An Esterase from Anaerobic Clostridium hathewayi Can Hydrolyze

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An esterase from anaerobic Clostridium hathewayi can hydrolyze aliphatic aromatic polyesters Veronika Perz, Altijana Hromic, Armin Baumschlager, Georg Steinkellner, Tea PavkovKeller, Karl Gruber, Klaus Bleymaier, Sabine Zitzenbacher, Armin Zankel, Claudia Mayrhofer, Carsten Sinkel, Ulf Kueper, Katharina Schlegel, Doris Ribitsch, and Georg M. Guebitz Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.5b04346 • Publication Date (Web): 15 Feb 2016 Downloaded from http://pubs.acs.org on February 17, 2016

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Environmental Science & Technology

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An esterase from anaerobic Clostridium hathewayi can hydrolyze

2

aliphatic-aromatic polyesters

3

4

Veronika Perz a, Altijana Hromic b,c, Armin Baumschlager b, Georg Steinkellner b,

5

Tea Pavkov-Keller b, Karl Gruber b,c, Klaus Bleymaier b, Sabine Zitzenbacher b,

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Armin Zankel d, Claudia Mayrhofer d, Carsten Sinkel e, Ulf Kueper e, Katharina Schlegel e,

7

Doris Ribitsch a,f,*, Georg M. Guebitz a,f

8

a

9

b

10

c

Institute of Molecular Biosciences, University of Graz, Humboldtstrasse 50/III, 8010 Graz, Austria

11

d

Institute of Electron Microscopy and Nanoanalysis, Graz University of Technology, Steyrergasse 17/III, 8010

12

Graz, Austria

13

e

BASF SE, Carl-Bosch-Straße 38, 67056 Ludwigshafen, Germany

14

f

Institute of Environmental Biotechnology, University of Natural Resources and Life Sciences, Vienna, Konrad

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Lorenz Strasse 20, 3430 Tulln, Austria

Austrian Centre of Industrial Biotechnology ACIB, Konrad Lorenz Strasse 20, 3430 Tulln, Austria Austrian Centre of Industrial Biotechnology ACIB, Petersgasse 14, 8010, Graz, Austria

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17

*Corresponding author:

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Doris Ribitsch, [email protected], phone: ++43 1 47654 4974

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Keywords

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Microbial polyesterase, anaerobic esterase, polyester biodegradation, biogas batch, anaerobic

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biodegradation, Clostridium hathewayi, Hungatella hathewayi 1 ACS Paragon Plus Environment


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Abstract

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Recently a variety of biodegradable polymers have been developed as alternatives to

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recalcitrant materials. While many studies on polyester biodegradability have focused on

26

aerobic environments, there is much less known on biodegradation of polyesters in natural

27

and artificial anaerobic habitats. Consequently, the potential of anaerobic biogas sludge to

28

hydrolyze the synthetic compostable polyester PBAT (poly(butylene adipate-co-butylene

29

terephthalate) was evaluated in this study. Based on RP-HPLC analysis, accumulation of

30

terephthalic acid (Ta) was observed in all anaerobic batches within the first 14 days.

31

Thereafter a decline of Ta was observed, which occurred presumably due to consumption by

32

the microbial population. The esterase Chath_Est1 from the anaerobic risk 1 strain

33

Clostridium hathewayi DSM-13479 was found to hydrolyze PBAT. Detailed characterization

34

of this esterase including elucidation of the crystal structure was performed. The crystal

35

structure indicates that Chath_Est1 belongs to the α/β-hydrolases family. This study gives a

36

clear hint that also microorganisms in anaerobic habitats can degrade manmade PBAT.

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2 ACS Paragon Plus Environment

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Introduction

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Polymeric materials and polyesters are widely used in everyday life due to interesting material

40

properties such as low density, chemical resistance, simple process ability and low production

41

costs. Synthetic polymers have been designed to resist degradation since long-lasting

42

polymers were urgently needed for a multitude of applications in e.g. automotive industry,

43

electronics or building and construction. However, for other applications in agriculture (e.g.

