Structural and functional studies of Aspergillus oryzae cutinase: Enhanced thermostability and hydrolytic activity of synthetic ester and polyester degradation

Article abstract of DOI:10.1021/ja9046697

Cutinases are responsible for hydrolysis of the protective cutin lipid polyester matrix in plants and thus have been exploited for hydrolysis of small molecule esters and polyesters. Here we explore the reactivity, stability, and structure of Aspergillus oryzae cutinase and compare it to the well-studied enzyme from Fusarium solani. Two critical differences are highlighted in the crystallographic analysis of the A. oryzae structure: (i) an additional disulfide bond and (ii) a topologically favored catalytic triad with a continuous and deep groove. These structural features of A. oryzae cutinase are proposed to result in an improved hydrolytic activity and altered substrate specificity profile, enhanced thermostability, and remarkable reactivity toward the degradation of the synthetic polyester polycaprolactone. The results presented here provide insight into engineering new cutinase-inspired biocatalysts with tailor-made properties.

Full text of DOI:10.1021/ja9046697

Published on Web 10/07/2009
Structural and Functional Studies of Aspergillus oryzae
Cutinase: Enhanced Thermostability and Hydrolytic Activity of
Synthetic Ester and Polyester Degradation
Zhiqiang Liu,† Yuying Gosser,‡ Peter James Baker,† Yaniv Ravee,† Ziying Lu,‡
Girum Alemu,‡ Huiguang Li,§ Glenn L. Butterfoss,| Xiang-Peng Kong,§
Richard Gross,† and Jin Kim Montclare*,†,⊥
Department of Chemical and Biological Sciences, Polytechnic Institute of New York UniVersity,
Brooklyn, New York 11201, Pathways Bioinformatics & Biomolecular Center, The City College,
CUNY, New York, New York 10031, Department of Biochemistry, NYU School of Medicine, 550
First AVenue, New York, New York 10016, Department of Biology and Computer Science, New
York UniVersity, New York, New York 10003, Department of Biochemistry, SUNY-Downstate
Medical Center, Brooklyn, New York 11203
Received June 8, 2009; E-mail: jmontcla@poly.edu
Abstract: Cutinases are responsible for hydrolysis of the protective cutin lipid polyester matrix in plants
and thus have been exploited for hydrolysis of small molecule esters and polyesters. Here we explore the
reactivity, stability, and structure of Aspergillus oryzae cutinase and compare it to the well-studied enzyme
from Fusarium solani. Two critical differences are highlighted in the crystallographic analysis of the A.
oryzae structure: (i) an additional disulfide bond and (ii) a topologically favored catalytic triad with a continuous
and deep groove. These structural features of A. oryzae cutinase are proposed to result in an improved
hydrolytic activity and altered substrate specificity profile, enhanced thermostability, and remarkable reactivity
toward the degradation of the synthetic polyester polycaprolactone. The results presented here provide
insight into engineering new cutinase-inspired biocatalysts with tailor-made properties.
Introduction
Cutinases are R/ꢀ hydrolases commonly secreted by fungal
phytopathogens that enable them to penetrate the protective
surface cutin layer of plants.15-19 Cutin is an insoluble, cross-
linked, lipid-polyester matrix comprised of n-C17 and n-C18
hydroxy and epoxy fatty acids that serve as a barrier against
dehydration and invasion. Cutinases hydrolyze and break down
these complex polyesters into smaller hydroxyacids.15,19
Because of the ability of cutinases to hydrolyze cutin, they
have been exploited for reactions with small molecule esters
and synthetic polyesters.1 The utility of cutinase or any enzyme
With environmental concerns over waste build-up and
limited petroleum resources, the demand has never been
greater for enzymatic or “green” approaches for efficient
chemical synthesis and degradation reactions.1-3 Several
enzymes have been exploited not only to perform stereo- and
regioselective chemical transformations on small molecules
but also to break down or modify synthetic polymers useful
for applications in chemical, pharmaceutical, and textile
industries,2,4,5 Serine hydrolases have been employed most
extensively for these biotransformations.1,6-14
(8) Borreguero, I.; Carvalho, C. M. L.; Cabral, J. M. S.; Sinisterra, J. V.;
Alcantara, A. R. J. Mol. Catal. B: Enzym. 2001, 11, 613–622.
(9) Bogel-Lukasik, R.; Lourenco, N. M. T.; Vidinha, P.; da Silva, M.;
Afonso, C. A. M.; da Ponte, M. N.; Barreiros, S. Green Chem. 2008,
10, 243–248.
(10) Chen, B.; Hu, J.; Miller, E. M.; Xie, W. C.; Cai, M. M.; Gross, R. A.
Biomacromolecules 2008, 9, 463–471.
