society.
However, no copyright claim is made to original U.S. Government works, or works
produced by employees of any Commonwealth realm Crown government in the course
of their duties.

Page 1 of 45
Biochemistry
1
2
3
4
5
6
7
8
The roles of small laccases from Streptomyces in lignin degradation
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Sudipta Majumdar‡,#,*, Tiit Lukk‡,¥, Jose O. Solbiati‡, Stefan Bauer±, Satish K. Nair‡, John E.
Cronan‡, and John A. Gerlt‡
‡Institute for Genomic Biology, University of Illinois at UrbanaꢀChampaign, Illinois 61801 and
±Energy Biosciences Institute, University of California, Berkeley, California 94720
#Current address: Columbia University, New York City, New York 10027, ¥Cornell University,
Ithaca, New York 14853
*To whom correspondence should be addressed: Sudipta Majumdar, Department of Chemical
Engineering, Columbia University; telephone, (212) 854ꢀ1444; email, sm3577@columbia.edu
Funding information:
This research was supported by Energy Biosciences Institute, University of Illinois at Urbanaꢀ
Champaign, Illinois 61801. Use of the Advanced Photon Source was supported by the U.S.
Department of Energy, Office of Sciences, under Contact No. DEꢀAC02ꢀ06CH11357; LSꢀCAT
Sector 21 is supported by the Michigan Economic Development Corporation and the Michigan
Technology TriꢀCorridor (Grant 085P1000817).
The authors declare no competing financial interest.
ACS Paragon Plus Environment
1

Biochemistry
Page 2 of 45
1
2
3
4
5
6
7
ABBREVIATIONS
8
9
LiP, lignin peroxidase; MnP, manganese peroxidase; SCLAC, Streptomyces coelicolor laccase;
SLLAC, Streptomyces lividans laccase; SVLAC, Streptomyces viridosporus laccase; AMLAC,
Amycolatopsis laccase; TAT, twinꢀarginine translocation; LMꢀOMe, 1ꢀ(3,4ꢀdimethoxyphenyl)ꢀ2ꢀ
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
(2ꢀmethoxyphenoxy)ꢀ1,3ꢀdihydroxypropane;
methoxyphenoxy)ꢀ1,3ꢀdihydroxypropane;
LMꢀOH,
1ꢀ(3ꢀmethoxyꢀ4ꢀhydroxy)ꢀ2ꢀ(2ꢀ
ABTS,
2,2’ꢀazinoꢀbis(3ꢀethylbenzothiazolineꢀ6ꢀ
sulfonate); DMP, 2,6ꢀdimethoxy phenol; AV, acetovanillone; HOBt, hydroxybenzotiazole;
CH3CN, acetonitrile; DMSO, dimethyl sulfoxide; MgCl2, magnesium chloride; NaCl, sodium
chloride; HCl, hydrochloric acid; KCl, potassium chloride; Na2HPO4, disodium hydrogen
phosphate; KH2PO4, potassium dihydrogen phosphate; CuSO4, copper sulfate; DMF, dimethyl
formamide; PBS, phosphate buffered saline; dNTP, deoxyribinucleotide; DNA,
deoxyribonucleic acid; FAD, flavin adenine dinucleotide; LB, luriaꢀbertani; IPTG, isopropyl βꢀ
Dꢀ1 thipgalactopyranoside; 3ꢀHAA, 3ꢀhydroxyanthranilic acid; OD, optical density; HPLC,
highꢀperformance liquid chromatography; LCMSꢀITꢀTOF, liquid chromatography mass
spectrometry ionꢀtrap timeꢀofꢀflight; PCR, polymerase chain reaction; UVꢀVis, ultravioletꢀ
visible; krel, relative rate constant; Kapp, apparent KM.
ACS Paragon Plus Environment
2

Page 3 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
ABSTRACT: Laccases (EC 1.10.3.2) are multicopper oxidases able to oxidize a range of
substrates including phenols, aromatic amines and nonꢀphenolic substrates. In order to
investigate the involvement of the small Streptomyces laccases in lignin degradation, we
generated acidꢀprecipitable polymeric lignin (APPL) obtained in the presence of wild type
Streptomyces coelicolor A3(2) (SCWT) and its laccaseꢀless mutant (SCꢁLAC) in presence of
Miscanthus x giganteus lignocellulose. The results showed that strain SCꢁLAC was inefficient
in degrading lignin compared to strain SCWT, thereby supporting the importance of laccase for
lignin degradation by Streptomyces coelicolor A3(2). We also studied the lignin degradation
activity of laccases from Streptomyces coelicolor A3(2), Streptomyces lividans TK24,
Streptomyces viridosporus T7A and Amycolatopsis sp. 75iv2 using both lignin model
compounds and ethanosolv lignin. All four laccases degraded a phenolic model compound (LMꢀ
OH), but only able to oxidize a nonꢀphenolic model compound (LMꢀOMe) in the presence of
redox mediators. Their activities are highest at pH 8.0 with a low krel/Kapp for LMꢀOH,
suggesting that the enzymes’ natural substrates must be different in shape or chemical nature.
Crystal structures of the laccases from Streptomyces viridosporus T7A (SVLAC) and
Amycolatopsis sp. 75iv2 (AMLAC) were determined both with and without bound substrate.
This is the first report of a crystal structure for any laccase bound to a nonꢀphenolic βꢀOꢀ4 lignin
model compound. An additional zinc metal binding site in SVLAC was also identified. The
ability to oxidize/rearrange ethanosolv lignin further provides evidence for the utility of laccase
activity for lignin degradation/modification.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
ACS Paragon Plus Environment
3

Biochemistry
Page 4 of 45
1
2
3
4
5
6
7
8
9
Lignocellulosic biomass, composed of lignin, cellulose and hemicellulose, has been
targeted as a potential source of renewable bioenergy. Lignin accounts for 20% of lignocellulosic
material and is an integral part of all higher plants and the second most abundant organic
polymer in nature after cellulose. Lignin serves as a key structural component and an outer
protective shield against biochemical hydrolysis of the energy rich and more easily
metabolizable cellulose and hemicellulose (1, 2). Therefore, lignin removal is required for
efficient utilization of lignocellulosic biomass for the production of biofuels and other celluloseꢀ
based chemicals. Lignin has a complex, heterogeneous, polymeric structure derived from
oxidativeꢀcoupling of three phenylpropanoid monomers, pꢀcoumaryl, coniferyl and sinapyl
alcohols (Figure S1) (3). The complexity of lignin structure plus its covalent connectivity to
hemicellulose provides not only mechanical and structural stability but also resistance to
chemical and biological degradation. Most of the available chemical pretreatment methods for
lignin degradation have adverse effects on the later stages of energy conversion from cellulosic
biomass and generate toxic waste products (4ꢀ6). For more ecologically favorable applications,
methods using ligninꢀdegrading microbes and/or their enzymes may provide a more efficient and
environmentally sound approach for accessing renewable lignocellulosic biomass.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
A variety of microorganisms, including certain fungi and bacteria, can degrade lignin,
thus enabling access to plant carbohydrates as a rich energy source (7ꢀ10). Among these
microorganisms whiteꢀrot fungi have attracted widespread attention because of their powerful
ligninꢀdegrading enzymatic systems (11, 12). However, utilization of these enzymes is
problematic because of their poor stability in industrial processes, as well as the difficulty in
developing efficient heterologous systems for enzyme expression and purification.
Consequently, much attention has been paid to lignin degradation by bacterial species (13ꢀ15).
ACS Paragon Plus Environment
4