44

mulch films), food packaging, organic waste- or carrier bags the persistence of polymer is not

45

needed or even undesirable. For that reason, biodegradable polyesters are in the focus of

46

research since a few years as they combine beneficial material properties with

47

biodegradability in a range of habitats.

48

The aliphatic-aromatic co-polyester PBAT (poly(butyleneadipate-co-butyleneterephthalate) is

49

a synthetic polyesters used e.g. in food packaging or organic waste bags. When using as

50

packaging materials, PBAT offers special barrier properties, e.g. its high water vapor

51

permeability, which predestines it for fruit and vegetable packaging, where it helps to prevent

52

growth of molds. PBAT is proven to be compostable

53

material for many compostable and biobased plastics. In microbial mineralization of

54

polyesters, hydrolysis by extracellular enzymes is the first step to produce oligomers /

55

monomers which can be further metabolized in the microbial cell. Thus, several research

56

groups have studied the enzymatic hydrolysis of PBAT. Generally, aerobic hydrolases like the

57

cutinase from Thermobifida cellulosilytica or Humicola insolens 3, the cutinase-like enzymes

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from Saccharomonospora viridis 4, 5, lipases from Pseudomonas sp.

59

Thermobifida fusca 7 are known to hydrolyze PBAT.

60

In recent years anaerobic digestion has gained importance in the biological treatment of

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organic

household wastes,

1, 2

which makes it an important raw

6

or a hydrolase from

often integrated in existing composting plants. The 3 ACS Paragon Plus Environment


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biodegradability of aliphatic-aromatic polyesters under composting conditions was

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demonstrated by different research groups. In contrast to aerobic composting there is

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considerably less information available on the hydrolysis in anaerobic environments. Here, an

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esterase from Clostridium hathewayi 8 namely Chath_Est1 was discovered. C. hathewayi was

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lately reclassified to Hungatella hathewayi 9, a representative of the new genus Hungatella of

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the family Clostridiaceae. The risk 1 strain was found to hydrolyze PBAT, which shows for

68

the first time that anaerobic microorganisms are able to degrade synthetic polymer.

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Experimental Section

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Chemicals and reagents

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Milled PBAT (poly(butylene adipate-co-butylene terephthalate), PBAT oligomers (oPBAT)

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and PHB (Polyhydroxybutyrate) as well as oPBAT and PBAT pellets were kindly provided

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by BASF SE (Germany). The PET model substrate bis(benzoyloxyethyl)terephthalate

75

(BaETaEBa) was synthesized according to the method described by Heumann et al. 10.

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Oligomeric

77

mono(4-hydroxybutyl) terephthalate

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O,O’-(butane-1,4-diyl) bis(4-hydroxybutyl) diterephthalate (BTaBTaB) were synthesized as

79

previously described 11. NMR and HPLC-MS were used to confirm purity.

80

Methanol and acetonitrile were of HPLC grade and purchased from Roth (Karlsruhe,

81

Germany). All other chemicals were of analytical grade and purchased from Sigma-Aldrich

82

(Germany).

83

Carrez reagent I and Carrez reagent II were prepared by dissolving 1.06 g K4[Fe(CN)6]·3H2O

84

or 2.88 g ZnSO4·7H2O in 10 ml ddH2O respectively.

