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Discovery of Polyesterases from Moss-
Associated Microorganisms
Christina Andrea Müller,
a,b
Veronika Perz,
c
Christoph Provasnek,
a,b
Felice Quartinello,
d
Georg M. Guebitz,
c,d
Gabriele Berg
a
Institute of Environmental Biotechnology, Graz University of Technology, Graz, Austria
a
; ACIB GmbH, Graz,
Austria
b
; ACIB GmbH, Tulln an der Donau, Austria
c
; Institute of Environmental Biotechnology, University of
Natural Resources and Life Sciences, Tulln an der Donau, Austria
d
ABSTRACT The growing pollution of the environment with plastic debris is a
global threat which urgently requires biotechnological solutions. Enzymatic recy-
cling not only prevents pollution but also would allow recovery of valuable
building blocks. Therefore, we explored the existence of microbial polyesterases
in microbial communities associated with the Sphagnum magellanicum moss, a
key species within unexploited bog ecosystems. This resulted in the identifica-
tion of six novel esterases, which were isolated, cloned, and heterologously ex-
pressed in Escherichia coli. The esterases were found to hydrolyze the copolyes-
ter poly(butylene adipate-co-butylene terephthalate) (PBAT) and the oligomeric
model substrate bis[4-(benzoyloxy)butyl] terephthalate (BaBTaBBa). Two promis-
ing polyesterase candidates, EstB3 and EstC7, which clustered in family VIII of
bacterial lipolytic enzymes, were purified and characterized using the soluble es-
terase substrate p-nitrophenyl butyrate (K
m
values of 46.5 and 3.4
M, tempera-
ture optima of 48°C and 50°C, and pH optima of 7.0 and 8.5, respectively). In
particular, EstC7 showed outstanding activity and a strong preference for hydro-
lysis of the aromatic ester bond in PBAT. Our study highlights the potential of
plant-associated microbiomes from extreme natural ecosystems as a source for
novel hydrolytic enzymes hydrolyzing polymeric compounds.
IMPORTANCE In this study, we describe the discovery and analysis of new en-
zymes from microbial communities associated with plants (moss). The recovered
enzymes show the ability to hydrolyze not only common esterase substrates but
also the synthetic polyester poly(butylene adipate-co-butylene terephthalate),
which is a common material employed in biodegradable plastics. The widespread
use of such synthetic polyesters in industry and society requires the develop-
ment of new sustainable technological solutions for their recycling. The discov-
ered enzymes have the potential to be used as catalysts for selective recovery of
valuable building blocks from this material.
KEYWORDS enzyme discovery, esterases, functional metagenomics, plant-associated
microbiomes, polymer hydrolysis
Different types of natural aliphatic polyesters can be found in nature, such as
poly-3-hydroxybutyrate and other polyhydroxyalkanoates produced by bacteria
(1) and cutin, a main component of the cuticle in higher plants (2). In addition to these
natural polyesters, several types of synthetic polyesters, mainly aliphatic polyesters and
aliphatic-aromatic copolyesters, have been developed in the past century (3). Some
synthetic polyesters have characteristics and chemical structures that allow good
biodegradability; however, the most popular and widespread polyesters, namely, poly-
ethylene terephthalate (PET) and polybutylene terephthalate (PBT), exhibit rather re-
stricted biodegradability and were long considered to be resistant to microbial attack
Received 22 September 2016 Accepted 7
December 2016
Accepted manuscript posted online 9
December 2016
Citation Müller CA, Perz V, Provasnek C,
Quartinello F, Guebitz GM, Berg G. 2017.
Discovery of polyesterases from moss-
associated microorganisms. Appl Environ
Microbiol 83:e02641-16. https://doi.org/
10.1128/AEM.02641-16.
Editor Haruyuki Atomi, Kyoto University
Copyright © 2017 American Society for
Microbiology. All Rights Reserved.
Address correspondence to Christina Andrea
Müller, christina.mueller@tugraz.at.
C.A.M. and V.P. are co-first authors.
BIOTECHNOLOGY
crossm
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 1Applied and Environmental Microbiology
(4). The increasing pollution of nature with microplastic debris is a global threat and a
key challenge for future generations (5, 6). Biodegradation of polyesters is a heteroge-
neous process consisting of initial depolymerization to water-soluble intermediates by
extracellular enzymes and subsequent assimilation of intermediates into metabolic
pathways to achieve final degradation (4). The initial depolymerization is considered
the limiting biodegradation step (4). Systematic studies by Marten et al. revealed that
the mobility of the polymer chains is the key factor affecting the biodegradability of
aliphatic and aromatic polyesters (7, 8). By combining aliphatic and aromatic monomers
in the synthesis of copolyesters, it is possible to achieve greater biodegradability while
keeping the material properties acceptable (8). The copolyester poly(butylene adipate-
co-butylene terephthalate) (PBAT), for instance, belongs to this category. PBAT is
synthetized on an industrial scale by several companies, through polycondensation of
the aliphatic monomers adipic acid (Ada) and 1,4-butanediol (B) and the aromatic
monomer terephthalic acid (Ta) (9). PBAT can also be used as a component of polymer
blends with, for example, poly(lactic acid) (PLA) (10).
The ever-growing use of plastic-based materials strengthens the need to investigate
not only biodegradable polyesters but also microorganisms and whole microbiomes for
new enzymes. Novel biocatalysts capable of hydrolyzing polyesters could allow recy-
cling of valuable building blocks. Microbial enzymes from various sources (e.g., com-
post) that can degrade or modify natural and synthetic polyesters have been studied
previously (11, 12). Esterases and lipases not only are prevalent in most natural
ecosystems and microorganisms but also are some of the most widely used biocatalysts
in the chemical industry (13). According to the family classification system defined first
by Arpigny and Jaeger (14), bacterial esterases and lipases can be grouped into lipolytic
enzyme families I to VIII. This classification system was later extended up to family XIV
(15). Several hydrolases with polyester-degrading capabilities have been isolated from
bacteria belonging to the phyla Actinobacteria (e.g., Thermobifida spp. and Saccha-
romonospora viridis), Proteobacteria (Pseudomonas mendocina), and Firmicutes (e.g.,
Bacillus subtilis and Clostridium spp.) and from fungi (e.g., Fusarium solani,Humicola
insolens,Aspergillus oryzae, and Rhizopus spp.) (6, 11, 16–21). While searching for new
microorganisms, Yoshida et al. recently discovered the novel bacterium Ideonella
sakaiensis, which contains two PET hydrolases and is capable of utilizing PET as an
energy and carbon source (22). It was demonstrated that metagenomes are also
promising sources for novel enzymes (23). However, there are only seven examples of
polyester-hydrolyzing enzymes from metagenomic sources, including the LC-cutinase
from a leaf-branch compost metagenomic library, which is capable of hydrolyzing not
only cutin but also PET (24), the PLA depolymerase PlaM4, which is also from a compost
metagenome (25), and five esterases from marine metagenomic libraries with hydro-
lytic activities toward several polyester substrates, including PLA (26). The potential of
plant-associated metagenomes as a source of novel enzymatic activities has been less
well explored (27).
The objective of our study was to explore the Sphagnum bog metagenome for
identification of novel enzymatic activities. Sphagnum magellanicum is the dominant
species in bog ecosystems. It has been shown that the Sphagnum microbiome supports
the host in terms of metabolism, growth, and health (28), as well as ecosystem
functioning under extreme conditions (29–31). In contrast to the reported low enzy-
matic activities in the anaerobic areas of bogs (32, 33), analysis of the Sphagnum moss
metagenome revealed very high levels of taxonomic and functional diversity (31). The
influence of specialized microorganisms, harboring novel enzymes, on ecosystem
functioning was also reported for peatland soils (34). The cell wall components of
Sphagnum leaves contain polysaccharides similar to those found in higher plants,
possibly cellulose, mannans, xylans (35), pectin-like polysaccharides, xyloglucans, and
uronic acid (36). Moreover, extracts of different moss species, including Sphagnum
magellanicum, also contain high concentrations of secondary metabolites such as
sterols, polyphenols, and terpenoids (37). Based on the tight interactions and functional
diversity of the Sphagnum moss microbiome, we expected significant abundance and
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 2
high levels of diversity of hydrolases that play essential roles in plant metabolism,
especially in regard to degradation and/or assimilation of plant polysaccharides, poly-
mers, and secondary metabolites, while exhibiting hydrolytic activities toward synthetic
polymers.