(11) Kulshrestha, A. S.; Gao, W.; Fu, H. Y.; Gross, R. A. Biomacromol-
ecules 2007, 8, 1794–1801.
(12) Chen, B.; Miller, E. M.; Miller, L.; Maikner, J. J.; Gross, R. A.
Langmuir 2007, 23, 1381–1387.
† Polytechnic Institute of New York University.
‡ The City College, CUNY.
§ NYU School of Medicine.
| New York University.
⊥ SUNY-Downstate Medical Center.
(1) Carvalho, C. M. L.; Aires-Barros, M. R.; Cabral, J. M. S. Biotechnol.
Bioeng. 1999, 66, 17–34.
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38.
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ecules 2006, 7, 3093–3097.
(14) Hu, J.; Gao, W.; Kulshrestha, A.; Gross, R. A. Macromolecules 2006,
39, 6789–6792.
(3) Maeda, H.; Yamagata, Y.; Abe, K.; Hasegawa, F.; Machida, M.;
Ishioka, R.; Gomi, K.; Nakajima, T. Appl. Microbiol. Biotechnol. 2005,
67, 778–788.
(4) Costa, L.; Brissos, V.; Lemos, F.; Ribeiro, F. R.; Cabral, J. M. S.
Bioprocess Biosyst. Eng. 2008, 31, 323–327.
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(6) Masaki, K.; Kamini, N. R.; Ikeda, H.; Iefuji, H. Appl. EnViron.
Microbiol. 2005, 71, 7548–7550.
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Mol. Biol. 1997, 268, 779–799.
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A R T I C L E S
Liu et al.
60 to 0.9375 µM were used for the kinetic study.21,22 Assays were
performed using a final concentration of 1.1 µM of A. oryzae and
F. solani cutinases in 14.5 mM Tris-HCl buffer pH 7.5, 0.75%
glycerol. Since pNPA and pNPH were dissolved in DMSO, pNPB
in methanol, and pNPV in TritonX, there was approximately 25%
DMSO, methanol or 0.5% Triton X in the final mixtures.23
Reactions were initiated by the addition of enzyme, and reaction
progress was monitored spectrophotometrically (Molecular Devices
Spectramax M2) at 405 nm. Softmax Pro v5 software was used to
analyze data. Enzyme-catalyzed initial rates were corrected by
subtracting background hydrolysis rate. All reactions were per-
formed in triplicate. The Km and kcat values were determined by a
double-reciprocal Lineweaver-Burk plot (1/V vs 1/[pNPA/pNPB/
pNPV/ pNPH]) (Supporting Information).
Thermoactivity. The thermoactivity or residual activity of
cutinases was investigated by incubating enzymes at temperatures
ranging from 25 to 60 °C at a final concentration of 1.1 µM in
14.5 mM Tris-HCl buffer pH 7.5, 0.75% glycerol. The incubation
took place in an Eppendorf Thermomixer at the specified temper-
ature with a constant mixing of 350 rpm for 1 h. The samples were
incubated on ice for 5 min followed by incubation at room
temperature for 15 min. Reactions were initiated by addition of 30
µM pNPB, monitored at 405 nm for 1 min, and performed in
triplicate. The data obtained was presented as activity without
normalization.
for chemical reactions is commonly limited by intolerance to
high temperatures and the constraints of the substrate recognition
pocket.1 Thus, the identification of enzymes exhibiting enhanced
thermostability and altered reactivity for various substrates
would greatly expand the potential of cutinase for industrial
and environmental applications.
Aspergillus oryzea is a filamentous fungus that has been
employed in fermentation to produce traditional consumable
products such as rice wine, soybean paste, and soy sauce in the
food industry for approximately 1000 years.9 It has been used
recently as an expression host for recombinant proteins20 and
to degrade poly(butylene succinate) (PBS) as well as emulsified
poly(butylenes succinate-co-adipate) (PBSA).3 Because of the
remarkable ability of A. oryzae cutinase to readily break down
such synthetic plastics,3 it represents an excellent target to
perform detailed structure-activity analysis. Here we report the
reactivity, stability, and crystal structure of A. oryzea cutinase
and perform a comparison to the well-characterized Fusarium
solani cutinase.