PBAT

model

substrates

bis(4-hydroxybutyl) terephthalate (BTa)

(BTaB),

and


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86

Anaerobic batch cultures

87

To study the ability of anaerobic microbial populations to adapt to the synthetic polyester

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PBAT and the oligomeric PBAT model substrate oPBAT, anaerobic batch cultures were

89

prepared by mixing 170 ml anaerobic inoculum with 330 ml mineral salt medium in 1000 ml

90

flasks. Inoculation sludge was originally collected at the biogas plant Strem (Burgenland,

91

Austria) utilizing energy crops, which were fully fermented under mesophilic conditions

92

(37°), sieved to exclude biomass particles > 2.0 mm and pooled before application in

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cultivation experiments. Before usage in the described experiments, the sludge was kept under

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mesophilic conditions for several months and fed sporadically with maize silage. The mineral

95

salt medium (EN ISO 11734 L47 “Evaluation of the ultimate anaerobic biodegradability of

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organic compounds in digested sludge”, 1998) had the following composition: 0.27 g

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KH2PO4, 1.12 g Na2HPO4·12H2O, 0.53 g NH4Cl, 0.075 g CaCl2·2H2O, 0.1 g MgCl2·6H2O,

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0.02 g FeCl2·4H2O, 0.001 g resazurin, 0.1 g Na2S, 1000 ml ddH2O, 1 ml trace element

99

solution. The pH was adjusted to 7.0 using 1 M HCl. Trace element solution 12 contained:

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70 mg ZnCl2, 100 mg MnCl2·4H2O, 200 mg CoCl2·6H2O, 100 mg NiCl2·6H2O, 20 mg

101

CuCl2·2H2O, 50 mg NaMoO4·2H2O, 26 mg Na2SeO3·5H2O, 1 ml HCl (25 %), 1000 ml

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ddH2O. The sludge was flushed with oxygen-free nitrogen gas for 20 min l-1 to achieve

103

anaerobic cultures. In order to induce polyesterase expression, 0.5 g of the microbial

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polyesters PHB was fed-batch-wise added on day 0 and subsequently over 30 days to the

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sample batches oPBAT+ and PBAT+. The PBAT- batch that was not induced with PHB, was

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fed with 0.5 g maize silage instead. 1 g of milled oPBAT or PBAT (particle size 100-300 µm)

107

was introduced to batch experiments on day 0 to reach a final sample concentration of

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0.2 % w/v. The control biogas batch was spiked with 6.5 mM of Ta and adipic acid (Ada) on

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day 0. Anaerobic control batches containing mineral salt medium, 0.02 % w/v sodium azide 5 ACS Paragon Plus Environment


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and the respective substrates were incubated simultaneously at 37 °C. Hydrolysis of PBAT

111

and oPBAT was monitored over a time period of 60 days by means of RP-HPLC analysis.

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Samples were taken as triplicates in regular intervals.

113

114

HPLC analysis of hydrolysis products in anaerobic batch cultures

115

Prior to HPLC analysis, a modified Carrez precipitation

116

interfering substances from the complex sample matrix. One milliliter was withdrawn from

117

each biogas batch or abiotic control. Samples were centrifuged (Hermle Z300K, MIDSCI,

118

Missouri) at 14,000 g for 10 min at 20 °C. Two hundred microliters of each supernatant were

119

then transferred into fresh 1.5 ml reaction tubes, mixed with 780 µl ddH2O and 10 µl Carrez

120

reagent I. The mixture was vortexed and incubate for 1 min at 20 °C. Then, 10 µl of Carrez

121

reagent II were added and the mixture was vortexed again. Precipitation was conducted at

122

20 °C for 20 min followed by a centrifugation step (14,000 g, 4 °C, 30 min). 400 µl of the

123

supernatant were mixed with 400 µl methanol, acidified with formic acid to reach a pH of 3.0

124

and incubated for 60 min at 4°C. Samples were centrifuge again (14,000 g, 15 min, 0 °C) and

125

analyzed by means of RP-HPLC with the method that was previously described 3.

13, 14

was performed to remove

126

127

Extraction of terephthalic acid from Biogas sludge

128

To ensure that a measured decrease of Ta concentration in the batch cultures was not caused

129

by precipitation of Ta, a DMSO extraction was performed 133 days after the start of the

130

experiment. Forty milliliters sludge were withdrawn from each batch and centrifuged

131

(4,000 g, 20 min, 20 °C). Pellets were resuspended in 100 ml DMSO and rotated for 15 min at

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30 rpm on a rotation mixer (WiseMix RT-10, Wisd Laboratory Instruments, Czech Republic). 6 ACS Paragon Plus Environment

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The mixture was centrifuged again (4,000 g, 20 min, 20 °C), supernatants were Carrez

134

precipitated and prepared for RP-HPLP as described above.