Page 5 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
The most extensively studied are actinomycetes, particularly Streptomyces species because of
their reported efficiency in lignin degradation, established genetic and molecular engineering
tools and the availability of complete genome sequences (16, 17). In spite of this, very little is
known about either the involvement of specific enzymes or the molecular mechanism of lignin
degradation. Extensive work has been performed with Streptomyces viridosporus T7A in which
an extracellular lignin peroxidase (LiP) was reported to possess significant lignin degradation
activity (18ꢀ20). Despite some reports in the literature, bona fide extracellular lignin degrading
peroxidases have yet to be identified in Streptomyces viridosporus T7A (9, 18, 19, 21, 22).
Recently an extracellular DyP peroxidase from Amycolatopsis sp. 75iv2 was characterized and
reported as lignin degrading enzyme (14, 23). Efforts to find enzymes responsible for lignin
degradation by Streptomyces had resulted in our discovery of small laccase, a copper dependent
phenol oxidase (24).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Laccases (pꢀdiphenol: dioxygen oxidoreductase) are copperꢀcontaining enzymes that can
oxidize a range of aromatic and nonꢀaromatic compounds containing hydroxyl and amine groups
in presence of atmospheric oxygen (25, 26). These enzymes are produced not only by eukaryotes
such as fungi and plants but also by prokaryotes, including a wide range of Gramꢀpositive and
Gramꢀnegative bacteria (25, 27, 28). Although laccases are believed to be involved in lignin
synthesis in plants and in cell pigment formation, as well as metal oxidation in fungi and
bacteria, their physiological roles are unclear. The most detailed biochemical characterization of
laccases was done with the enzymes from the basidomycetes, Pycnoporus cinnabarinus and
Trametes versicolor, wherein laccases were shown to play important roles in lignin degradation
(29, 30). Recently, laccases from actinomycetes were reported to have in vitro activity against a
wide range of substrates. These enzymes were designated as “small laccases” because of their
ACS Paragon Plus Environment
5

Biochemistry
Page 6 of 45
1
2
3
4
5
6
7
8
9
sequence similarity but smaller size compared to fungal laccases (24). Small laccases have been
extensively exploited in the pulp and paper industries for dyeꢀdecolorization and bioꢀbleaching
due to their high oxidizing power, pHꢀversatility and thermal stability (24, 31). Despite the
advantages bacterial enzymes may offer over fungal enzymes, no detailed biochemical studies
have been done using either native lignocellulosic biomass or lignin model compounds as
substrates. Improved knowledge about bacterial laccases and their role in lignin degradation will
have significant impact on a wide array of biotechnologies focused on lignin degradation.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
The purpose of the present work is to investigate the importance of the small laccases
from Streptomyces in both in vivo and in vitro lignin degradation activity assays using
lignocellulosic biomass from Miscanthus x giganteus, ethanosolv lignin and βꢀOꢀ4 lignin model
compounds (3, 32). A Streptomyces coelicolor A3(2) laccase deficient mutant (SCꢁLAC) was
constructed to assess its in vivo lignin degradation activity. Using ethanosolv lignin and βꢀOꢀ4
lignin model compounds we characterized four different small laccases from Streptomyces
coelicolor A3(2) (SCLAC), Streptomyces lividans TK24 (SLLAC), Streptomyces viridosporus
T7A (SVLAC) and Amycolatopsis sp. 75iv2 (AMLAC) (previously known as Streptomyces
setonii and Streptomyces griseus 75iv2) for their in vitro lignin degradation/oxidation activity.
For better solubility, ethanosolv lignocellulose was used as the substrate for HPLC assays (3).
Two different βꢀOꢀ4 lignin model compounds, with and without a phenolic group (Figure S2),
were also used as substrates for spectrophotometric assays. Xꢀray crystal structures of small
laccases, with and without substrate bound, from Streptomyces viridosporus T7A and
Amycolatopsis sp. 75iv2 are also reported.
ACS Paragon Plus Environment
6

Page 7 of 45
Biochemistry
1
2
3
4
MATERIALS AND METHODS
5
6
7
8
9
All reagents and chemicals used as buffers and substrates were purchased of the highest
grade commercially available from SigmaꢀAldrich, Alfa Aesar, Acros, MP Biomedicals, or
Fisher. The lignin model compounds, 1ꢀ(3, 4ꢀdimethoxyphenyl)ꢀ2ꢀ(2ꢀmethoxyphenoxy)ꢀ1, 3ꢀ
dihydroxypropane [LMꢀOMe, Figure S2] and 1ꢀ(3ꢀmethoxyꢀ4ꢀhydroxy)ꢀ2ꢀ(2ꢀmethoxyphenoxy)ꢀ
1, 3ꢀdihydroxypropane [LMꢀOH, Figure S2] were purchased from AstaTech, PA, USA and used
as received. The ethanosolv lignin was prepared as described previously (32). The Trametes
versicolor fungal laccase was purchased from SigmaꢀAldrich. Enzymes for gene cloning were
purchased from New England Biolabs. The buffers were made by mixing 0.1 M acetic acid, 0.1
M sodium acetate, 0.1 M potassium phosphate monobasic, 0.1 M potassium phosphate dibasic,
0.1 M sodium carbonate and 0.1 M sodium bicarbonate to the desired pH. UVꢀVis
spectrophotometric data were collected using a Cary 300 Bio Spectrophotometer. HPLC data
were collected using Beckman Coulter System Gold Spectrophotometer. Mass Spectrometry data
were collected using Shimadzu LCMSꢀITꢀTOF. Protein purifications were done using
AKTAxpress (GE) protein purification system. Xꢀray diffraction data were collected at Argonne
National Laboratory, IL, USA.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Microorganisms. The strains of Streptomyces coelicolor A3(2) and Streptomyces lividans
TK24 were kindly supplied by Prof. William W. Metcalf (University of Illinois at Urbanaꢀ
Champaign) and the strains of Streptomyces viridosporus T7A (ATCC 39115) and
Amycolatopsis sp. 75iv2 (ATCC 39116) were obtained from American Type Culture Collection
(Rockville, MD). All strains were grown in Difco ISP Medium 1 and genomic DNAs were
isolated using Wizard Genomic DNA Purification Kit (Promega) following manufacture’s
protocol.
ACS Paragon Plus Environment
7