RESULTS AND DISCUSSION
Screening for esterases in the Sphagnum moss metagenome. In order to
identify enzymes for hydrolysis of aromatic-aliphatic polyesters, we investigated the
microbiome associated with the moss Sphagnum magellanicum as a source for
esterases. Our screening strategy consisted of three steps (Fig. 1), allowing identi-
fication of clones displaying not only esterase activity but also, in particular,
polyester-hydrolyzing activity.
The initial high-throughput screening of around 90,000 fosmid clones from the moss
metagenome library resulted in identification of 83 clones showing hydrolytic activity
(halo formation) on tributyrin agar plates (Fig. 1A and data not shown). During the
second screening step, the 83 identified clones were evaluated for hydrolytic activity
toward p-nitrophenyl butyrate (pNPB) by using cell-free lysates (see Table S2 in the
supplemental material). The 11 most active clones were selected for a third screening
step using two different synthetic copolyesters. In addition, cell-free lysates were tested
for the presence of hydrolases by labeling active serine hydrolases in the samples with
a fluorogenic probe prior to SDS-PAGE analysis (Fig. S1); this allowed detection of
distinct and significantly intense hydrolase bands in the analyzed clones with molecular
masses of 30 to 50 kDa, providing evidence for the presence of putative esterases.
Finally, six nonredundant clones (clones B3, B11, C5, C7, C9, and G4) showed significant
hydrolytic activity with the aliphatic-aromatic copolyester PBAT (ecoflex; BASF) and the
oligomeric low-molecular-weight PBAT model substrate bis[4-(benzoyloxy)butyl] te-
rephthalate (BaBTaBBa), compared to the control reaction (library host carrying the
empty vector) (Fig. S2).
Identification and classification of metagenome esterases. Subcloning of the 6
fosmid clones with polyester-hydrolyzing activities was performed to gain shorter DNA
fragments for sequencing (Fig. 1B). The sequencing results allowed identification of
open reading frames (ORFs) coding for putative esterases. The esterases coded by the
six individual ORFs were termed EstB3, EstB11, EstC5, EstC7, EstC9, and EstG4. Table 1
shows characteristic properties of the six esterase coding sequences.
Gene lengths ranged between 888 and 1,227 bp, showing rather high GC contents
of 62.2 to 65.1%, with the exception of EstG4 (40.3%). Possible N-terminal signal
peptides were predicted for the ORFs of EstC5 and EstG4 using SignalP 4.1 software
(38). Sequence similarities among the six esterases were estimated by alignment and
FIG 1 Schematic presentation of the screening process for metagenome-derived clones showing polyester-hydrolyzing
activity and retrieval of sequences. (A) Three screening steps for identification of fosmid clones containing the target
hydrolytic activity, as follows: first step, tributyrin agar plates; second step, pNPB assay in 96-well plates; third step, HPLC
analysis of PBAT and BaBTaBBa reaction products. Positively identified clones harbor the library vector pCC2FOS containing
the metagenome insert DNA with the targeted hydrolase gene (marked with a star). (B) Subcloning of shorter insert DNA
fragments after digestion with restriction enzymes (EcoRI and HindIII or BamHI). Subclones displaying the same hydrolytic
activity were sequenced for identification of the hydrolase ORF.
Polyesterases from Moss Metagenome Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 3
pairwise comparison of amino acid sequences. EstB11 and EstC7 had the greatest
sequence identity (69.8%), followed by lower levels of identity between EstB3 and EstC7
(29.9%), EstB3 and EstB11 (27.5%), and EstC5 and EstC9 (24.7%). All other sequence
pairs demonstrated levels of identity below 14.4%.
Phylogenetic analysis of the ORF sequences revealed amino acid sequence homol-
ogy with annotated esterases from the GenBank protein database of 37 to 63% (Table
1). Multiple sequence alignments of the six metagenome esterases and reference
sequences belonging to all 14 known lipolytic families revealed the presence of
conserved esterase motifs (Fig. S3). Based on phylogenetic analysis, the metagenome
esterases could be classified into three previously reported families (see the phyloge-
netic tree in Fig. S4).
EstB3, EstB11, and EstC7 clustered into family VIII of carboxylesterases, which share
sequence identity with class C

-lactamases and penicillin-binding proteins (14). They
harbor an active site serine residue in the consensus sequence SMTK (SXXK motif), with
a catalytic triad of Ser-Lys-Tyr, comparable to the other enzymes in this family and
especially the recently reported crystal structure of EstU1 (PDB accession no. 4IVK)(39).
Promiscuous

-lactamase activity has been demonstrated for some members of this
family, such as the metagenome-derived carboxylesterases EstU1 (40) and Est22 (41),
with first-generation cephalosporin-based derivatives. EstG4 clustered in family II of
lipolytic enzymes, which show the conserved motif GDSL containing the active site
serine. In this family, the catalytic triad is the classic Ser-Asp-His sequence found in
␣
/

-hydrolases (42). EstC5 and EstC9 showed similarities to enzymes grouped into
family IV, which contains hormone-sensitive lipase (HSL)-like esterases that display the
conserved pentapeptide motif GXSXG, containing the active site serine (43). EstC5
shows the typical triad Ser-Asp-His, while EstC9 may utilize glutamic acid (Ser-Glu-His)
instead of aspartic acid. To compare the affiliations of the metagenome esterases with
those of previously characterized polyesterases with documented activity toward
PBAT, the following enzymes were included in the phylogenetic analysis: esterases
Cbotu_EstA, Cbotu_EstB (Clostridium botulinum)(
20), and Chath_Est1 (Clostridium
hathewayi)(
21), lipase Pfl1 (Pelosinus fermentans)(44), and cutinases Cut190 (Saccha-
romonosporas viridis)(
19) and Thc_Cut1 (Thermobifida cellulosilytica)(45). These six
known PBAT hydrolases share the same catalytic triad Ser-Asp-His (or Ser-Glu-His, in the
case of Chath_Est1). Cbotu_EstA, Cbotu_EstB (conserved motif GHSMGG), and Pfl1
(AHSMG) do not directly cluster with any of the 14 reported lipolytic enzyme families
and seem to represent a new cluster of lipolytic enzymes in close proximity to families
I, V, and IX. In contrast, the esterase Chath_Est1 (GQSGG) clusters in family VII and the
TABLE 1 Properties of identified ORFs and protein coding sequences (wild type) for six novel metagenome esterases
ORF
Length
(bp)
GC content
(%)
Protein sequence
Size
(amino
acids)
Estimated
molecular
mass (kDa)
Predicted
isoelectric
point
Predicted
signal peptide
a
Maximal amino acid sequence identity
(GenBank accession no.)
b
EstB3 1,227 64.9 408 44.54 6.28 No 63% vs serine hydrolase from Tardiphaga sp.
Vaf07 (WP_068735481)
EstB11 1,152 62.2 383 40.31 5.33 No 55% vs 1,4-butanediol diacrylate esterase
from Thermorudis peleae (WP_038038941)
EstC5 918 63.9 305 32.76 6.50 N-terminal (⬃34
amino acids)
61% vs esterase/lipase from Caulobacter
vibrioides (WP_004617716)
EstC7 1,155 63.6 383 40.20 5.43 No 53% vs 1,4-butanediol diacrylate esterase
from Thermorudis peleae (WP_038038941)
EstC9 888 65.1 295 30.82 7.78 No 56% vs
␣
/

-hydrolase from Acidocella sp.
MX-AZ02 (WP_008494015)
EstG4 960 40.3 319 35.80 5.61 N-terminal (⬃19
amino acids)
37% vs hypothetical protein
ACD_45C00716G0002 from uncultured
bacterium (EKD72285)
a
Signal peptide prediction was performed with SignalP 4.1, with a cutoff value for discrimination scores of 0.500.
b
BLASTP analysis against the GenBank database of full-length nonredundant protein sequences (23 August 2016 version) yielded the maximal identity to the closest
annotated hit.
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 4
cutinases Cut190 and Thc_Cut1 (GXSMG) in family III. The phylogenetic analysis indi-
cated that the previously reported PBAT-degrading enzymes are not affiliated with the
same lipolytic families as the esterases discovered in this study. Therefore, the observed
polyesterase activities do not seem to be strictly dependent on sequence similarity.