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136

Scanning electron microscopy of biofilm formation on polyester pellets

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To investigate biofilm formation on the polyesters, pellets of PBAT and oPBAT (diameter

138

0.5 cm) were introduced to a separate batch culture using in-situ concentrate bags

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(6.75×12 cm; Ankom, Macedon, NY, USA) in order to facilitate recovery of the pellets after

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batch operation. Pellets were removed from the anaerobic batches after 86 days and prepared

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for scanning electron microscopy. For morphological investigations scanning electron

142

microscopy (Zeiss ULTRA 55, Carl Zeiss Micro Imaging GmbH, Germany) was performed at

143

primary electron energy of 5 keV. The (poly)ester pellets carrying biological material were

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imaged with the high-efficiency “in-lens SE detector” using secondary electrons (SE), in

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order to get topographic contrast 15, 16. Before application, fresh samples were fixed overnight

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using glutaraldehyde (3 % in 0.1 mol cacodylic acid sodium salt trihydrate, pH 7.2 at 4 °C),

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stained and double fixed for at least 3 h at 4 °C with 2 % OsO4 (in 0.1 mol cacodylic acid

148

sodium salt trihydrate) dehydrated in a graded ethanol series with a final step in propylene

149

oxide and ultimately dried by critical point drying (CO2). For observation, prepared samples

150

were mounted on aluminum stubs using double-sided carbon tape and sputter coated with

151

Au/Pd.

152

153

Investigation of hydrolytic activity of C. hathewayi

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The potential of C. hathewayi to hydrolyze PBAT in vivo was analyzed. For this,

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C. hathewayi cultures (Strain DSM-13479 obtained from DSMZ) were grown at 30 °C as

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recommended from DSMZ under anaerobic conditions in 6 × 50 ml PYX medium (DSMZ 7 ACS Paragon Plus Environment


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medium 104b) supplemented with 0.04 g/l glucose and 10 µM potassium buffer (pH 7.0). The

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cells were harvested anaerobically in the late exponential growth phase (OD 0.18). The cells

159

were centrifuged (3000 g, 10 min, 4 °C) and washed with PYX medium for two times to

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remove all remaining supplements. After washing procedure, the cells were resuspended in

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PYX medium to a final OD of 0.6. Each 30 ml of the cell suspension were supplemented with

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0.04 g/l glucose and 10 µM phosphate buffer, or PBAT and oPBAT and 10 µM phosphate

163

buffer, or 10 µM phosphate buffer as a negative control. The sample were incubated for 24 h

164

at 30 °C. Afterwards, 25 ml of cell suspension were harvested (3,200 g, 4 °C, 30 min) and

165

supernatant as well as cells were frozen separately. To analyze the hydrolytic activity of

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enzymes from C. hathewayi the supernatants (after centrifugation of the cells) were tested. To

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do this, 10 mg of PBAT were incubated with 2 ml supernatant for 6 days at 100 rpm and

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37 °C in a thermomixer (Thermomixer comfort, Eppendorf, Germany). The control (2 ml

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PYX medium with 10 mg PBAT) was treated simultaneously. Additionally, supernatants of

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all three cultures as well as PYX medium alone were analyzed for hydrolysis products prior to

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incubation with PBAT. Prior to HPLC quantification, a Carrez precipitation was performed.