Biochemistry
Page 8 of 45
1
2
3
4
5
6
7
8
9
Double crossꢀover inꢀframe deletion of SCO6712 by PCR targeting. The deletion of the
gene encoding the Streptomyces coelicolor A3(2) small laccase (SCO6712) was performed
following the procedures described in the REDIRECT manual (33). The gene was replaced with
an apramycin resistance cassette, apr, using the PCRꢀtargeted double crossꢀover inꢀframe
deletion technique (34). The extended resistance cassette including apr resistance marker,
aac(3)IV, was amplified from pIJ773 by PCR using SCO6712ꢀKOꢀF and SCO6712ꢀKOꢀR
primers (Table S1). Cosmid St4C6 containing the SCO6712 gene was introduced into E. coli
BW25113/pIJ790 (λ RED recombination plasmid) by electroporation and the resulting
transformants were then electroporated with the extended resistance cassette to target the
SCO6712 gene. The resulting mutant cosmid, St4C6ꢁSCO6712::apr, was transformed into E.
coli ET12567/pUZ8002 (nonmethylating strain) and then transferred into wildꢀtype Streptomyces
coelicolor A3(2) (SCWT), yielding the desired SCꢁLAC mutant. The mutant was selected by
virtue of its apramycin resistance and kanamycin sensitivity. The disruption of SCO6712 by the
apr cassette was confirmed by PCR on the isolated genomic DNAs of both the SCꢁLAC and
SCWT using the primers KOꢀCKꢀF and KOꢀCKꢀR (Table S1). The mutation also was confirmed
by DNA sequencing of PCR products.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Lignocellulose preparation. Miscanthus x giganteus from an experimental field at
University of Illinois at UrbanaꢀChampaign was used as the source of lignocellulose. The ground
lignocellulose was prepared by passage of airꢀdried lignocellulose through an 80 micron sieve
(SR 300 rotor beater mill, Retsch, Germany). Dry lignocellulose (3.5 g) was placed in a 1 L
Erlenmeyer flask with several glass beads and autoclaved for 1 hr uncovered and then additional
30 min covered with aluminum foil.
ACS Paragon Plus Environment
8

Page 9 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
Inoculum preparation. Spores of the SCWT or the SCꢁLAC (100 ꢂL stock) were added
to 200 mL of autoclaved Difco ISP Medium 1 and incubated aerobically at 30 °C for 3 days by
shaking at 220 rpm with glass beads, at which time the cells had entered into late logarithmic
growth phase (35). The cells were collected by centrifugation, washed and resuspended in 30 mL
of yeast extractꢀmineral salt (YEMS) medium (35).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Acidꢀprecipitable polymeric lignin (APPL) production from lignocellulose. Autoclaved
YEMS medium (1 L) was added to the sterile lignocellulose described previously and 10 mL of
diluted logꢀphase cell suspension of Streptomyces was added; the mixture was incubated in a 2 L
flask at 30 °C for one week with shaking at 220 rpm. Control reactions with YEMS medium
only, YEMS medium with autoclaved lignocellulose, YEMS medium with the SCWT lacking
lignocellulose and YEMS medium with the SCꢁLAC strain lacking lignocellulose were
performed under similar conditions. All reactions were performed in triplicate. After seven days,
the residual lignocellulose and bacterial cells were removed by centrifugation. The supernatants
°
then were acidified at 4 C to pH 5.0 with concentrated HCl. The precipitated APPL was
collected by centrifugation, airꢀdried, weighed and analyzed chemically.
Chemical characterization of APPL. The APPL (from the SCWT or SCꢀLAC) was
incubated at room temperature with 0.5 mL of 72% (w/w) sulfuric acid in a modified Hungate
vial capped with a rubber stopper; the mixture was vortexed every 15 min. After 1 hr of
incubation 14 mL of deionized water was added and the mixture was autoclaved for 60 min. A
sugar recovery standard containing the same sulfuric acid concentration was prepared in a
similar way and coꢀautoclaved with the samples. The mixture was kept in the refrigerator
overnight and 2 mL of the clear supernatant was filtered (0.45 µm, PES) and used for highꢀ
performance liquid chromatography (HPLC) analysis.
ACS Paragon Plus Environment
9

Biochemistry
Page 10 of 45
1
2
3
4
5
6
7
8
9
The precipitated solids were resuspended by vortexing and the suspension was filtered
through a glass micro filter. Both the vial and filter were extensively rinsed with deionized water
and dried at 105 °C overnight; the weight was determined after cooling in a desiccator for 30
min. The filter and solids were then incubated at 575 °C (ramp: 105 °C for 10 min, 200 °C for 10
min, 300 °C for 30 min, 575 °C for 3 h, cooling to 105 °C); the weight was determined after
cooling in desiccator for 30 min.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Calculation [Klason lignin %] = [m(filter + dried solid) – m(filter +ash)]/m(dried initial
APPL) * 100 %
For ash determination, the APPL (SCWT or SCꢀLAC) was incubated on a preꢀweighed
and conditioned aluminum pan at 575 °C (ramp: 105 °C for 10 min, 200 °C for 10 min, 300 °C
for 30 min, 575 °C for 3 h, cooling to 105 °C); the weight was determined after cooling in
desiccator for 30 min.
Calculation [ash %] = [m(pan + ash) – m(pan)]/m(dried initial APPL) * 100 %
For HPLC, samples were analyzed at 50 °C on an HPXꢀ87H (300 x 7.8 mm, Bio Rad)
column on an Agilent 1200 series liquid chromatography instrument equipped with a refractive
index detector. Elution was performed with 0.005 M sulfuric acid at a flow rate of 0.6 mL/min.
Cloning, expression and purification of small laccases from Streptomyces coelicolor
(SCLAC), Streptomyces lividans (SLLAC), Streptomyces viridosporus (SVLAC) and
Amycolatopsis (AMLAC). The genes encoding the small laccase from Streptomyces coelicolor
A3(2) (GI:21225006), Streptomyces lividans TK24 (GI:256783840), Streptomyces viridosporus
T7A (GI:2518361192) and Amycolatopsis sp. 75iv2 (GI:2513525025) were PCRꢀamplified from
their corresponding genomic DNA without TAT secretion tag, with the primers (Integrated DNA
ACS Paragon Plus Environment
10

Page 11 of 45
Biochemistry
1
2
technologies) listed in Table S1 and all of which had a 5׳ꢀNdeI restriction site and 3׳ꢀBamHI
restriction site, respectively. The PCR mixture (50 ꢂL) contained 5 ꢂL of 5 ng/ꢂL template
genomic DNA, 10 ꢂL of Expand High FidelityPLUS reaction buffer (5X) with 7.5 mM MgCl2, 2
ꢂL 10 mM dNTP mixture, 2.5 ꢂL DMSO, 2 ꢂL of each 20 ꢂM forward and reverse primers, 1
ꢂL Expand High FidelityPLUS (5 U/ꢂL) enzyme blend (Roche) and 25.5 ꢂL of ddH2O. The PCR
were performed in a Veriti 96 Well Thermal Cycler (Applied Biosystems) with the following
parameters: 94 °C for 4 min followed by 35 cycles of 94 °C for 30 sec, 70 °C for 45 sec, 72 °C for
1.15 min and the final extension time was 7 min at 72 °C. The PCR products were purified by gel
extraction (Qiagen) following the manufacturer’s protocol. The amplified DNA products were
digested with NdeI and BamHI restriction enzymes (New England Biolabs) and ligated into
pET15b expression vector (Novagen) cut with the same enzymes. The Nꢀterminal His tagged
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
°
proteins were expressed in E. coli BL21 (DE3) cells at 30 C. A bacterial culture for a typical
°
preparation utilized 4 x 1 L of LB medium shaken at 37 C until the OD600 reached 0.6 after
°
which it was induced with 1 mM IPTG. The culture was then shaken at 30 C for another 20 h
and harvested by centrifugation. The cell pellets were resuspended in 50 mL of buffer containing
5 mM imidazole, 500 mM NaCl and 20 mM TrisꢀHCl (pH 7.9). The suspension was sonicated to
lyse the cells and the lysate was cleared by centrifugation. The supernatant was applied to a 5 mL
HisTrapTM HP column (GE Healthcare) and eluted with a linear gradient (100 mL) of 0 to 1 M
imidazole buffered with 500 mM NaCl and 20 mM TrisꢀHCl (pH 7.9). Fractions containing the
small laccase was collected and dialyzed against 1X PBS (Phosphate Buffered Saline) buffer (pH
7.4) containing 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4 and 1.76 mM KH2PO4. After
cleavage of the Nꢀterminal Hisꢀtag by thrombin (GE Healthcare) at room temperature, the
proteins were dialyzed against 20 mM TrisꢀHCl (pH 7.9) before being applied to a 5 mL
ACS Paragon Plus Environment
11