Methods
Enzyme Expression. The cutinase gene was expressed in Pichia
pastoris, and recombinant cutinase was produced by using the strong
methanol-induced AOX1 promoter. Single colonies were picked
and cultured in BMGY medium (g/L) composed of 5 g of yeast
extract, 10 g of peptone, supplemented with 50 mL of 1 M KH2PO4
buffer pH 6.0, 1.7 g of yeast nitrogen base, 5 g of ammonium
sulfate, 5 mL of glycerol, 500 × l mL biotin, and 96 × 5.2 mL
histidine. Precultures of P. pastoris harboring cutinase genes were
incubated at 30 °C, 200 rpm, overnight. After centrifugation at 6,000
rpm for 10 min, cells were transferred into a parallel fermentor
(DASGIP, Germany) containing 1 L of basal salt medium composed
of glycerol 40 mL/L, CaSO4 0.9 g/L, K2SO4 14.67 g/L,
MgSO4 ·7H2O 11.67 g/L, (NH4)2SO4 9 g/L, 12 mL/L hexameta-
phosphate, trace salts (cupric sulfate ·5H2O 6.0 g/L, sodium iodide
0.08 g /L, manganese sulfate H2O 3.0 g/L, sodium molybdate ·2H2O
0.2 g/L, boric acid 0.02 g/L, cobalt chloride 0.5 g/L, zinc chloride
20.0 g/L, ferrous sulfate ·7H2O 65 g/L, biotin 0.2 g/L). The constant
dissolved oxygen was set to 40%, glycerol (50%, v/v) feeding time
was 6 h, and the rate of methanol feeding was 5 mL/h. When the
activity reached its maximum, the fermentation was stopped. After
centrifugation, the supernatant was collected for further use.
Enzyme Purification. Fermentation broth was centrifuged at
8,000 rpm for 10 min at 4 °C, and then supernatant was concentrated
about 10 times using an ultrafiltration unit (Millipore TFE system)
with a 10 kDa membrane. Cutinase was purified by FPLC using
VISION Workstation (Applied Biosystems Co.) with a 16 mmD/
100 mmL POROS MC 20 µm column (Applied Biosystems Co.).
The metal site in the column was saturated with NiCl2 solution
according to operating instructions for the column. Approximately
50 mM NaH2PO4 buffer with 0.5 mM imidazole (pH 8.0) was used
as a starting buffer, and 50 mM NaH2PO4 buffer with 100 mM
imidazole (pH 8.0) was used as an elution buffer at a flow rate of
30 mL/min. Samples filtered by a 0.45 µm filter were loaded onto
the column. Approximately 2 column volumes (CVs) of starting
buffer were run to wash out any proteins that were bound
nonspecifically to the column, and then 3 CVs of elution buffer
with imidazole were run with a concentration linear gradient from
0 to 100 mM. The peak fraction in the gradient step was collected.
The collected samples were desalted by an ultrafiltration unit with
a 10 kDa membrane and then freeze-dried. SDS-PAGE analysis
of the purified proteins was performed (Supporting Information).
Kinetic Analysis for pNP Ester Substrates. The pNP esters
(pNPA, pNPB, pNPV, and pNPH) with concentration ranging from
Circular Dichroism (CD) Measurements. CD spectra were
recorded on a JASCO J-815 Spectropolarimeter using Spectra
Manager 228 software. Temperature was controlled using a Fisher
Isotemp Model 3016S water bath. A protein concentration of 29
µM in 10 mM sodium phosphate buffer, pH 8.0 was used for both
cutinases. Data were collected at 1-nm intervals from 190 to 250
nm for wavelength scans and 0.3 °C/min from 4 to 85 °C for
temperature scans in duplicate (Supporting Information). Small
signaling arising from buffer was subtracted.
Thermodynamic Parameters by Differential Scanning
Calorimetry (DSC). Calorimetric measurements of melting tem-
peratures (Tm) were carried out on a NanoDSC differential scanning
calorimeter (TA Instruments) with a sample cell volume of 0.3 mL.
Unfolding data of both cutinases were obtained by heating the
samples, at a concentration 5 mg/mL, from 4 to 80 °C at a rate of
1 °C/min in duplicate (Supporting Information). The protein samples
were present in water. Water was used in the reference cell to obtain
the molar heat capacity by comparison. The observed thermograms
were baseline corrected, and normalized data were analyzed using
NanoAnalyze software.
Degradation of Polymer Thin Films. Thin films of poly(ε-
caprolactone) (PCL) were cut to 1.0 cm × 1.0 cm with an
approximate thickness of 250 µm (30-35 mg) and placed in 20
mL scintillation screw cap vials containing 2.5 mL of 100 mM
Tris-HCl buffer pH 8.0 with a final concentration of 8.8 µM
enzyme. The control vials contained a film with buffer solution.
All measurements were performed in triplicate, incubated for 6 h
at 40 °C in an incubator shaker at 200 rpm, and weighed after
drying.
Crystallization. The protein was crystallized by mixing equal
volumes of protein solution (15 mg/mL, in 100 mM Tris buffer
pH 8.5) with mother liquor (30% PEG2KMME, 100 mM potassium
thiocyanate) at 296 K. The screening was conducted with 96
crystallization conditions at 296 K using the hanging drop vapor
diffusion technique. Crystals appeared within 10-15 days.