Substrate profiles and thermostability of cloned esterases. The six metagenome
esterase genes were cloned into E. coli BL21(DE3) or Lemo21(DE3) using the inducible
vector pET28a(⫹). Heterologous expression of soluble enzymes was confirmed by
SDS-PAGE analysis and visualization using a fluorogenic serine hydrolase label (Fig. S5).
Substrate specificity profiles of the cloned esterases were measured using cell-free
lysates and p-nitrophenyl (pNP) aliphatic esters of different chain lengths (Fig. 2A). To
enable comparison of the substrate profiles despite strong differences in the expression
levels of the esterases in the cell-free lysates, volumetric activities were calculated as
percentages of the activity measured with the commonly employed reference substrate
pNPB. In the cases of EstB3 and EstC7, substrate profiles were also recorded with
purified enzyme (Fig. 2B).
All six esterases showed strong preferences for substrates with short (C
2
to C
4
)or
medium (C
6
to C
10
) chain lengths. The smallest substrate (pNP acetate [C
2
]) was
accepted by all esterases, while the largest tested substrate (pNP palmitate [C
16
]) was
converted only by EstB11, EstC5, EstC7, and EstG4. The esterases EstB3, EstC5, EstC7,
and EstG4 showed their highest hydrolysis rates with pNP hexanoate (C
6
), while EstB11
was more active with pNP decanoate (C
10
) and EstC9 preferentially hydrolyzed pNPB
(C
4
). The substrate profiles were corroborated for EstB3 and EStC7 using purified
enzymes. Based on the substrate profiles, it can be concluded that the identified
enzymes displayed esterase character, belonging to the group of true esterases. The six
moss metagenome esterases presented here showed clear differences in their substrate
profiles. The majority of metagenome-derived esterases discovered to date also show
preferences for short-chain to medium-chain p-nitrophenyl aliphatic esters (46).
The thermostability of the esterases in the cell-free lysates was evaluated for a
period of at least 96 h. The stability was measured as residual activity at 30°C and 50°C
using the pNPB assay (Fig. S6). During incubation for 96 h at 30°C, no significant loss of
hydrolytic activity was observed for the six esterases. At 50°C, the esterases in lysates
B3, B11, C5, C7, and C9 were quickly inactivated. In contrast to the other esterases,
esterase G4 showed a high level of stability at 50°C during a prolonged incubation of
96 h, without a decrease in residual activity. Control samples (cell-free lysates of E. coli
carrying the empty vector) did not show any significant activity. Due to the excellent
stability of all six esterases at 30°C for 72 h, the hydrolysis experiments with PBAT were
performed at this temperature.
Activity with the polyester PBAT. In addition to the activities determined with the
common soluble ester substrates, the hydrolysis of the polyester PBAT and the more
FIG 2 Substrate profiles of metagenome esterases, as either cell-free lysates (A) or purified enzymes (B),
using pNP aliphatic esters with various chain lengths. The pNP aliphatic esters were pNP acetate (C
2
),
pNPB (C
4
), pNP hexanoate (C
6
), pNP octanoate (C
8
), pNP decanoate (C
10
), pNP dodecanoate (C
12
), and pNP
palmitate (C
16
). Bars, mean values of at least 2 measurements; error bars, standard deviations.
Polyesterases from Moss Metagenome Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 5
persistent polyester PET was investigated. Our goal was to evaluate the potential of
each metagenome esterase for polyester degradation and to select the best degraders
for further study. The polyester cleavage pattern depends on each enzyme’s specificity
and its ability to accommodate the less-flexible, high-molecular-weight aliphatic-
aromatic copolyester chain in the enzyme’s active site (8). Enzymatic hydrolysis of PBAT
(Fig. 3) leads to the release of the possible hydrolytic products Ada, bis(4-hydroxybutyl)
terephthalate (BTaB), and B, resulting from hydrolysis of the aliphatic ester bonds, as
well as mono(4-hydroxybutyl) terephthalate (BTa) and Ta, resulting from hydrolysis of
the aromatic ester bonds.
The polyester was applied to the reactions in two different forms, i.e., either as
milled powder (Fig. 4A) or as a piece of polyester foil (Fig. 4B). Product quantification
by means of reverse-phase high-performance liquid chromatography (RP-HPLC) with
UV-visible (UV-Vis) detection focused on detection of the most important products from
hydrolytic cleavage of the aromatic ester bond, namely, Ta and the Ta-containing
oligomers BTa and BTaB.
Generally, higher concentrations of Ta versus BTaB and BTa indicate more efficient
enzymatic hydrolysis of the aromatic ester bond, i.e., the Ta-B bond (12). The highest
concentrations of released Ta (up to 35
M) were measured with milled PBAT and
EstB3, EstB11, and EstC7 (Fig. 4A). In contrast, EstC5 and EstC9 showed minor accumu-
lation of BTa and BTaB with both substrate forms but poor or no release of Ta.
Moreover, the hydrolytic activity of EstG4 toward PBAT was negligible in both cases. To
FIG 3 Chemical structure (A) and enzymatic hydrolysis (B) of the aliphatic-aromatic copolyester PBAT.
PBAT contains an adipic acid/terephthalic acid ratio of approximately 50:50. Possible hydrolysis products
are BTaB, BTa, B, Ada, and Ta.
FIG 4 Hydrolysis of PBAT by cell-free lysates from strains containing heterologous expressed esterases
from the Sphagnum moss metagenome. Reactions with milled PBAT (A) and PBAT foil (B) were performed
as described in Materials and Methods (2-ml reaction mixtures at pH 7.5, with incubation at 30°C and 100
rpm for 72 h). The hydrolytic products Ta, BTa, and BTaB were quantified by HPLC/UV-Vis analysis. Values
for hydrolysis products released during control and blank reactions were subtracted in order to exclude
the influence of hydrolytic enzymes from the E. coli host. Bars, mean values (n⫽3); error bars, standard
deviations.
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 6
exclude the influence of hydrolytic activities of other enzymes present in the cell-free
lysates, values for control and blank reactions were subtracted from those for the
measured samples; however, it was not possible to exclude synergistic effects of the
heterologous expressed esterases and the native enzymatic activity of E. coli during
hydrolysis of the polyester.
Esterases displaying higher concentrations of released Ta are of special interest,
since the enzymatic cleavage of aromatic ester bonds in a polyester chain is more
difficult to achieve than the cleavage of aliphatic ester bonds, due to lower chain
mobility and melting temperatures (3, 7, 8). Released Ta concentrations were nearly
4-fold higher after incubation of the same enzymes with PBAT applied as milled
particles, in contrast to PBAT foil (Fig. 4B). The higher activity levels with milled and
resuspended PBAT particles is not surprising, since the particles offer greater surface
area for enzyme attachment and consequently greater amounts of exposed substrate
chains for catalytic attack than does the PBAT foil. Unfortunately, no significant
amounts of hydrolysis products could be measured for the reaction mixtures contain-
ing PET foil. Interestingly, the three esterases (EstB3, EstB11, and EstC7) that exhibited
high levels of activity with PBAT belong to the same family of lipolytic enzymes, i.e.,
family VIII, which has a different catalytic triad (Ser-Lys-Tyr), compared to the classic
Ser-Asp/Glu-His triad found in the previously characterized PBAT hydrolases Thc_Cut1,
Cbotu_A, Cbotu_B, Pfl1, Cut190, and Chath_Est1 (19–21, 44, 45). In summary, the
esterases EstB3, EstB11, and EstC7 showed higher levels of activity in polyester hydro-
lysis than did EstC5, EstC9, and EstG4 and therefore are the most promising candidates
for further study.
Kinetic characterization of purified EstB3 and EstC7. The esterases EstB3, EstB11,
and EstC7 showed significant hydrolytic activity with PBAT during prescreening exper-
iments. For that reason, these enzymes were selected for purification by affinity
chromatography with Ni-Sepharose columns, using the N-terminal His-tagged proteins.
Purification of EstB3 and EstC7 was achieved with purities of ⬎98%, as estimated by
SDS-PAGE analysis (Fig. S7). Unfortunately, His tag purification of EstB11 was not
successful under several conditions. Therefore, this study focused on characterization of
purified EstB3 and EstC7, in terms of kinetic parameters and hydrolysis of the PBAT
polyester.