Biochemistry
Page 12 of 45
1
2
HiTrapTM Q HP column (GE Healthcare). The proteins were eluted with a linear gradient (100
mL) of 0 to 1 M NaCl buffered with 20 mM TrisꢀHCl (pH 7.9). Fractions containing pure (>
95%) proteins were collected and dialyzed twice against 20 mM TrisꢀHCl (pH 7.9) containing
CuSO4 at four fold greater molar ratio to given 4 copper atoms per active site and then stored at ꢀ
80 °C.
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Determination of laccase activity at different pH. The small laccase activities at different
°
pH (3ꢀ10) were determined at 25 C using 2,2׳ꢀazinoꢀbis(3ꢀethylbenzothiazolineꢀ6ꢀsulfonate)
[ABTS] or 2,6ꢀdimethoxy phenol [DMP] as the substrate. The oxidation of substrates was
detected by measuring the absorbance at 420 nm for ABTS (ε420 = 36000 Mꢀ1 cmꢀ1) at pH 3ꢀ5
and at 468 nm (ε468 = 14800 Mꢀ1 cmꢀ1) for DMP at pH 6ꢀ10. The reaction mixture (200 ꢂL)
contained 10 mM ABTS (dissolved in water) and 5 mM DMP (dissolved in DMF) and the small
laccase (2.0 ꢂM) with the appropriate buffer, respectively.
Determination of krel and Kapp of small laccases. LMꢀOH, a phenolic βꢀOꢀ4 ligninꢀmodel
compound (Figure S2), was used as ligninꢀmodel substrate to determine krel and Kapp of small
laccases. Spectrophotometric assays were performed at room temperature and monitored at 340
nm (ε340 = 1500 Mꢀ1 cmꢀ1 for the formation of vanillin, one of the LMꢀOH oxidation products,
Figure S9). Reaction mixtures (200 ꢂL) were buffered at varying pH (6ꢀ10) with 100 mM
potassium phosphate buffer (pH 6ꢀ8) and 100 mM sodium carbonate buffer (pH 9ꢀ10). It
contained 1ꢀ100 mM LMꢀOH (dissolved in DMF) as substrate and the enzyme (20ꢀ30 ꢂM) at
room temperature. The steadyꢀstate kinetic parameters were evaluated by fitting the initial rates
in MichaelisꢀMenten equation using GraphPad Prism (36).
ACS Paragon Plus Environment
12

Page 13 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
Ethanosolv lignin degradation by small laccase. In order to detect activity of small
laccase with a heterogeneous mixture of lignin building blocks, the enzyme was incubated for 16
h in the presence of ethanosolv lignin. The resultant cloudy reaction mixture with some
precipitate was mixed vigorously while adding an equal volume of 20% acetonitrile (CH3CN) in
water to a final concentration of 10% to produce a homogeneous suspension. The mixture
containing 10% CH3CN was then centrifuged for 10 min at 15,000g. Cleared supernatant was
applied to a C18 column (ꢂBondapakTM C18, 10 ꢂM 125 Å, 4.6 x 150 mm, Waters) and eluted
with a 10% isocratic gradient of CH3CN/H2O for 5 mL, followed by a linear gradient of 10ꢀ60%
for 25 mL at a flow rate of 1 ml/min and monitored at 285 nm.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Size exclusion chromatography was utilized to elucidate the relative size distribution of
small laccase degraded ethanosolv lignin. For size exclusion, the cloudy reaction mixture was
mixed with an equal volume of 60% CH3CN/H2O to solubilize the otherwise insoluble lignin
species. A sample (20 ꢂL) was then injected into size exclusion column (BioSuiteTM 125, 4 ꢂM
UHR SEC, 4.6 x 300 mm, Waters) with a mobile phase consisting of 30% CH3CN/H2O, eluted at
0.4 mL/min and monitored at 315 nm.
Crystallization and data collection for AMLAC. Two crystal forms of Amycolatopsis sp.
75iv2 small laccase (AMLAC) were obtained utilizing the Hampton Research sparse matrix
crystallization screens. Crystals were obtained both in the presence and absence of a nonꢀ
phenolic βꢀOꢀ4 lignin model compound (LMꢀOMe) [Figure S2]. Both crystal forms appeared
within two weeks of the initial setup in the Crystal Screen II condition #3 and Index Screen
condition #11. Centered orthorhombic crystals were obtained in the presence of the LMꢀOMe
substrate analogue in the first and rhombohedral ligandꢀfree crystals in the second condition,
respectively, using the hanging drop vapor diffusion method at 4 ˚C.
ACS Paragon Plus Environment
13

Biochemistry
Page 14 of 45
1
2
For the crystals of AMLAC•Cu2+•LMꢀOMe (PDB 3TA4), the protein solution contained
AMLAC (20 mg/mL) in 20 mM TrisꢀHCl (pH 7.9) and 20 mM of LMꢀOMe ligand; the
precipitant contained 25% ethylene glycol. Crystals appeared within two weeks and exhibited
diffraction consistent with the space group C2221, with three molecules per asymmetric unit.
Prior to data collection, the crystals were dipped in fresh mother liquor, followed by flash
cooling in liquid nitrogen; the ligand was not included in cryoprotectant solution.
3
4
5
6
7
8
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
For the crystals of AMLAC•Cu2+ (PDB 3T9W), the protein solution contained AMLAC
(20 mg/mL) in 20 mM TrisꢀHCl (pH 7.9); the precipitant contained 3.0 M NaCl and 100 mM
Hepes (pH 7.5). Crystals appeared within two weeks and exhibited diffraction consistent with the
space group R32, with one molecule per asymmetric unit. Prior to data collection, the crystals
were immersed in a cryoprotectant solution composed of 30% glycerol and of 70% of the
original mother liquor. After 30 seconds of incubation, the crystals were vitrified by immersion
in liquid nitrogen.
The diffraction data for AMLAC•Cu2+ was recorded at LSꢀCAT (Sector 21 IDꢀG,
Advanced Photon Source, Argonne, IL) using a MAR 300 CCD detector. The diffraction data for
AMLAC•Cu2+•LMꢀOMe was recorded at LSꢀCAT (Sector 21 IDꢀF, Advanced Photon Source,
Argonne, IL) using a MAR 225 CCD detector. Diffraction intensities were integrated and scaled
with the XDS package (37). Relevant data collection and data reduction statistics are given in
Table 4.
Crystallization and data collection for SVLAC. Two crystal forms were also obtained for
the Streptomyces viridosporus T7A small laccase (SVLAC). Approximately one month after the
initial setup of the Precipitant Synergy sparse matrix screen, orthorhombic crystals appeared in
condition #16, and trigonal crystals in condition #8, using the hanging drop vapor diffusion
ACS Paragon Plus Environment
14