Structure Determination. X-ray diffraction data of the crystal
were collected at beamline X4A (λ ) 0.96785 Å) of the synchrotron
light source in the Brookhaven National Laboratory. Prior to data
(21) Pedersen, S.; Nesgaard, L.; Baptista, R. P.; Melo, E. P.; Kristensen,
S. R.; Otzen, D. E. Biopolymers 2006, 83, 619–629.
(22) Goncalves, A. M.; Serro, A. P.; Aires-Barros, M. R.; Cabral, J. M. S.
Biochim. Biophys. Acta 2000, 1480, 92–106.
(20) Christensen, T.; Woeldike, H.; Boel, E.; Mortensen, S. B.; Hjortshoejk,
(23) Camacho, R. M.; Mateo, J. C.; Gonza´lez-Reynoso, O.; Prado, L. A.;
Co´rdova, J. J. Ind. Microbiol. Biotechnol. 2009, 36, 901–909.
K.; Thim, L.; Hansen, M. T. Biotechnology 1988, 6, 1419–1422.
9
15712 J. AM. CHEM. SOC. VOL. 131, NO. 43, 2009

Structural and Functional Studies of A. oryzae Cutinase
A R T I C L E S
Scheme 1. (a) Synthetic Ester and (b) Polyester Used As
collection, the crystal was soaked for 5 min in a mixture of the
mother liquid (30% PEG2000MME and 100 mM potassium
thiocyanate) and 15% glycerol for cryoprotection. A data set of
1.75 Å resolution was collected from a single crystal at 100 K with
98.5% completeness. This data set was processed using HKL2000
software.24 The crystal belongs to the trigonal space group P3221
with one subunit in the asymmetric unit. The unit cell parameters
were a ) 45.299 Å, b ) 45.299 Å, c ) 157.111 Å and R ) 90.00°,
ꢀ ) 90.00°, ꢁ ) 120.00°. The structure was solved by molecular
replacement using the Molrep program in the CCP4 suite.25 The
structure of the native F. solani cutinase (PDB accession number
1CEX) was used as the search model. The initial model was built
using O program. The structure was refined with CNS.26 The
iterative model building and refinement was carried out to a final
R-work factor of 19.4% and a final free-R factor of 19.9%. The
stereochemistry of the final structure was verified using the
Procheck_NT and Sfcheck programs in the CCP4 suite.25 Procheck
validation of local geometry revealed that 91.5% of residues are
located in the most favored regions of the Ramachandran plot, 7.2%
are in the additional allowed regions, and 1.3% are in the generously
allowed region. The electron densities of the first 10 N-terminal
residues (17-25) were not observed. Structural analysis was
performed using UCSF Chimera software (http://www.cgl.ucsf.edu/
chimera).27 Electrostatic surface rendering was performed using
ICN and the groove by the active site was generated by the
PocketFinder function.28 For all above analysis 1CEX pdb file was
used for F. solani cutinase.
Modeling. 1CEX and the A. oryzae cutinase crystal structure
presented here were modeled with the butyrate, valerate, and
hexanoate esters of the active site serine. Hydrogens were added,
and starting structures of the serine esters were generated by
sampling all combinations of -60°, 60°, and 180° for each ꢁ angle
(with the exception of the ester C-O torsion, which was sampled
at -180° and 180°). This generated 162, 486, and 1458 conforma-
tions for the butyrate, valerate, and hexanoate esters, respectively.
The modified serine residues were then subject to 11 × 20 steps of
conjugate gradient energy minimization using the CHARMM22
potential29 while keeping all other protein atoms fixed. Modeling
was done using the SIGMA package.30
Substrates for Hydrolytic Reactions with Cutinasea
a Abbreviations: p-nitrophenylacetate, pNPA; p-nitrophenylbutyrate,
pNPB; p-nitrophenylvalerate, pNPV; p-nitrophenylhexanoate, pNPH; poly-
caprolactone, PCL.
on cutinase activity was studied via Michaelis-Menten kinetics.