The kinetic data for EstC7 were fitted according to the Michaelis-Menten equation,
resulting in a V
max
of 70.6 U mg
⫺1
and a K
m
of 3.43
M. The best fitting of the EstB3
kinetic parameters was achieved by using the Hill equation instead of the Michaelis-
Menten equation, and n
H
was estimated to be 0.687. V
max
and K
m
values for EstB3 were
340.8 U mg
⫺1
and 46.45
M, respectively. The K
m
values determined for EstB3 and
EstC7 were very low, suggesting high affinity for pNPB as a substrate. For comparison,
K
m
values for other family VIII carboxylesterases from metagenome sources, such as
those for EstC (58.7
M) (47) and EstU1 (6.0
M) (40), are in a similar range. The recently
described esterase Cbotu_EstA, which is also capable of hydrolyzing PBAT, has a V
max
value similar to that of EstC7 (Cbotu_EstA V
max
⫽83.4 U mg
⫺1
); however, the K
m
value
for Cbotu_EstA is 3 orders of magnitude higher (1.95 mM) (20). Lower K
m
values for
pNPB were reported for the eukaryotic liver carboxylesterases from sheep (Ovis aries)
(0.43
M), chicken (Gallus gallus) (0.55
M), and horse (Equus caballus) (1.1
M) (48).
The temperature and pH profiles of EstB3 and EstC7 were determined as well (Fig.
5). For the temperature profiles, the standard pNPB assay was used. Determination of
pH profiles with the chromogenic substrate pNPB was not feasible at pH values higher
than 9.0, due to rapid base-catalyzed autohydrolysis of the substrate. The fluorogenic
substrate 4-methylumbelliferyl butyrate (4-MUB) showed less susceptibility to autohy-
drolysis at pH values of ⬎9.0 and therefore was employed for the measurements.
The optimal temperatures for enzyme-catalyzed hydrolysis were determined to be
around 47.7°C for EstB3 and 50°C for EstC7 (Fig. 5A). EstB3 showed a broader optimal
pH range between 6.0 and 8.0, with a maximum at pH 7.0, whereas EstC7 had a narrow
optimal pH range, with a maximum at pH 8.5 (Fig. 5B).
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Polyesterase activity of EstB3 and EstC7 with PBAT (milled and foil). The
polyesterase activity was measured by employing purified esterases EstB3 and EstC7
(Fig. 6). Hydrolytic activities were normalized to the applied esterase concentration (0.6
M).
The polyesterase activities of purified EstB3 and EstC7 with both milled PBAT (Fig.
6A) and PBAT foil (Fig. 6B) showed distinct hydrolysis patterns. After incubation with
both enzymes and substrate forms, Ta and BTa were determined to be the main
hydrolysis products. No BTaB was observed; however, accumulation of BTa was prom-
inent for EstB3 with milled PBAT, with a 6-fold higher concentration of BTa (60.4
M
[101 mol mol of enzyme
⫺1
]) versus Ta (8.0
M [17 mol mol of enzyme
⫺1
]). The total
concentration of quantified products in the EstB3 reaction was higher with the milled
substrate than with the substrate foil. In contrast, the total concentrations of quantified
monomeric subunits for EstC7 were equal with the two substrate forms (318
M [530
mol mole enzyme
⫺1
]). Total concentrations of quantified products were higher in the
reactions with purified esterases than in the reactions with cell-free lysates (Fig. 4),
probably due to greater catalyst loads. An unexpected finding was the change in the
product profiles for the reactions with purified EstB3 and EstC7, compared to the
reactions with the respective cell-free lysates. The higher concentrations of hydrolysis
products measured with purified enzymes led to greater accumulation of BTa (Fig. 6).
A possible reason is inactivation of the esterase before BTa is hydrolyzed further to the
monomeric subunits (Ta and B). In addition, although control and blank reaction values
were subtracted in the cell-free lysate experiments, a synergistic effect between the
esterases and other cell lysate components, leading to a different distribution of
hydrolysis products and less accumulation of BTa with cell-free lysates, cannot be fully
excluded. Purified EstC7 yielded higher concentrations of Ta overall, especially with the
FIG 5 Temperature profiles (A) and pH profiles (B) of purified EstB3 and EstC7. Activities were measured
as described in Materials and Methods, using p-nitrophenyl butyrate for the temperature measurements
(at pH 7.5) and 4-methylumbelliferyl butyrate for pH measurements (at 30°C). Mean values of at least
three separate measurements are shown; error bars indicate the standard deviations. For reactions at
34.2, 47.7, and 61.5°C with EstC7 in panel A, single measurements were performed.
FIG 6 Hydrolysis of milled PBAT (A) and PBAT foil (B) by purified esterases EstB3 and EstC7. Reactions were
performed as described in Materials and Methods (2-ml reaction mixtures at pH 7.5 with 0.6
M enzyme,
with incubation at 30°C and 150 rpm for 72 h). Hydrolysis products Ta, BTa, and BTaB were quantified by
HPLC/UV-Vis analysis. The sum of products (Ta, BTa, and BTaB) is also given. Bars, mean values (n⫽3); error
bars, standard deviations.
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 8
PBAT foil (191.1
M [318.5 mol mol of enzyme
⫺1
]), indicating more efficient hydrolysis
to the monomeric subunits and more effective hydrolysis of the aromatic Ta-B ester
bond. Similar differences in hydrolytic cleavage patterns for aliphatic-aromatic polyes-
ters between enzymes were reported previously, for example, for the cutinases from
Humicola insolens (HiC) and Thermobifida cellulosilytica (Thc_Cut1). The cutinase Thc_
Cut1 shows greater specificity for the hydrolysis of aromatic ester bonds (Ta-B) than
does the cutinase HiC (12). Moreover, the polyesterase activities of the novel enzymes
EstB3 and EstC7 with milled PBAT were significantly higher than the activities of the
recently presented esterases from the anaerobic bacteria Clostridium hathewayi
(Chath_Est1) (21) and Clostridium botulinum (Cbotu_Est and Cbotu_EstB) (20).
Conclusion. The plant-associated microbiome of moss plantlets was successfully
investigated as a source of novel enzymes. To date, only a few examples reporting the
identification of enzymes in plant-associated microbiomes through functional metag-
enomics are available (27). Moreover, we demonstrate here the hydrolytic activities of
esterases obtained from a plant-associated microbiome with a synthetic copolyester.
EstB3 and EstC7 showed significant hydrolytic activities with the synthetic aliphatic-
aromatic copolyester PBAT. Since EstB3 and EstC7 belong to family VIII of carboxyles-
terases, polyesterase activity emerges as an unusual activity of this lipolytic enzyme
family. In particular, EstC7 showed a strong preference for cleavage of the aromatic
(Ta-B) ester bond, which is an attractive and challenging hydrolytic activity for degra-
dation or modification of polyester-based materials.
MATERIALS AND METHODS
Chemicals and reagents. Milled PBAT, with a melting point of 125.3°C, a glass transition tempera-
ture of ⫺34°C, a molecular number average of 19,000 g mol
⫺1
, and a molecular weight average of 65,000
g mol
⫺1
, milled to a particle size of 100 to 300
m as described elsewhere (12), and PBAT foil (ecoflex,
with a film thickness of 50
m) were kindly provided by BASF SE. BaBTaBBa, with a particle size of 100
to 300
m, BTaB, and BTa were synthesized and purified according to the methods described by Perz et
al. (49). Amorphous solvent-casted polyethylene terephthalate films (film thickness of 0.25 mm) were
purchased from Goodfellow (United Kingdom). The polymers used were of analytical grade, in order to
exclude any influences of additives. All other chemicals were of analytical grade and purchased from
Sigma-Aldrich (Germany), TCI Europe (Germany), Carl Roth (Germany), or Merck (Germany). DNA-
modifying enzymes were obtained from New England BioLabs (Germany) or Thermo Scientific (Germany)
and oligonucleotides from Sigma-Aldrich (Germany). Plasmid DNA was isolated using the GeneJet
plasmid miniprep kit (Thermo Scientific). The Wizard SV PCR cleanup kit (Promega, USA) was used for
purification and agarose gel extraction of PCR products.
Initial screening of the Sphagnum moss metagenome library. The microbiome of moss
(Sphagnum magellanicum) from a sampling site in the Alpine bog Pirker Waldhochmoor
(N46°37=38.66⬙, E14°26=5.66⬙) in Austria was employed for extraction of metagenomic DNA and
generation of a fosmid clone library using the CopyControl fosmid library production kit (Epicentre),
as described previously (50). Activity screening of the fosmid library in E. coli EPI300 pCC2FOS
(Epicentre) was performed on LB medium (pH 7.5) agar plates containing 1% (vol/vol) tributyrin
(emulsified using an Ultra Turrax homogenizer [IKA Werke, Germany]), 15 g liter
⫺1
agar, 1.2 g liter
⫺1
gum arabic, 1⫻fosmid autoinduction solution (Epicentre), and 12.5
gml
⫺1
chloramphenicol. The
plates were incubated at 30°C for up to 7 days until a clear zone (halo) was detected around the
colonies. Positive clones were evaluated for esterase activity using cell-free lysates and the soluble
esterase substrate pNPB.