Page 15 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
method at 9 ˚C. The trigonal crystal form was also subjected to soaking experiments with
acetovanillone (AV), a naturally occurring mediator (38, 39). Although cocrystallization with the
LMꢀOMe compound was attempted, no ligand was detected in resultant electron density maps
for either of the two crystal forms. The presence of the Zn2+ atom in the first crystal form was
established via the identities of its bindingꢀresidues (40). In addition, zinc content of the enzyme
was analyzed with Sciex Elan DRCe ICPꢀMS (Perkin Elmer) at the University of Illinois
Microanalytical Laboratory.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
For the crystals of SVLAC•Cu2+•Zn2+ (PDB 3TBB), the protein solution contained
SVLAC (20 mg/mL) in 50 mM HepesꢀK+ (pH 8.0) and 100 mM KCl; the precipitant contained
2.0 M Li2SO4, 5% polyethylene glycol 400, 100 mM MgSO4 and 100 mM acetate (pH 5.5).
Crystals appeared in approximately one month and exhibited diffraction consistent with the
space group I222, with three molecules per asymmetric unit. Prior to data collection, the crystals
were immersed in a cryoprotectant solution composed of 4.0 M Naꢀmalonate. After 30 seconds
of incubation, the crystals were vitrified by immersion in liquid nitrogen.
For the crystals of SVLAC•Cu2+ (PDB 3TAS) and SVLAC•Cu2+•AV (PDB 3TBC), the
protein solution contained SVLAC (20 mg/mL) in 20 mM TrisꢀHCl (pH 7.9); the precipitant
contained 5% 2ꢀpropanol and 2.5 M dibasic potassium phosphate/monobasic sodium phosphate
(pH 5.5). Crystals appeared in approximately one month and exhibited diffraction consistent with
the space group P3121, with three molecules per asymmetric unit. Prior to data collection, the
crystals of SVLAC•Cu2+ were immersed in a cryoprotectant solution composed of 70% of the
original mother liquor and 30% of glycerol. After 30 seconds of incubation, the crystals were
vitrified by immersion in liquid nitrogen. For SVLAC•Cu2+•AV, the ligandꢀfree crystals were
transferred into fresh mother liquor containing 20 mM acetovanillone and incubated at 9˚ C for
ACS Paragon Plus Environment
15

Biochemistry
Page 16 of 45
1
2
3
4
5
6
four hours. The crystals were vitrified as described earlier and the cryoprotectant solution did not
contain the ligand.
7
8
9
The diffraction data set for SVLAC•Cu2+•Zn2+ was recorded at LSꢀCAT (Sector 21 IDꢀG,
Advanced Photon Source, Argonne, IL) using a MAR 300 CCD detector. The diffraction data set
for the crystals of SVLAC•Cu2+ and SVLAC•Cu2+•AV were recorded at LSꢀCAT (Sector 21 IDꢀ
F, Advanced Photon Source, Argonne, IL) using a MAR 225 CCD detector. Diffraction
intensities were integrated and scaled with the XDS package (37). The data collection statistics
are given in Table 4.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Structure determination and model refinement for AMLAC and SVLAC. The structures of
both liganded and unliganded AMLAC and SVLAC were solved by molecular replacement with
PHASER (41), using the atomic coordinates of the small laccase from Streptomyces coelicolor
A3(2) (PDB 3CG8) as the search model. The partial solution from PHASER was then subjected
to automated model building with ARP/wARP (42), followed by multiple iterative cycles of
manual and automated rebuilding with COOT (43), interspersed with rounds of refinement using
PHENIX (44). Water molecules were automatically built with ARP/wARP, followed by manual
assessment in COOT. Final refinement statistics are provided in Table 4 and Table 5.
RESULTS AND DISCUSSION
Identification of the small laccase. The genome of Streptomyces coelicolor A3(2)
encodes a “small” laccase (SCLAC) that is homologous to fungal laccases, although the bacterial
laccase contains two domains and the larger fungal laccases contain three domains. The gene
encoding SCLAC is genome proximal (separated by eight genes) to a sevenꢀgene operon
encoding enzymes in the βꢀketoadipate pathway to the tricarboxylic acid cycle intermediates
acetylꢀCoA and succinylꢀCoA (45). Homologous gene clusters can be found in the genomes of
ACS Paragon Plus Environment
16

Page 17 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
several Streptomyces species, including Streptomyces lividans TK24, Amycolatopsis sp. 75iv2
and Streptomyces griseus NBRC 13350. Because lignin is composed of aromatic moieties, we
investigated whether SCLAC is involved in lignin degradation (24, 31) by disruption of the gene
encoding the small laccase by insertion of an apramycin resistance cassette, apr, using PCRꢀ
targeted Streptomyces gene replacement method, yielding the mutant designated SCꢁLAC.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Lignocellulose degradation by the small laccase. Quantitative studies of lignin
degradation are hampered by the lack of convenient assay methods due to the complexity and
heterogeneity of its polymeric structure (Figure S1). The mineralization of lignin can be
14
14
quantified by the conversion of Cꢀlabeled lignin to CO2. However, this method requires the
biosynthetic preparation of the isotopically labeled substrate and, therefore, may allow for the
incorporation of radioactivity into nonꢀlignin components of the lignocellulose, e.g. proteins,
aromatic acids esterified to hemicelluloses or to lignin and other aromatics such as lignans (8).
Also, a spectrophotometric assay recently was reported using chemically nitrated lignin, but the
assay sensitivity is low and we were unable to demonstrate lignin degradation using this method
(46). Monomeric and βꢀOꢀ4 dimeric lignin model compounds have been used, but these are not
perfect representations of the complex native lignin structures (47). Finally, some organisms
show lignin degradation activity only in the presence of cellulosic biomass, i.e. in native
lignocellulosic state (48). Therefore, we considered the use of native lignocellulose, a
combination of lignin, cellulose and hemicellulose, as an important realꢀworld criterion for
determining the lignin degradation activity of microorganisms.
Considering the molecular size and structure of lignin, it is reasonable to postulate that its
microbial degradation will involve extracellular (secreted) enzymes. Actinomycetes are known
to secrete enzymes that produce water soluble, polymeric lignin fragments from different
ACS Paragon Plus Environment
17