The A. oryzea cutinase demonstrated a preference for pNPV
resulting in a Km of 0.04 ( 0.01 µM, followed by pNPB and
pNPH with Km’s of 0.21 ( 0.04 and 0.29 ( 0.09 µM,
respectively. The short pNPA chain yielded a >120-fold higher
Km of 4.96 ( 0.11 µM (Table 1). The highest catalytic efficiency
was observed for pNPB and pNPV with a kcat/Km of 3.49 (
0.51 and 3.32 ( 0.74 µM-1 min-1 (Table 1). This was followed
by pNPH and PNPA with a kcat/Km of 1.34 ( 0.48 and 0.07 (
0.01 µM-1 min-1, respectively. This is in stark contrast to F.
solani cutinase, where an overall preference for short chain
substrate pNPA was highest. Although recognition for all three
substrates was relatively similar with Km’s ranging from 0.67
( 0.23 to 1.50 ( 0.19 µM, the catalytic efficiencies reveal a
strong preference for pNPA with a kcat/Km of 2.53 ( 1.11 µM-1
min-1 followed by pNPV, pNPB, and pNPH, respectively (Table
1). For pNPV, pNPB and pNPH, a 6-, 10-, and 20-fold decrease
in kcat/Km was observed relative to pNPA.
Results
Higher Thermostability of A. oryzae Cutinase. To investigate
the ability of the two enzymes to function at higher temperatures,
we examined the residual activities for pNPB hydrolysis for a
range of temperatures. Selection of pNPB was based on the
demonstration of reasonable reactivity by both cutinases. The
A. oryzea cutinase exhibited a higher activity for pNPB relative
to that of F. solani at room temperature, corresponding to
previous kinetic measurements. The level of activity was
maintained even up to 40 °C in the case of A. oryzea cutinase
(Figure 1a). By contrast, F. solani enzyme revealed a 40% drop
at 30 °C and demonstrated a continuous decline in activity as
a function of increasing temperature. Surprisingly, under all
temperatures investigated, A. oryzea cutinase displayed higher
overall residual activity when compared to that of F. solani,
suggesting an improved tolerance to heat. This is corroborated
by thermostability measurements by CD in which A. oryzea
cutinase exhibited a higher melting temperature (Tm) of 59 °C,
whereas the Tm of F. solani is 56 °C (Table 1). This increase in
Tm is manifested by a 39.6 kJ/mol increase in enthalpy by A.
oryzea cutinase as measured by DSC (Table 1).
Altered Specificity of A. oryzae Cutinase for pNP Esters.
Initially, the A. oryzea and F. solani cutinases were assessed
for hydrolytic activity on a set of model p-nitrophenylester
substrates: p-nitrophenylacetate (pNPA), p-nitrophenylbutyrate
(pNPB), p-nitrophenylvalerate (pNPV), and p-nitrophenylhex-
anoate (pNPH) (Scheme 1a).3,10 The influence of chain length
(24) Otwinowski, Z.; Minor, W. Methods Enzymol. 1997, 276, 307–326.
(25) Bailey, S. Acta Crystallogr., Sect. D: Biol. Crystallogr. 1994, 50, 760–
763.
(26) Brunger, A. T.; Adams, P. D.; Clore, G. M.; DeLano, W. L.; Gros,
P.; Grosse-Kunstleve, R. W.; Jiang, J. S.; Kuszewski, J.; Nilges, M.;
Pannu, N. S.; Read, R. J.; Rice, L. M.; Simonson, T.; Warren, G. L.
Acta Crystallogr., Sect. D: Biol. Crystallogr. 1998, 54, 905–921.
(27) Pettersen, E. F.; Goddard, T. D.; Huang, C. C.; Couch, G. S.;
Greenblatt, D. M.; Meng, E. C.; Ferrin, T. E. J. Comput. Chem. 2004,
25, 1605–1612.
(28) Abagyan, R.; Totrov, M.; Kuznetsov, D. J. Comput. Chem. 2004, 15,
488–506.
(29) MacKerell, A. D.; Bashford, D.; Bellott, M.; Dunbrack, R. L.;
Evanseck, J. D.; Field, M. J.; Fischer, S.; Gao, J.; Guo, H.; Ha, S.;
Joseph-McCarthy, D.; Kuchnir, L.; Kuczera, K.; Lau, F. T. K.; Mattos,
C.; Michnick, S.; Ngo, T.; Nguyen, D. T.; Prodhom, B.; Reiher, W. E.;
Roux, B.; Schlenkrich, M.; Smith, J. C.; Stote, R.; Straub, J.; Watanabe,
M.; Wiorkiewicz-Kuczera, J.; Yin, D.; Karplus, M. J. Phys. Chem. B
1998, 102, 3586–3616.
Higher Reactivity of A. oryzae Cutinase for Synthetic
Polyester. Since the natural function of cutinase is to degrade
the complex biopolyester of cutin,15 we assessed the ability of
both A. oryzae and F. solani enzymes to react on the surrogate,
well-defined polyester, poly ε-(caprolactone) (PCL) (Scheme
1b). PCL is known to be degraded by various microorganisms
including fungi.31,32 Murphy and co-workers demonstrated that
(30) Mann, G.; Yun, R. H.; Nyland, L.; Prins, J.; Board, J.; Hermans, J.,
Computationalmethodsformacromolecules:ChallengesandApplications-
Proceedings of the 3rd International Workshop on Algorithms for
Macromolecular Modeling; Schlick, T., Gan, H. H., Eds.; Springer
Verlag: Berlin, 2002; pp 129-145.