Preparation of cell-free lysates. Fosmid clones (E. coli EPI300) were cultured at 30°C at 120 rpm for
16 h in 50 ml LB medium containing 1⫻fosmid autoinduction solution (Epicentre, USA) and 12.5
g
ml
⫺1
chloramphenicol. Cells were harvested by centrifugation (2,900 ⫻gat 4°C for 15 min). Cell pellets
from 50- or 400-ml cultures were resuspended in 6 or 40 ml, respectively, of 0.1 M Tris-HCl buffer (pH 7.5).
Cell-free lysates were prepared by sonication (twice for 1 min, at 45% amplitude) with a digital sonifier
(Branson, USA), followed by lysis in a FastPrep instrument (MP Biomedicals, USA) for 60 s at 6.0 m s
⫺1
,
using 20% (vol/vol) glass beads (0.1 to 0.25 mm; Retsch GmbH, Germany). Cell debris was removed by
centrifugation (16,000 ⫻gat 4°C for 30 min), and lysates were filtered (0.45
m; Roth, Germany) and
stored at 4°C until further use. For polyester degradation assays, lysates were shock-frozen in liquid
nitrogen and lyophilized for 48 h in a FreeZone 4.5 freeze dryer (Labconco, USA).
Subcloning, sequencing of fosmids, and cloning of esterases. A subclone library of each fosmid
was prepared by restriction of isolated plasmids with EcoRI and HindIII for B11, C5, C7, C9, and G4 or
BamHI for B3, followed by cleanup of restriction products, ligation into the same vector backbone used
for library construction (pCC2FOS), and transformation into the library strain E. coli EPI300 (Epicentre).
Clones were subjected again to activity screening on tributyrin agar plates, as described above. From
each subclone library, one colony displaying halo formation was selected for sequencing by primer
walking at LGC Genomics (Germany) or Microsynth (Switzerland). Primers used for the initial sequencing
are specified in Table S1 in the supplemental material. CLC Main Workbench software (Qiagen) was
Polyesterases from Moss Metagenome Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 9
employed for sequence assembly, BLASTX analysis, and identification of ORFs. The esterase-coding genes
for EstB11, EstC5, EstC7, EstC9, and EstG4 were cloned into pET28a(⫹) (Novagen, USA) through restriction
cloning, while EstB3 was cloned with a ligase-independent method (phosphorothioate-based ligase-
independent gene cloning [PLICing]), as described elsewhere (51). A standard PCR mixture (50
l)
contained 1⫻Phusion HF buffer,1UofPhusion DNA polymerase (New England BioLabs), 0.2 mM
deoxynucleoside triphosphates, 0.2
M each primer, and 10 ng of plasmid DNA. The PCR primers
employed are listed in Table S1. Ligated/hybridized plasmids were transformed into chemically compe-
tent E. coli BL21(DE3) cells (Invitrogen, USA) and E. coli Lemo21(DE3) cells (New England BioLabs,
Germany).
Protein expression. The strains employed for expression of the six cloned esterases in pET28a(⫹)
were E. coli Lemo21(DE3) for EstB3, EstB11, EstC5, EstC9, and EstG4 and E. coli BL21(DE3) for EstC7. For
control reactions, E. coli strains BL21(DE3) and Lemo(DE3) containing the empty vector pET28a(⫹) were
cultivated under the same conditions. Flasks containing 400 ml of LB medium, 50
gml
⫺1
kanamycin,
and 12.5
gml
⫺1
chloramphenicol [for Lemo21(DE3) strains] were inoculated with 2% (vol/vol) of an
overnight culture and incubated at 37°C and 120 rpm until the respective optical density at 600 nm
(OD
600
) was obtained (OD
600
of 0.6 for EstB3 and OD
600
of 0.8 for the remaining strains). Protein
expression was induced by the addition of isopropyl-

-D-thiogalactopyranoside (IPTG) (1 mM for EstB11
and EstC5 and 0.4 mM for the remaining strains). Cultures containing EstB3, EstB11, EstC5, and EstC7
were incubated further at 25°C and 200 rpm, and those containing EstC9, EstG4, and the empty vector
controls were incubated at 30°C and 200 rpm. The optimal incubation time after induction was4hfor
EstC7, EstC9, and EstG4, 5 h for EstB3, and 20 h for EstB11 and EstC5. Additionally, in the case of EstB3,
0.5 mM L-rhamnose was added upon induction and agitation was performed at 240 rpm. Cell-free lysates
for activity assays were prepared as indicated above.
Esterase activity assay with p-nitrophenyl butyrate. Hydrolytic activity was assayed spectropho-
tometrically at 410 nm and 30°C using a U-2001 spectrophotometer (Hitachi, Japan) or a Tecan M200
plate reader (Tecan, Switzerland). A typical reaction mixture contained 0.5 mM pNPB, 5% (vol/vol)
acetonitrile as cosolvent, and 50
l of cell-free lysate (65 to 250
g total protein) or 0.2 to 2.5
g purified
enzyme, in a final volume of 1 ml (cuvette) or 200
l (96-well plate) of 0.1 M Tris-HCl buffer (pH 7.5). The
reaction was initiated by the addition of enzyme or lysate. One unit was defined as the amount of activity
required for the release of 1
mol of p-nitrophenol per minute. The initial hydrolysis rates (linear signal,
up to 20 s) were employed for activity determination. Concentrations of 3.12 to 100
Mp-nitrophenol
(extinction coefficient of 10.2 mM
⫺1
cm
⫺1
) were used to generate a calibration curve.
Detection of active serine hydrolases by SDS-PAGE and fluorescence labeling. In order to
identify serine hydrolases in cell-free lysates and purified proteins, SDS-PAGE was performed according
to the method described by Laemmli (52). Protein samples were pretreated with the ActiveX
6-carboxytetramethylrhodamine fluorophosphonate (TAMRA-FP) serine hydrolase probe (Thermo Scien-
tific, USA), according to the manufacturer’s protocol, to label active serine hydrolases prior to SDS-PAGE
separation. After fluorescence imaging of the gel with a Cy3 filter to detect labeled serine hydrolases,
proteins were stained with Coomassie brilliant blue R-250. The PageRuler prestained protein ladder (10
to 170 kDa; Thermo Scientific, Germany) was used as a molecular mass marker.
Screening for polyester hydrolytic activity using PBAT, model substrate BaBTaBBa, and PET.
Cell-free lysates of nonredundant fosmid clones (clones B3, B11, C5, C7, C9, and G4) were evaluated for
their hydrolytic activities with milled PBAT and the oligomeric model substrate BaBTaBBa. Lyophilized
cell-free lysates (80 to 84 mg, from 50-ml cultures) were redissolved in 3.5 ml double-distilled water
(ddH
2
O) to obtain a 0.17 M Tris-HCl buffered solution (pH 7.5). Esterase activity of the samples was first
confirmed with the pNPB assay. Rehydrated lysates (1 ml; 23 to 24 g liter
⫺1
) were incubated for 72 h at
30°C and 100 rpm with 10 mg milled PBAT, 7 mg BaBTaBBa, or PET foil (0.5 by 1 cm), followed by
RP-HPLC/UV-Vis analysis. Prior to incubation, PET foils were washed as described previously (45).