Biochemistry
Page 18 of 45
1
2
3
4
lignocellulosic materials.
These intermediates precipitate from aqueous solution upon
5
6
7
8
acidification and are known as acidꢀprecipitable polymeric lignin (APPL) (35). Quantification of
APPL production can provide a measure of the lignin degradation efficiency; chemical
characterization of the APPL has the potential to elucidate the molecular processes involved in
lignin degradation (35). We reasoned that if the small laccase is involved in lignin degradation,
less APPL would be produced by the SCꢁLAC mutant than by the wild type strain SCWT.
Also, chemical analysis of the APPL produced by the mutant and wild type strains would
provide a measure of the extent of lignin degradation and therefore the importance of the small
laccase in the APPL production.
9
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
APPL production assays were performed with the SCWT and SCꢁLAC mutant strains
using milled, airꢀdried lignocellulose from Miscanthus x giganteus as the source of lignin
substrate (3). Controls were performed using i) the medium alone, ii) the medium with
lignocellulose and iii) the medium with either the SCWT or SCꢁLAC mutant strain in the
absence of lignocellulose. The growth of SCWT and SCꢁLAC mutant strains were similar
under experimental conditions (the growth was measured by monitoring the OD at 600 nm).
After one week of incubation the residual lignocellulose and bacteria were removed by
°
centrifugation. The supernatants were acidified at 4 C to minimize acid hydrolysis and the
resulting precipitates (APPL) were dried and weighed. APPL was found only in the supernatants
of the cultures containing both bacteria and lignocellulose but not in any of the control
experiments (Figure 1). This indicates that APPL is the breakdown product of lignocellulosic
substrates in the presence of the SCWT and SCꢁLAC mutant strains and not a hydrolysis
product of lignocellulose.
ACS Paragon Plus Environment
18

Page 19 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
Although the SCꢁLAC strain produced APPL, the total amount of APPL was
significantly less (33.1%) than that produced by the wild type strain (Table 1). That the mutant
was deficient in APPL production may support the possible importance of the small laccase in
lignin degradation. The results of the compositional characterization and chemical analysis of the
APPLs produced by the SCWT and SCꢁLAC strains are shown in Table 2. The results are the
mean of triplicate analysis, except for ash (duplicate analysis) and nitrogen (single analysis). The
level of lignin degradation (Klason lignin) as measured from the total amount of APPL produced
by SCWT was much greater than that produced by the SCꢁLAC strain (Table 2). As a result of
the larger extent of lignin degradation by the SCWT strain a larger amount of carbohydrate
(glucan, xylan and arabinan) was released by the SCWT strain (15.9 mg) than by the SCꢁLAC
strain (11.9 mg), suggesting the hydrolytic enzymes had easier access to the carbohydrate
biomass. The acetyl content of the APPL quantitates the acetyl released from acetyl esters of
both lignin and hemicellulose; the larger amount of acetyl released by the SCWT strain again
suggests higher accessibility to the carbohydrate biomass due to the greater lignin degradation by
the SCWT strain. Ash and nitrogen measure the amounts of inorganic salts and protein present
in APPL, respectively, and were unchanged by the disruption of the gene encoding the small
laccase (Table 2).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Functional characterization of purified small laccases. To investigate further the
involvement of small laccases in lignin degradation, four small laccase enzymes, SCLAC,
SLLAC, SVLAC and AMLAC, were expressed and purified from E. coli with a high yield (15ꢀ
20 mg/L cell culture). SDSꢀPAGE of pure laccase enzymes show both monomeric and trimeric
form, an observation found previously (Figure S8) (24). They are very stable enzymes that are
active across a broad pH range (pH 3ꢀ10), compared to fungal laccases which are mostly active
ACS Paragon Plus Environment
19

Biochemistry
Page 20 of 45
1
2
3
4
5
6
7
8
9
at low pH (49, 50). Their activities were determined using ABTS (pH 3ꢀ5) and DMP (pH 6ꢀ10)
as substrates (data not shown). The high yields of the small laccases from Streptomyces using
E.coli as heterologous expression system and their stability over a broad pH range make them
excellent candidates for various industrial applications.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Oxidative degradation of lignin model compounds. Both phenolic and nonꢀphenolic
dimeric βꢀOꢀ4 lignin model compounds are widely used to study CαꢀCβ bond cleavage in lignin
degradation (8). The ability of small laccases to degrade/oxidize the dimeric βꢀOꢀ4 lignin model
compounds indirectly reflects their ability of lignin oxidative degradation, as more than 50% of
lignin structure is comprised of βꢀOꢀ4 bonds (3). Scheme S1 demonstrates a possible oxidative
degradation pathway of βꢀOꢀ4 lignin model compounds by laccases. It was postulated that
laccase is only responsible for the radical initiation step, with the downstream CꢀC bond
deconstruction the result of spontaneous reactions (51). The catalytic activities of the
Streptomyces laccases were determined by measuring the release of degradation product(s) due
to the CαꢀCβ bond cleavage/rearrangement of the LMꢀOH phenolic βꢀOꢀ4 lignin model
°
compound at pH 8.0 and at 25 C. A mixture of oxidative degradation products was formed,
which were not fully characterized. However, one of the products was identified and
characterized as vanillin which absorbs at 340 nm (Figure S9). Product(s) formation at this
wavelength was found to show MichaelisꢀMenten kinetics, and thereby the relative kcat (krel) and
apparent KM (Kapp) values were calculated for small laccases (Table 3). The low krel/Kapp (low
krel and high Kapp) values suggest that the βꢀOꢀ4 model compound is not the natural substrate,
prompting us to use the slightly modified native lignin substrate (ethanosolv lignin) for further
studies. The pH optimum for SCLAC against LMꢀOH was determined; the values of krel and
krel/Kapp were highest at pH 8.0 (Figure S3). Different mediators, ABTS and HOBt, were used to
ACS Paragon Plus Environment
20

Page 21 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
oxidize nonꢀphenolic βꢀOꢀ4 lignin model compound (LMꢀOMe) in presence of small laccases.
No CαꢀCβ bond cleavage were observed, instead there was very slow formation of the
corresponding ketone product [Figure 2, identified by MS (Figure S7)], a behavior previously
observed with fungal laccase (52). However, the rate of conversion of LMꢀOMe to quinone
product by small laccases was slower compared to fungal laccase from T. versicolor (data not
shown). This result suggests that even in the presence of mediators neither bacterial nor fungal
laccases were able to cleave the CαꢀCβ bond of LMꢀOMe compound, but they were able to
modify the structure by forming the quinone product at different rates. These activities explains
laccases’ role in enhancing the access of energy rich cellulose and hemicellulose by modifying
lignin structure (16).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Degradation/modification of ethanosolv lignin. To verify the activity of the laccases
against native lignin, ethanosolv lignin was used because of its better water solubility.
Ethanosolv lignin was prepared by preꢀtreating native Miscanthus lignocellulose with 80%
aqueous ethanol and 0.5% w/w (based on raw material dried mass) sulfuric acid as catalyst at
high temperature (2, 32). Ethanosolv lignin retains the same lignin and carbohydrate composition
as native lignocellulose except that the molecular weight is lower, which makes it more water
soluble and convenient for enzyme assays (2). After reacting ethanosolv lignin with purified
SCLAC for 16 h at 37 °C, the reaction mixture was passed through a reverse phase C18 column;
the elution profile is shown in Figure 3. As determined by LCꢀMS, most of the ionizable peaks
from this elution profile belong to a molecular weight range of 200ꢀ1000 Da. The major species
(~ 13 min) was determined, by Agilent 1100 LC/MSD Trap XCT Plus (LC/MS) at the University
of Illinois Roy J. Carver Biotechnology center, to be pꢀcoumaric acid ethyl ester with a
molecular weight of 192.08, an expected lignin degradation product and a natural laccase
ACS Paragon Plus Environment
21