9
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A R T I C L E S
Liu et al.
Table 1. Kinetic and Thermodynamic Parameters
pNPA
pNPB
Km (µM-1>) kcat/Km (µM-1 min-1
pNPV
pNPH
cutinase
Km (µM-1
)
kcat/Km (µM-1 min-1
)
)
Km (µM-1
)
kcat/Km (µM-1 min-1
)
Km (µM-1
)
kcat/Km (µM-1 min-1
)
Tm (°C) ∆H (kJ/mol)
F. solani 0.67 ( 0.23
A. oryzae 4.96 ( 0.1
2.53 ( 1.11
0.07 ( 0.01
1.26 ( 0.28
0.21 ( 0.04
0.26 ( 0.06
3.49 ( 0.51
1.48 ( 0.56
0.04 ( 0.01
0.61 ( 0.40
3.32 ( 0.74
1.50 ( 0.19
0.29 ( 0.09
0.14 ( 0.02
1.34 ( 0.48
56
59
562.21
622.61
of residues 87-93 within helix 3, and residues 186-194 that
represents the loop between helices 9 and 10 as well as the first
3 residues of the latter. A. oryzea cutinase bears an oxyanion
hole composed of Ser48 and Gln127 backbone amides that are
critical in polarizing the ester bond of the substrate and
stabilizing the transition state of the formed substrate oxyanion
(Figure 2a).12
Upon overlay, the A. oryzea and F. solani structures have a
similar fold with an overall rms deviation of 1.02 Å, main chain
deviation of 0.87 Å, and CR deviation of 0.84 Å (Figure 3a).
However, the A. oryzae enzyme bears several structural features
that differ significantly from that of F. solani. Comparison of
the two sequences reveal that the A. oryzea cutinase is shorter
than that of F. solani as exhibited by smaller loops in the N-
and C-terminal regions as well as a missing ꢀ-strand in the A.
oryzae structure (Figures 2a and 3a). Although the F. solani
cutinase possesses 6 ꢀ-strands and 10 R-helices, the A. oryzae
structure bears the 5 ꢀ-strands and the same number of R-helices.
The N-terminal region (residues Thr26 to Asp30), the loops in
between helix 1 and strand 1 (residues Gly35 to Pro49), as well
as helix 2 and strand 2 (residues Ser71 to Asp73), helix 10
(residues Ser199 to Asp203), and C-terminal residues beyond
helix 10 based on the A. oryzae structure deviate significantly
in the overlay as highlighted in Figure 3a.
Distinct Disulfide Bond. The A. oryzae structure contains a
unique disulfide bond between Cys63 and Cys76 that ties helix
2 to strand 2 of the central ꢀ-sheet (Figure 2a). This disulfide
bond has not been previously reported for any cutinase or
hydrolase structures.7,12-17 The other two disulfide bonds,
Cys37-Cys115 connecting the loop between helix 1 and strand
1 with the loop between helix 4 and strand 3 and
Cys177-Cys184 linking the loop following strand 5 to helix
9, are well-conserved in previously published cutinase structures,
including that from F. solani.14 Sequence analysis suggests that
this disulfide bond is unique for the cutinases from the
Aspergillus family (sharing sequence identity of 50-77% with
A. oryzae) and a few other filamentous fungi, such as Neosa-
rtorya fischeri (53% sequence identity) and Emericella nidulans
52% sequence identity) (Figure 2a). The cutinases from F. solani
and Glomerella cingulata represent another group of filamentous
fungi that do not have this special disulfide bond, although they
share about 50% sequence identity with the A. oryzae enzyme.
Figure 1. (a) Thermoactivity profile as measured in terms of residual
activity for pNPB upon incubation at increasing temperatures. (b) Weight
loss measurements of defined PCL films.
F. solani strains bearing cutinase were able to degrade PCL
and use it as a carbon source as well as to induce the production
of cutinase,33 suggesting that the cutinase should be active for
PCL hydrolysis in vitro. PCL films with dimensions of 1 cm2
and 250 µm thickness were incubated at 40 °C in the presence
of either A. oryzea or F. solani cutinase in 100 mM Tris-HCl
at pH 8.0. Remarkably, nearly complete degradation of 87%
PCL was achieved within 6 h in the presence of A. oryzea
enzyme. Degradation of PCL films by F. solani cutinase
proceeded more slowly, reaching 30% degradation by 6 h
(Figure 1b). These data support the results on the aforementioned
substrate specificity in which the A. oryzea cutinase preferred
longer chain esters such as pNPB and pNPH, since PCL
represents a longer chain polyester. Furthermore, this supports
the thermoactivity data in which a dramatic loss in activity was
observed for the F. solani enzyme at 40 °C while that of A.
oryzae maintained a high level activity.