Substrate specificity for p-nitrophenyl aliphatic esters. Esterase activity toward p-nitrophenyl
aliphatic esters with different chain lengths was determined according to the standard ester-hydrolyzing
activity assay described for pNPB, using a U-2001 spectrophotometer (Hitachi). Hydrolytic activity was
measured using the following pNP esters: pNP acetate (pNPC
2
), pNPB (pNPC
4
), pNP hexanoate (pNPC
6
),
pNP octanoate (pNPC
8
), pNP decanoate (pNPC
10
), pNP dodecanoate (pNPC
12
), and pNP palmitate
(pNPC
16
). To enable comparison of the substrate profiles using cell-free lysates, independent of different
expression levels in each lysate, volumetric activities (units per liter) were calculated as percentages of
the activity with the reference substrate pNPB. For purified enzymes (EstB3 and EstC7), the hydrolytic
activity was calculated as units per milligram of enzyme. To increase the solubility of the long-chain and
less-soluble substrates and to provide equal conditions for all reactions, 1% (vol/vol) Triton X-100 was
added to the reaction buffer. The reactions in 0.1 M Tris-HCl buffer (pH 7.5) contained 0.5 mM pNP
substrate, 5% (vol/vol) acetonitrile, 1% (vol/vol) Triton X-100, and cell-free lysate or purified enzyme. For
purified enzymes, protein concentrations were measured using the Pierce bicinchoninic acid (BCA)
protein assay kit (Thermo Scientific), with bovine serum albumin as the protein standard. Background
activity was measured in a blank reaction without the addition of lysate. Moreover, control reactions were
performed using lysates of E. coli carrying the empty vector. The sample signals were then corrected by
subtraction of the background and control reaction signals.
Thermostability (residual activity). The thermostability of the six cloned and expressed esterases
was measured using sterile filtered cell-free lysates. The lysates were incubated continuously at 30°C and
50°C for 168 h and 96 h, respectively. Samples were withdrawn upon initiation of the incubation (initial
activity), after 6 h, and every 24 h. Initial and residual activities were measured in triplicate using the pNPB
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 10
assay, as described above. Control reactions using cell-free lysates of E. coli carrying the empty vector
were performed simultaneously, and results were subtracted from the sample signals.
Purification of EstC7 and EstB3. His-tagged esterases EstC7 and EstB3 were purified under non-
denaturing conditions using a 1-ml HisTrap HP column (GE Health Care, USA), according to the
manufacturer’s protocol. After expression, cell pellets were resuspended in 14 ml of 20 mM Tris-HCl
buffer (pH 7.0) containing 0.5 M NaCl and 50 mM imidazole. Cell-free lysates were prepared as described
above. Fractions containing active esterase were eluted with 500 and 750 mM imidazole for EstC7 and
EstB3, respectively. Buffer exchange was performed through dilution and centrifugation using Amicon
Ultra-15 centrifugal filters (10-kDa cutoff; Merck Millipore), using 20 mM Tris-HCl buffer (pH 7.0)
containing 0.5 M NaCl. For the polyesterase activity determination, the purified enzymes were shock-
frozen in liquid nitrogen and lyophilized as described above.
Temperature optima, pH optima, and kinetic characterization of EstC7 and EstB3. Kinetic data
(K
m
and V
max
) were recorded at 30°C and pH 7.5, using purified enzyme (0.78 to 1,000
M) and the pNPB
assay described above. Protein concentrations were measured using the Pierce BCA protein assay kit
(Thermo Scientific), with bovine serum albumin as the protein standard. Specific activities are given as
units per milligram of enzyme. Temperature optima were also determined using the pNPB assay, at a pH
of 7.5 and various temperatures (15 to 64°C). For the pH-dependent activity measurements, the more
suitable fluorogenic substrate 4-MUB was employed, since the chromogenic substrate pNPB is more
susceptible to autohydrolysis above pH 9, making measurements at pH values of ⬎9 unfeasible. The
4-MUB assay consisted of 0.5 mM 4-MUB in 0.2 ml of the Britton-Robinson universal buffer system (40
mM acetic acid, 40 mM phosphoric acid, and 40 mM boracic acid, at pH values of 5.0 to 12.0). The reaction
was started by the addition of enzyme. The increase in fluorescence was recorded using a Tecan M200
plate reader, and quantification was performed using a 4-methylumbelliferone standard curve for each
pH value. The signal was corrected for autohydrolysis of the substrate at each pH by subtracting the
background signal (control reaction without enzyme). One unit was defined as the amount of activity
required for the release of 1
mol of 4-methylumbelliferone per minute.
Polyesterase activity of all six esterases (cell-free lysates) and purified EstB3 and EstC7 with
PBAT (milled and foil). Lyophilized cell-free lysates (0.3 g, from 200-ml cultures) from E. coli
Lemo21(DE3) or E. coli BL21(DE3) containing the expressed esterases or the empty vector (control
reaction) and Tris-HCl buffer were resuspended in 13 ml ddH
2
O, while lyophilized purified enzymes EstB3
(48.08 kDa) and EstC7 (43.74 kDa) were dissolved in ddH
2
O to yield an enzyme concentration of 0.6
M.
The reaction mixtures had a final pH of 7.5 and contained one piece of PBAT foil (0.5 by 1 cm) or 10 mg
milled PBAT in 2 ml of rehydrated cell-free lysate (23.1 g liter
⫺1
) or 2 ml of purified enzyme (0.6
M).
Samples were incubated at 30°C and 150 rpm for 72 h, followed by HPLC analysis. Blank reactions,
without the addition of resuspended cell-free lysates or purified enzymes, and control reactions,
containing resuspended cell-free lysates of E. coli carrying the empty vector, were performed simulta-
neously. The concentrations of hydrolysis products released during control and blank reactions were
subtracted from the concentrations reached in sample reactions in order to exclude the influence of
hydrolytic enzymes from the E. coli host.
HPLC quantification of released hydrolysis products. Hydrolysis reactions were stopped by
methanol precipitation through the addition of 0.5 ml of ice-cooled methanol to 0.5-ml sample, control,
and blank reaction mixtures. The hydrolysis products Ta, Ada, benzoic acid (Ba), BTa, and BTaB were
quantified by means of RP-HPLC using a Dionex system (Dionex, USA), consisting of an UltiMate 3000
pump, an ASI-100 automated sample injector, an UltiMate 3000 column compartment, and a UVD 340U
photodiode array detector, in combination with a XTerra RP
18
column (3.5
m; 3.0 mm by 150 mm) with
a precolumn (Waters Corp., USA). Separation of the hydrolysis products was performed as described
previously (12). Signals from blank and control reactions were subtracted from the sample signals in
order to correct the data for background activity (autohydrolysis). Quantification of hydrolysis products
was performed using calibration curves constructed with authentic standards.
Phylogenetic analysis. Phylogenetic analysis on the basis of amino acid sequence similarities was
performed using sequences from closest hits from BLASTX analysis with the NCBI GenBank protein
database. Multiple sequence alignments were performed using ClustalW software (53), Gonnet protein
weight matrices, and default parameters, with visualization by ESPript 3.0 (54). A neighbor-joining
phylogenetic tree was constructed with MEGA6 software (55), using the number-of-differences method
and a bootstrap of 1,000 replicates. Positions containing gaps and missing data were eliminated, which
yielded a total of 92 positions in the final data set.
Accession number(s). The nucleotide sequences were deposited in the GenBank database under the
following accession numbers: KX533504 for EstB3, KX533505 for EstB11, KX533506 for EstC5, KX533507
for EstC7, KX533508 for EstC9, and KX533509 for EstG4.
SUPPLEMENTAL MATERIAL
Supplemental material for this article may be found at https://doi.org/10.1128/
AEM.02641-16.
TEXT S1, PDF file, 3.1 MB.
ACKNOWLEDGMENTS
This work was supported by the Federal Ministry of Science, Research, and Economy,
the Federal Ministry of Traffic, Innovation, and Technology, the Styrian Business Pro-
Polyesterases from Moss Metagenome Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 11
motion Agency, the Standortagentur Tirol, the Government of Lower Austria, and the
Technology Agency of the City of Vienna through the COMET Funding Program,
managed by the Austrian Research Promotion Agency.
We thank Cornelia Rainer, Tanja Nottendorfer, and Christina Laireiter (Graz Univer-
sity of Technology), as well as Johanna Aigner (FH Joanneum Graz), for valuable support
and Daria Rybakova (Graz University of Technology) for proofreading of the manuscript
and valuable discussions.
REFERENCES
1. Lee SY. 1996. Bacterial polyhydroxyalkanoates. Biotechnol Bioeng 49:
1–14. https://doi.org/10.1002/(SICI)1097-0290(19960105)49:1⬍1::AID
-BIT1⬎3.3.CO;2-1.
2. Pollard M, Beisson F, Li Y, Ohlrogge JB. 2008. Building lipid barriers:
biosynthesis of cutin and suberin. Trends Plant Sci 13:236–246. https://
doi.org/10.1016/j.tplants.2008.03.003.
3. Witt U, Müller R-J, Deckwer W-D. 1995. New biodegradable polyester-
copolymers from commodity chemicals with favorable use properties. J
Environ Polym Degrad 3:215–223. https://doi.org/10.1007/BF02068676.