Biochemistry
Page 22 of 45
1
2
3
4
5
6
7
8
9
mediator (53, 54). The chromatogram seems to indicate loss of soluble material, which can be
equated to polymerization of the smaller lignin units into larger pieces that are less soluble in
aqueous solution and were not retained during work up on the reaction mixture for C18
chromatography (vide infra). It is believed that laccases catalyze the oxidation of lignocellulosic
substrates to produce aryl cation radicals; these radicals spontaneously rearrange, leading to
further polymerization of degraded materials by the fission of carbonꢀcarbon or carbonꢀoxygen
bonds of the alkyl side chains or to the cleavage of aromatic rings (55, 56). Taking this into
account, our experiments aimed at reproducing natural lignin biodegradation in vitro indicate
that although the laccase is able to degrade lignin substrates, the products simultaneously
undergo competing radical polymerization.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
To probe this conclusion, sizeꢀexclusion chromatography was utilized to elucidate the
relative size distribution of ethanosolv lignin after reaction with SCLAC. The results are shown
in Figure 4. For this experiment, unlike the reverse phase C18 column chromatography
experiment, the cloudy reaction mixture was mixed with an equal volume of 60% CH3CN/H2O
to solubilize the polymerized lignin species. An observable shift towards larger molecular weight
lignin was observed, as was expected based on the loss of material in the C18 chromatograms
(vide supra). Due to the lack of appropriate molecular weight standards, the molecular weight
estimate is only qualitative. Nevertheless, we can conclude that during in vitro lignin degradation
there is competition between deꢀpolymerization and reꢀpolymerization, which would shift toward
deꢀpolymerization under in vivo conditions due to presence of a suitable radical quencher (56).
Considering the very complex structure of lignin and the subsequent complexity of reꢀ
condensation of reactive lignin degradation products, it is unlikely that this biopolymer can be
effectively degraded by one enzyme alone. In fungi, FADꢀdependent enzymes veratryl alcohol
ACS Paragon Plus Environment
22

Page 23 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
oxidase and glucose 1ꢀoxidase were considered to have coꢀoperative action with laccase to
prevent the reꢀpolymerization by reducing the reactive radical species (51, 56). For Streptomyces
discovery of such a coꢀoperative enzyme for prevention of reꢀpolymerization is an area of active
study.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Structure of small laccases. A multiple sequence alignment of all four small laccases shows that
all the active site residues involved in copper binding are conserved (Figure S4). These residues
are also highly conserved among other laccases and fourꢀcopper oxidases from fungi, plants and
bacteria although there is less similarity across the rest of the protein (24).
Laccases are known to bind four copper ions (II) in three different binding sites, each
characterized by unique spectroscopic properties and playing an important role in substrate
oxidation (31, 57ꢀ60). The paramagnetic Typeꢀ1 (T1) copper is coordinated by two His residues,
one Met residue and one Cys residue. The tight coordination of T1 copper to Cys residue is
responsible for an intense absorption band around 600 nm, giving the blue color to the enzyme.
Two His residues are coordinated to one paramagnetic Typeꢀ2 (T2) copper and three His
residues (six His residues total) are coordinated to each of the two Typeꢀ3 (T3) coppers. The T3
coppers are antiferromagnetically coupled with EPR silent pair absorption maximum at 330 nm.
The one T2 and two T3 coppers are arranged in a trinuclear cluster, which is believed to be the
site of molecular oxygen reduction (31, 57). The catalytic cycle is thought to be initiated by
oxidation of substrate near the T1 copper site by transfer of an electron from the substrate. In
total, four electrons (produced by oxidation of four substrates) are then sequentially transferred
from the T1 copper to the trinuclear cluster along a CysꢀHis pathway, where one oxygen
molecule is reduced to two water molecules to complete the cycle (31).
ACS Paragon Plus Environment
23

Biochemistry
Page 24 of 45
1
2
3
4
5
6
7
8
9
The structure of AMLAC was solved both in the presence (PDB 3TA4) and absence
(PDB 3T9W) of a nonꢀcleavable lignin βꢀOꢀ4 model compound LMꢀOMe (Figure S2). Although
the biologically active form of the enzyme is a trimer, the ligandꢀfree structure (space group
R32) contains one molecule per asymmetric unit. In this case the crystallographic symmetryꢀ
related molecules complete the trimeric composition. To date, the unliganded structure of
AMLAC reported here (PDB 3T9W) is the highest resolution structure of a small bacterial
laccase. The enzyme shares 58% sequence identity to the previously reported small laccase from
Streptomyces coelicolor A3(2) (PDB: 3CG8 and 3KW8) (SCLAC) and consists of an identical
twoꢀdomain fold with metalꢀbinding architecture (31).
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
The T1 copperꢀbinding site contains a highly ordered copper ion that is coordinated to
Cys 279, His 222, and His 284 with Met 298 as the axial ligand. The T2 copper is coordinated to
*
His 93 and His 225 (where the * denotes the residue from a symmetryꢀrelated polypeptide).
The first T3 copper is coordinated to His 95, His 147 and *His 280; the second T3 copper site is
*
made up of His 149 and His 278 and *His 227. The area between the T2 and T3 copper centers
contains strong positive electron density in the (FoꢀFc) omit map, suggestive of a bound dioxygen
molecule. However, the electron density is elongated so a hydrogen peroxide molecule may be
more appropriate. This dioxygen reduction intermediate, however, is not unusual and has
previously been reported in threeꢀdomain laccase structures with almost identical coordination
geometry (Figure S5) (61, 62).
Whereas the threeꢀdomain laccases contain a negatively charged Glu or Asp residue in
the tip of the access solvent channel, the water molecule coordinated to O1 of the peroxide is
bonded to a polar Gln 282 residue in the AMLAC structure. The ligandꢀbound structure of
AMLAC (PDB 3TA4) (space group C2221) contains three polypeptides in the asymmetric unit.
ACS Paragon Plus Environment
24

Page 25 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
The trimer contains three trinuclear and three mononuclear copper centers. Serendipitously, this
crystal form allows for the binding of the nonꢀphenolic lignin model compound LMꢀOMe, which
was included in the crystallization solution. Although the surface of the trimer lacks defined
cavities that may act as the binding pocket for smallꢀmolecule phenolic substrates, it is
conceivable that the rather flat trimer surface is optimal for binding large phenolic polymers.
Although it is questionable whether the binding of the model compound is biologically relevant,
its binding site is close to the T1 copper site, which is believed to be the main source of catalytic
activity for the oxidation of phenolic compounds (63, 64). The ligand in this crystal form is
trapped in a “wedge” between two crystallographic symmetryꢀrelated trimers (Figure 5). It is
also not possible to assess the orientation of the inert substrate molecule due to its threeꢀ
dimensional conformation, which is nearly symmetric (Figure 5). The ligand was modeled at an
occupancy of 0.46 as the molecule lies on a symmetry axis and occupies approximately half of
the site. The site lying between the T2 and T3 copper atoms that was occupied by a hydrogen
peroxide moiety in the ligandꢀfree structure contains a water molecule.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Structures of SVLAC were solved in two distinct crystal forms. SVLAC shares 87%
sequence identity to SCLAC and differs mainly in the sequence of the Cꢀterminal tail of the
enzyme. Both crystal forms are composed of asymmetric units that contain the biologically
active trimeric enzyme. However, regions that were ordered in the crystal form belonging to I222
space groups were disordered in the crystals belonging to P3121 space group due to crystal
packing. However, the hexagonal crystal form produced a condition similar to that of the trimeric
form of AMLAC, enabling the trapping of a substrate molecule at the crystallographic
symmetryꢀrelated trimer interface. The new feature that can be observed with the structure
ACS Paragon Plus Environment
25