Crystal Structure of A. oryzae Cutinase and Comparison
to That of F. solani. To understand the observed reactivity and
stability differences, the A. oryzea cutinase structure was
determined by molecular replacement method and refined to
1.75 Å resolution (PDB ID:3GBS, Supporting Information). The
crystal structure revealed a monomeric protein with an R/ꢀ fold
hallmarked by a central ꢀ-sheet of 5 parallel strands surrounded
by 10 R-helices. As a member of the R/ꢀ hydrolases, this
enzyme possesses an active site composed of the catalytic triad
residues Ser126, Asp181, and His194 (Figure 2a). The catalytic
site is surrounded by the two hydrophobic surfaces composed
Continuous Groove by the Active Site. Although the catalytic
triad is sequentially conserved in both cutinases, they are
topologically positioned differently. In the A. oryzea structure,
His194 Nδ1 to Asp181 Oδ2 is 2.71 Å, and the Ser126 Oγ to
His194 Nε2 is 2.63 Å (Figure 3b), which are within hydrogen
bond distance. The corresponding distances in the F. solani
cutinase are 2.84 Å for Asp175 Oδ2 to His188 Nδ1 and 2.98
Å from Ser120 Oγ to His188 Nε2, which is slightly larger than
the hydrogen bond distance presented in the A. oryzae struc-
ture.16 This average distance of 2.98 Å was calculated on the
basis of the presence of two possible positions of the Ser120
Oγ in the native F. solani crystal structure: where the A form
possesses a 73% occupancy and the B form a 27% occupancy
(PDB ID:1CEX, Figure 3b). Moreover, the A. oryzae cutinase
(31) Benedict, C. V.; Cook, W. J.; Jarrett, P.; Cameron, J. A.; Huang, S. J.;
Bell, J. P. J. Appl. Polym. Sci. 1983, 28, 327–334.
(32) Nishida, H.; Tokiwa, Y. J. EnViron. Polym. Degrad. 1993, 1, 227–
233.
(33) Murphy, C. A.; Cameron, J. A.; Huang, S. J.; Vinopal, R. T. Appl.
EnViron. Microbiol. 1996, 62, 456–460.
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15714 J. AM. CHEM. SOC. VOL. 131, NO. 43, 2009

Structural and Functional Studies of A. oryzae Cutinase
A R T I C L E S
Figure 2. (a) Sequence alignment of A. oryzae cutinase with cutinases originating from representative filamentous fungi with sequence identity g50%. The
R-helical regions are highlighted in red, ꢀ-strands in yellow, catalytic residues in magenta, oxyanion hole residues in cyan, and cysteines bearing disulfide
bonds in yellow. Residues identical in all sequences in the alignment are labeled with “*”, residues with conserved substations are labeled with “:”, and
semiconserved substitutions with “.”. The alignment was performed using NCBI BLAST (http://blast.ncbi.nlm.nih.gov/) and EBI ClustalW (http://www.ebi.ac.uk/
Tools/clustalw2/). (b) Ribbon structure rendering of A. oryzae cutinase with the helices, strands, and catalytic residues labeled as in the alignment. The
disulfide bonds between the cysteines are displayed in yellow. (c) Stereoview of A. oryzae cutinase with catalytic triad in magenta and oxyanion hole in
cyan. Abbreviations: Aspergillus oryzae, A.ory; Aspergillus fumigatus, A.fum; Aspergillus claVatus, A.cla; Emericella nidulans, E.nid; Neosartorya fischeri,
N.fis; Glomerella cingulata, G.cin; Fusarium solani, F.sol.
bears a longer and deeper groove by the active site relative to
that of F. solani as demonstrated in Figure 3c. While the A.
oryzae structure reveals a continuous groove that spans across
the active site, the F. solani structure shows a narrow, shallower
groove that abruptly stops. Models of both cutinases with the
esters of the active site demonstrate the presence of two
gatekeeper residues (Leu87 and Val190) in A. oryzae and (Leu81
and Val184) in F. solani that play a role in substrate recognition
(Figure 4), The two residues in A. oryzae residues are 9.87 Å
(Cꢀ to Cꢀ) apart, whereas the corresponding residues of the F.
solani cutinase are separated by 8.74 Å. This results in the alkyl
chain populations in A. oryzae laying across the wider groove
region surrounding the active site, whereas for the F. solani
model, there is a preponderance of alkyl chain structures that
are oriented down the narrow tail of the groove.