4. Mueller R-J. 2006. Biological degradation of synthetic polyesters: en-
zymes as potential catalysts for polyester recycling. Process Biochem
41:2124–2128. https://doi.org/10.1016/j.procbio.2006.05.018.
5. Woodall LC, Sanchez-Vidal A, Canals M, Paterson GLJ, Coppock R, Sleight
V, Calafat A, Rogers AD, Narayanaswamy BE, Thompson RC. 2014. The
deep sea is a major sink for microplastic debris. R Soc Open Sci 1:140317.
https://doi.org/10.1098/rsos.140317.
6. Zimmermann W, Billig S. 2010. Enzymes for the biofunctionalization of
poly(ethylene terephthalate), p 97–120. In Nyanhongo GS, Steiner W,
Gübitz G (ed), Biofunctionalization of polymers and their applications.
Springer, Berlin, Germany.
7. Marten E, Müller R-J, Deckwer W-D. 2003. Studies on the enzymatic
hydrolysis of polyesters. I. Low molecular mass model esters and ali-
phatic polyesters. Polym Degrad Stab 80:485–501.
8. Marten E, Müller R-J, Deckwer W-D. 2005. Studies on the enzymatic
hydrolysis of polyesters. II. Aliphatic-aromatic copolyesters. Polym De-
grad Stab 88:371–381.
9. Vroman I, Tighzert L. 2009. Biodegradable polymers. Materials
2:307–344. https://doi.org/10.3390/ma2020307.
10. Weng Y-X, Jin Y-J, Meng Q-Y, Wang L, Zhang M, Wang Y-Z. 2013. Biodeg-
radation behavior of poly(butylene adipate-co-terephthalate) (PBAT), poly-
(lactic acid) (PLA), and their blend under soil conditions. Polym Test 32:
918–926. https://doi.org/10.1016/j.polymertesting.2013.05.001.
11. Kleeberg I, Hetz C, Kroppenstedt RM, Müller R-J, Deckwer W-D. 1998.
Biodegradation of aliphatic-aromatic copolyesters by Thermonospora
fusca and other thermophilic compost isolates. Appl Environ Microbiol
64:1731–1735.
12. Perz V, Bleymaier K, Sinkel C, Kueper U, Bonnekessel M, Ribitsch D,
Guebitz GM. 2016. Substrate specificities of cutinases on aliphatic-
aromatic polyesters and on their model substrates. New Biotechnol
33:295–304. https://doi.org/10.1016/j.nbt.2015.11.004.
13. Wohlgemuth R. 2010. Large-scale applications of hydrolases in biocata-
lytic asymmetric synthesis, p 249–264. In Blaser HU, Federsel H-J (ed),
Asymmetric catalysis on industrial scale: challenges, approaches and
solutions, 2nd ed. Wiley-VCH, Weinheim, Germany.
14. Arpigny JL, Jaeger K-E. 1999. Bacterial lipolytic enzymes: classification
and properties. Biochem J 343:177–183. https://doi.org/10.1042/
bj3430177.
15. Rao L, Xue Y, Zhou C, Tao J, Li G, Lu JR, Ma Y. 2011. A thermostable
esterase from Thermoanaerobacter tengcongensis opening up a new
family of bacterial lipolytic enzymes. Biochim Biophys Acta 1814:
1695–1702. https://doi.org/10.1016/j.bbapap.2011.08.013.
16. Tokiwa Y, Suzuki T. 1977. Hydrolysis of polyesters by lipases. Nature
270:76–78. https://doi.org/10.1038/270076a0.
17. Ronkvist ÅM, Xie W, Lu W, Gross RA. 2009. Cutinase-catalyzed hydrolysis
of poly(ethylene terephthalate). Macromolecules 42:5128–5138. https://
doi.org/10.1021/ma9005318.
18. Ribitsch D, Heumann S, Trotscha E, Herrero Acero E, Greimel K, Leber R,
Birner-Gruenberger R, Deller S, Eiteljoerg I, Remler P, Weber T, Siegert P,
Maurer K-H, Donelli I, Freddi G, Schwab H, Guebitz GM. 2011. Hydrolysis
of polyethyleneterephthalate by p-nitrobenzylesterase from Bacillus sub-
tilis. Biotechnol Prog 27:951–960. https://doi.org/10.1002/btpr.610.
19. Kawai F, Oda M, Tamashiro T, Waku T, Tanaka N, Yamamoto M, Miz-
ushima H, Miyakawa T, Tanokura M. 2014. A novel Ca
2⫹
-activated,
thermostabilized polyesterase capable of hydrolyzing polyethylene te-
rephthalate from Saccharomonospora viridis AHK190. Appl Microbiol
Biotechnol 98:10053–10064. https://doi.org/10.1007/s00253-014-5860-y.
20. Perz V, Baumschlager A, Bleymaier K, Zitzenbacher S, Hromic A, Stein-
kellner G, Pairitsch A, Łyskowski A, Gruber K, Sinkel C, Küper U, Ribitsch
D, Guebitz GM. 2016. Hydrolysis of synthetic polyesters by Clostridium
botulinum esterases. Biotechnol Bioeng 113:1024–1034. https://doi.org/
10.1002/bit.25874.
21. Perz V, Hromic A, Baumschlager A, Steinkellner G, Pavkov-Keller T,
Gruber K, Bleymaier K, Zitzenbacher S, Zankel A, Mayrhofer C, Sinkel C,
Kueper U, Schlegel K, Ribitsch D, Guebitz GM. 2016. An esterase from
anaerobic Clostridium hathewayi can hydrolyze aliphatic-aromatic poly-
esters. Environ Sci Technol 50:2899–2907. https://doi.org/10.1021/
acs.est.5b04346.
22. Yoshida S, Hiraga K, Takehana T, Taniguchi I, Yamaji H, Maeda Y, Toyo-
hara K, Miyamoto K, Kimura Y, Oda K. 2016. A bacterium that degrades
and assimilates poly(ethylene terephthalate). Science 351:1196–1199.
https://doi.org/10.1126/science.aad6359.
23. Ferrer M, Golyshina OV, Chernikova TN, Khachane AN, Martins dos
Santos VAP, Yakimov MM, Timmis KN, Golyshin PN. 2005. Microbial
enzymes mined from the Urania deep-sea hypersaline anoxic basin.
Chem Biol 12:895–904. https://doi.org/10.1016/j.chembiol.2005.05.020.
24. Sulaiman S, Yamato S, Kanaya E, Kim J-J, Koga Y, Takano K, Kanaya S.
2012. Isolation of a novel cutinase homolog with polyethylene
terephthalate-degrading activity from leaf-branch compost by using a
metagenomic approach. Appl Environ Microbiol 78:1556–1562. https://
doi.org/10.1128/AEM.06725-11.
25. Mayumi D, Akutsu-Shigeno Y, Uchiyama H, Nomura N, Nakajima-Kambe
T. 2008. Identification and characterization of novel poly(DL-lactic acid)
depolymerases from metagenome. Appl Microbiol Biotechnol 79:
743–750. https://doi.org/10.1007/s00253-008-1477-3.
26. Tchigvintsev A, Tran H, Popovic A, Kovacic F, Brown G, Flick R,
Hajighasemi M, Egorova O, Somody JC, Tchigvintsev D, Khusnutdinova
A, Chernikova TN, Golyshina OV, Yakimov MM, Savchenko A, Golyshin
PN, Jaeger K-E, Yakunin AF. 2015. The environment shapes microbial
enzymes: five cold-active and salt-resistant carboxylesterases from ma-
rine metagenomes. Appl Microbiol Biotechnol 99:2165–2178. https://
doi.org/10.1007/s00253-014-6038-3.
27. Müller CA, Obermeier MM, Berg G. 2016. Bioprospecting plant-
associated microbiomes. J Biotechnol 235:171–180. https://doi.org/
10.1016/j.jbiotec.2016.03.033.
28. Berg G, Grube M, Schloter M, Smalla K. 2014. Unraveling the plant
microbiome: looking back and future perspectives. Front Microbiol
5:148. https://doi.org/10.3389/fmicb.2014.00148.
29. Raghoebarsing AA, Smolders AJP, Schmid MC, Rijpstra WIC, Wolters-Arts
M, Derksen J, Jetten MSM, Schouten S, Sinninghe Damsté JS, Lamers
LPM, Roelofs JGM, Op den Camp HJM, Strous M. 2005. Methanotrophic
symbionts provide carbon for photosynthesis in peat bogs. Nature
436:1153–1156. https://doi.org/10.1038/nature03802.