Biochemistry
Page 26 of 45
1
2
3
4
5
6
belonging to the I222 space group is the extra metalꢀbinding site (PDB 3TBB), which arises from
an insertion of two extra histidine residues in the Cꢀterminus that SCLAC lacks.
7
8
9
Based on the results from the ICPꢀMS analysis, we hypothesize that this site contains a
zinc ion. The binding pocket for the zinc ion is formed by Glu 158 and His 159 of one
polypeptide and a *His 315 residue of the neighboring polypeptide. In all three zinc binding sites
in the trimer, the metal ion is within ~2.3Å of the εꢀnitrogen of the histidine residues and within
~2.6Å of the carboxylate oxygen of the glutamate moiety. In each of the three sites, the zinc ion
is also coordinated to at least one water molecule (Figure S6). .
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
In the structure, the monoꢀ and triꢀ nuclear copper sites are conserved. The T1 copperꢀ
binding site contains a highly ordered copper ion that is coordinated to Cys 283, His 226, and
*
His 288 with Met 293 as the axial ligand. The T2 copper is coordinated to His 97 and His 229.
The first T3 copper is coordinated to His 99, His 151 and *His 284; the second T3 copper site is
*
*
made up of His 153 and His 282 and His 231. The site that was occupied by a hydrogen
peroxide in the highꢀresolution structure of AMLAC contains a dioxygen molecule in all three
polypeptides of SVLAC. Interestingly, a molecule of acetate, a component of the crystallization
buffer, is coordinated to one of the T3 copper atoms in all three polypeptides. The trigonal
crystal form, which lacks the Cꢀterminal tail involved in zinc coordination, was an excellent
candidate for soaking experiments.
Although multiple natural mediators were tested in soaking experiments, only
acetovanillone (AV) yielded detectable electron density in the resultant structure. Although the
presence of AV is again most likely attributable to crystal packing, this phenolic substrate is
bound only ~10Å away from the T1 copper site. The binding of AV, when compared to the LMꢀ
OMe binding to AMLAC, seems to be more specific with multiple hydrogenꢀbonding
ACS Paragon Plus Environment
26

Page 27 of 45
Biochemistry
1
2
3
4
5
6
7
8
9
interactions to the SVLAC polypeptide. The molecule of AV is stacked against what appears to
be a molecule of Tris from the purification buffer (Figure 6). The Tris, however, is ordered only
in the liganded structure and is missing from the electron density in the apoꢀform of this crystal
form.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
Conclusions. We provided evidence that 1) strains of Streptomyces that encode the small
laccase have the potential to be useful in lignin degradation, and 2) the extracellular small
laccase plays an important role in this process. The small laccases from four different
Streptomyces were cloned, expressed and purified from heterologous sources and proved to be
stable and active against different substrates under versatile pH conditions. These enzymes were
able to degrade phenolic βꢀOꢀ4 lignin model compound without any mediators and rearrange
nonꢀphenolic βꢀOꢀ4 lignin model compound in presence of mediators, which provide in vivo
validation for their lignin degradation activity. However, APPL production was not completely
eliminated by the laccase deficient strain, so other, to be discovered, enzymes likely are also
involved in lignin degradation.
ACS Paragon Plus Environment
27

Biochemistry
Page 28 of 45
1
2
3
4
5
6
7
SUPPORTING INFORMATION
8
9
Primers used in this study (Table S1), Structural motif of lignin (Figure S1), Lignin model
compounds (Figure S2), Dependence of steadyꢀstate kinetic parameters of SCLAC on pH
(Figure S3), Multiple sequence alignment of small laccases (Figure S4), Coordination geometry
of the H2O2 intermediate bound to the trinuclear center copper atoms and water molecules in
AMLAC (Figure S5), The zinc metalꢀion binding site of SVLAC (Figure S6), ESIꢀMS showing
the formation of ketone derivative (Figure S7), SDSꢀPAGE of purified small laccases (Figure
S8), ESIꢀMS showing the formation oxidative products (Figure S9), Proposed oxidative
mechanism of lignin model compound (Scheme S1). “This material is available free of charge
via the Internet at http://pubs.acs.org.”
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
ACCESSION CODES
PDB entries:
3TA4 (AMLAC•Cu2+•LMꢀOMe), 3T9W (AMLAC•Cu2+), 3TBB
(SVLAC•Cu2+•Zn2+), 3TAS (SVLAC•Cu2+), 3TBC (SVLAC•Cu2+•AV)
ACS Paragon Plus Environment
28

Page 29 of 45
Biochemistry
1
2
3
4
REFERENCES
5
6
7
8
9
1.
2.
Adler, E. (1977) Lignin chemistry ꢀ past, present and future, Wood Sci. Technol. 11, 169ꢀ
218.
10
11
12
13
14
15
16
17
18
19
20
21
22
23
24
25
26
27
28
29
30
31
32
33
34
35
36
37
38
39
40
41
42
43
44
45
46
47
48
49
50
51
52
53
54
55
56
57
58
59
60
El Hage, R., Chrusciel, L., Desharnais, L., and Brosse, N. (2010) Effect of autohydrolysis
of Miscanthus x giganteus on lignin structure and organosolv delignification, Bioresour
Technol 101, 9321ꢀ9329.
3.
4.
Villaverde, J. J., Li, J., Ek, M., Ligero, P., and de Vega, A. (2009) Native lignin structure
of Miscanthus x giganteus and its changes during acetic and formic acid fractionation, J.
Agric. Food Chem. 57, 6262ꢀ6270.
Klinke, H. B., Thomsen, A. B., and Ahring, B. K. (2004) Inhibition of ethanolꢀproducing
yeast and bacteria by degradation products produced during preꢀtreatment of biomass,
Appl. Microbiol. Biotechnol. 66, 10ꢀ26.
5.
6.
7.
8.
9.
Palmqvist, E., and HahnꢀHagerdal, B. (2000) Fermentation of lignocellulosic
hydrolysates. I: inhibition and detoxification, Bioresour. Technol. 74, 17ꢀ24.
Palmqvist, E., and HahnꢀHagerdal, B. (2000) Fermentation of lignocellulosic
hydrolysates. II: inhibitors and mechanisms of inhibition, Bioresour. Technol. 74, 25ꢀ33.
Ander, P., and Eriksson, K. E. (1978) Lignin degradation and utilization by
microorganisms, Prog. Ind. Microbiol. 14, 1ꢀ58.
Kirk, T. K., and Farrell, R. L. (1987) Enzymatic "combustion": the microbial degradation
of lignin, Annu Rev Microbiol 41, 465ꢀ505.
Moyson, E., and Verachtert, H. (1993) Microbial degradation of lignin, Meded. ꢀ Fac.
Landbouwkd. Toegepaste Biol. Wet. (Univ. Gent) 58, 1651ꢀ1653.
ACS Paragon Plus Environment
29