Discussion
The molecular detail provided by the crystal structure
elucidates the observed discrepancy in the activity and specificity
of the two cutinases. The preference for the A. oryzea enzyme
to hydrolyze longer chain substrates can be explained by the
deep continuous groove extending across the active site, while
Figure 3. (a) Superposition of A. oryzae (red) and F. solani (gray)
cutinases revealing nearly identical structural similarity. The nonover-
lapping regions are highlighted in yellow. (b) Overlay of the A. oryzae
that of F. solani favors short chain substrates due to the shallow
and interrupted groove. This is further confirmed by our models
demonstrating that the gatekeeper residues that line the groove
within the A. oryzae are farther apart than the equivalent residues
within the F. solani structure (Figure 4). In the A. oryzae
structure, modeled with the hexanoate ester (Figure 4a), this
additional space allows the carbon chains greater conformational
freedom and easier access to a large region of the groove
cutinase (colored) residues of the catalytic triad and oxyanion hole with
that of F. solani residues (gray). Residues are labeled accordingly.
Electrostatic surface rendering of A. oryzae (c) and F. solani (d) cutinases
as rendered by ICM software package.28 The green solid density
generated by the PocketFinder function of ICM illustrates the groove
on the surface proximal to the active site.28 Note that the green density
cannot fit in the narrow groove on the F. solani cutinase.
9
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A R T I C L E S
Liu et al.
Figure 4. Predicted low energy conformations of serine hexanoate esters for (a) A. oryzae and (b) F. solani cutinases. Ester heavy atoms are shown as sticks
for all conformations within 5 kcal mol-1 of the lowest energy conformation after systematic screening of torsions and minimization. The gatekeeper residues,
Leu87 and Val190 in A. oryzae and Leu81 and Val184 in F. solani, are shown as spheres.
surrounding the active site. However, in the F. solani structure
(Figure 4b), the close distance between the two gatekeeper
residues creates a barrier, constraining the alkyl chains to a
narrower region of the groove pointing away from the active
site. The presence of a wider, continuous groove by the opening
of the active site in A. oryzae cutinase can also explain its ability
to rapidly hydrolyze PCL relative to that of F. solani (Figures
3c and 4a).
Enhanced thermotolerance in comparison to that of F. solani
is observed for A. oryzae cutinase by thermoactivity and
thermodynamic stability experiments. This improved stability
can be attributed to the presence of an additional disulfide bond
as observed in the A. oryzea structure (Figure 2b). Specifically,
the disulfide bond between Cys63 and Cys76 links a peripheral
helix to the central ꢀ-sheet, providing extra stability. In general,
covalent bonds stabilize proteins; specifically disulfide bonds
connecting disparate and adjacent regions of a protein improves
its thermostability.34-36
Understanding how the cutinase structure plays an important
role in function and stability not only provides insight into the
reactivity of A. oryzae enzyme but also offers useful guidelines
for the design of proteins in general. Much of the research on
cutinases has thus far focused on that of F. solani. This is largely
attributable to the existence of a crystal structure and extensive
biochemical studies.16,18,37-40 Recently, the structure of Glom-
erella cingulata cutinase has been determined, which suggests
that the catalytic triad undergoes a significant conformational
rearrangement during the catalytic cycle,41 providing insight into
its reactivity. Although a large number of other cutinases have
been functionally investigated,18,37-40,42-45 none have been
structurally characterized and linked to their reactivities.
Cutinases have demonstrated unique properties that can be
exploited for degradation of various synthetic polymers and may
in the near future find use in plastic recycling.46 Critical features
highlighted in this work include (i) engineering a continuous
groove across the active site and (ii) including appropriate
disulfide bonds as structural features that can be used in the
redesign of other cutinases and related enzymes. From our
studies, we have obtained useful guidelines for the redesign of
proteins for environmentally compatible biocatalytic reactions
on small molecules and polymer substrates useful for industry
and academia.1-3 Further detailed characterization to elucidate
structural and functional information are underway.
Acknowledgment. This work was supported in part by NYU-
Poly Seed Fund (XPK, YG, RG and JKM), AFOSR DURIP (FA-
9550-08-1-0266) (JKM), IUCRC (RG), NSF GK-12 Fellows grant
0741714 (PJB), NIH grant GM70841 (HL and XPK) and the HHMI
Science education project at CCNY (YG, ZL and GA). We thank
Jeremy Minshull and Jonathan Ness from DNA 2.0 for their
assistance with DNA and protein expression and purification
optimization.
Supporting Information Available: The coordinates of the
A. oryzae cutinase structure are available in the protein data
bank with the ID 3GBS. SDS PAGE, CD, DSC and modeling
characterization as well as kinetics is available free of charge
via the Internet at http://pubs.acs.org.
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