30. Opelt K, Chobot V, Hadacek F, Schönmann S, Eberl L, Berg G. 2007.
Investigations of the structure and function of bacterial communities
associated with Sphagnum mosses. Environ Microbiol 9:2795–2809.
https://doi.org/10.1111/j.1462-2920.2007.01391.x.
31. Bragina A, Oberauner-Wappis L, Zachow C, Halwachs B, Thallinger GG,
Müller H, Berg G. 2014. The Sphagnum microbiome supports bog eco-
system functioning under extreme conditions. Mol Ecol 23:4498–4510.
https://doi.org/10.1111/mec.12885.
32. Moore T, Basiliko N. 2006. Decomposition in boreal peatlands, p
Müller et al. Applied and Environmental Microbiology
February 2017 Volume 83 Issue 4 e02641-16 aem.asm.org 12
125–143. In Wieder RK, Vitt DH (ed), Boreal peatland ecosystems.
Springer, Berlin, Germany.
33. Hájek T, Ballance S, Limpens J, Zijlstra M, Verhoeven JTA. 2010. Cell-wall
polysaccharides play an important role in decay resistance of Sphagnum
and actively depressed decomposition in vitro. Biogeochemistry 103:
45–57.
34. Steger D, Wentrup C, Braunegger C, Deevong P, Hofer M, Richter A,
Baranyi C, Pester M, Wagner M, Loy A. 2011. Microorganisms with
novel dissimilatory (bi)sulfite reductase genes are widespread and
part of the core microbiota in low-sulfate peatlands. Appl Environ
Microbiol 77:1231–1242. https://doi.org/10.1128/AEM.01352-10.
35. Kremer C, Pettolino F, Bacic A, Drinnan A. 2004. Distribution of cell
wall components in Sphagnum hyaline cells and in liverwort and
hornwort elaters. Planta 219:1023–1035. https://doi.org/10.1007/
s00425-004-1308-4.
36. Popper ZA. 2008. Evolution and diversity of green plant cell walls. Curr
Opin Plant Biol 11:286–292. https://doi.org/10.1016/j.pbi.2008.02.012.
37. Klavina L, Springe G, Nikolajeva V, Martsinkevich I, Nakurte I, Dzabijeva
D, Steinberga I. 2015. Chemical composition analysis, antimicrobial ac-
tivity and cytotoxicity screening of moss extracts (moss phytochemistry).
Molecules 20:17221–17243. https://doi.org/10.3390/molecules
200917221.
38. Petersen TN, Brunak S, von Heijne G, Nielsen H. 2011. SignalP 4.0:
discriminating signal peptides from transmembrane regions. Nat Meth-
ods 8:785–786. https://doi.org/10.1038/nmeth.1701.
39. Cha S-S, An YJ, Jeong C-S, Kim M-K, Jeon JH, Lee C-M, Lee HS, Kang SG,
Lee J-H. 2013. Structural basis for the

-lactamase activity of EstU1, a
family VIII carboxylesterase. Proteins 81:2045–2051. https://doi.org/
10.1002/prot.24334.
40. Jeon JH, Kim S-J, Lee HS, Cha S-S, Lee JH, Yoon S-H, Koo B-S, Lee C-M,
Choi SH, Lee SH, Kang SG, Lee J-H. 2011. Novel metagenome-derived
carboxylesterase that hydrolyzes

-lactam antibiotics. Appl Environ Mi-
crobiol 77:7830–7836. https://doi.org/10.1128/AEM.05363-11.
41. Mokoena N, Mathiba K, Tsekoa T, Steenkamp P, Rashamuse K. 2013.
Functional characterisation of a metagenome derived family VIII esterase
with a deacetylation activity on

-lactam antibiotics. Biochem Biophys
Res Commun 437:342–348.
42. Upton C, Buckley JT. 1995. A new family of lipolytic enzymes? Trends
Biochem Sci 20:178–179. https://doi.org/10.1016/S0968-0004(00)89002-7.
43. Hemilä H, Koivula TT, Palva I. 1994. Hormone-sensitive lipase is closely
related to several bacterial proteins, and distantly related to acetylcho-
linesterase and lipoprotein lipase: identification of a superfamily of
esterases and lipases. Biochim Biophys Acta 1210:249–253. https://
doi.org/10.1016/0005-2760(94)90129-5.
44. Biundo A, Hromic A, Pavkov-Keller T, Gruber K, Quartinello F, Haernvall
K, Perz V, Arrell MS, Zinn M, Ribitsch D, Guebitz GM. 2016. Characteriza-
tion of a poly(butylene adipate-co-terephthalate)-hydrolyzing lipase
from Pelosinus fermentans. Appl Microbiol Biotechnol 100:1753–1764.
https://doi.org/10.1007/s00253-015-7031-1.
45. Herrero Acero E, Ribitsch D, Steinkellner G, Gruber K, Greimel K, Eiteljo-
erg I, Trotscha E, Wei R, Zimmermann W, Zinn M, Cavaco-Paulo A, Freddi
G, Schwab H, Guebitz G. 2011. Enzymatic surface hydrolysis of PET: effect
of structural diversity on kinetic properties of cutinases from Thermobi-
fida. Macromolecules 44:4632–4640. https://doi.org/10.1021/
ma200949p.
46. Ferrer M, Bargiela R, Martínez-Martínez M, Mir J, Koch R, Golyshina OV,
Golyshin PN. 2015. Biodiversity for biocatalysis: a review of the
␣
/

-
hydrolase fold superfamily of esterases-lipases discovered in meta-
genomes. Biocatal Biotransformation 33:235–249. https://doi.org/
10.3109/10242422.2016.1151416.
47. Rashamuse K, Magomani V, Ronneburg T, Brady D. 2009. A novel family
VIII carboxylesterase derived from a leachate metagenome library ex-
hibits promiscuous

-lactamase activity on nitrocefin. Appl Microbiol
Biotechnol 83:491–500. https://doi.org/10.1007/s00253-009-1895-x.
48. Stoops JK, Hamilton SE, Zerner B. 1975. Carboxylesterases (EC 3.1.1): a
comparison of some kinetic properties of horse, sheep, chicken, pig, and
ox liver carboxylesterases. Can J Biochem 53:565–573. https://doi.org/
10.1139/o75-077.
49. Perz V, Bleymaier K, Sinkel C, Kueper U, Bonnekessel M, Ribitsch D,
Guebitz GM. 2016. Data on synthesis of oligomeric and polymeric po-
ly(butylene adipate-co-butylene terephthalate) model substrates for the
investigation of enzymatic hydrolysis. Data Brief 7:291–298. https://
doi.org/10.1016/j.dib.2016.02.029.
50. Müller CA, Oberauner-Wappis L, Peyman A, Amos GCA, Wellington EMH,
Berg G. 2015. Mining for nonribosomal peptide synthetase and
polyketide synthase genes revealed a high level of diversity in the
Sphagnum bog metagenome. Appl Environ Microbiol 81:5064–5072.
https://doi.org/10.1128/AEM.00631-15.
51. Blanusa M, Schenk A, Sadeghi H, Marienhagen J, Schwaneberg U. 2010.
Phosphorothioate-based ligase-independent gene cloning (PLICing): an
enzyme-free and sequence-independent cloning method. Anal Biochem
406:141–146. https://doi.org/10.1016/j.ab.2010.07.011.
52. Laemmli UK. 1970. Cleavage of structural proteins during the assembly
of the head of bacteriophage T4. Nature 227:680–685. https://doi.org/
10.1038/227680a0.
53. Thompson JD, Higgins DG, Gibson TJ. 1994. CLUSTAL W: improving the
sensitivity of progressive multiple sequence alignment through se-
quence weighting, position-specific gap penalties and weight matrix
choice. Nucleic Acids Res 22:4673–4680. https://doi.org/10.1093/nar/
22.22.4673.
54. Robert X, Gouet P. 2014. Deciphering key features in protein structures
with the new ENDscript server. Nucleic Acids Res 42:W320–W324.
https://doi.org/10.1093/nar/gku316.
55. Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. 2013. MEGA6:
Molecular Evolutionary Genetics Analysis version 6.0. Mol Biol Evol
30:2725–2729. https://doi.org/10.1093/molbev/mst197.
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