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Hence, 960 µl sample and 20 µl Carrez reagent I were vortexed and incubate for 1 min at

173

20 °C. Then, 20 µl of Carrez reagent II were added, the mixture was vortexed and precipitated

174

at 20 °C for 20 min followed by a centrifugation step (14,000 g, 4 °C, 30 min). The

175

supernatant was acidified with 6 N HCl to reach a pH of 3.0 and the released hydrolysis

176

products Ta, BTa and BTaB were detected and quantified by means of RP-HPLC 3.

177

178

General recombinant DNA techniques

179

Molecular cloning of the esterase gene was performed by standard methods as described by

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Sambrook et al. 17. Digestion of DNA with restriction endonucleases NdeI and HindIII (New

181

England Biolabs, USA), dephosphorylation with alkaline phosphatase (Roche, Germany) and 8 ACS Paragon Plus Environment

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ligation with T4 DNA-ligase (Fermentas, Germany) were performed as described in the

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manufacturer’s protocols. Plasmid DNA was prepared using the Plasmid Mini Kit from

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Qiagen (Germany). DNA fragments and plasmids were purified by the Qiagen DNA

185

purification kits (Qiagen, Germany) and vector pET26b(+) (Novagen) was used for

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expression of the esterases carrying a C-terminal 6×HisTag in E. coli BL21-Gold(DE3)

187

(Stratagene).

188

189

Cloning of Chath_Est1

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The gene coding for the partial para-nitrobenzyl esterase from Clostridium hathewayi

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DSM13479 (accession# EFC94627.1) was completed according to the sequence alignment

192

with homologous proteins (Supporting information Figure S1) and codon optimized for

193

expression in E. coli and synthesized by GeneArt® (Life Technologies). The nucleotide

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sequence coding for Chath_Est1 fused to the C-terminal 6×His Tag comprises 1592 base pairs

195

with a GC content of 55 %. The plasmid carrying the target gene was digested, purified, and

196

ligated into expression vector pET26b(+) (Novagen, Merck KGaA, Germany) over the NdeI

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and HindIII restriction sites. The expression vector was then transformed into electro

198

competent E. coli BL21-Gold(DE3) cells.

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The gene was expressed as a fusion protein carrying a C-terminal 6×HisTag with a molecular

200

weight of 59.2 kDa.

201

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DNA sequencing, alignments and deposition of sequence data

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DNA was sequenced as custom service by Agowa (Germany) and DNA analysis was

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performed with Vector NTI Suite 10 (Invitrogen, USA). ExPASy (Expert Protein Analysis 9 ACS Paragon Plus Environment


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System) proteomics server of the Swiss Institute of Bioinformatics was used to perform a

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BLAST search, and sequences of related proteins were aligned using the Clustal W program

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(Swiss EMBnet node server). The complete and codon optimized nucleotide sequence of

208

Chath_Est1 was deposited in the GenBank database under accession number KT222913.

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210

Expression, purification and protein concentration of Chath_Est1

211

The expression at 20 °C and purification of Chath_Est1 was performed in accordance to the

212

previously described expression and purification of Cbotu_EstA and Cbotu_EstB

213

concentration of the purified enzyme solution was measured using the BIO-Rad Protein Assay

214

(BIO-RAD) and bovine serum albumin as a standard. SDS Page was performed in accordance

215

to Laemmli et al.

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R-250.

19

18

Protein

and staining of the proteins was done with Coomassie Brilliant Blue

217

218

Esterase activity and stability

219

Esterase activity and kinetic parameters were determined at pH 7.0 and 25 °C as previously

220

described

221

substrates.

222

To determine the temperature optimum Chath_Est1 was incubated in 50 mM potassium

223

phosphate buffer pH 7.0 at 25 °C, 37 °C, 50 °C and 300 rpm. Activity was measured using

224

pNPB as a substrate with the final concentrations of 7.9 mM. The activity determined at time

225

point zero was defined to be 100 %, the decrease in activity was followed for 35 days and the

226

activity half-life of Chath_Est1 was calculated.

18

using para-nitrophenylacetate (pNPA) and para-nitrophenylbutyrate (pNPB) as


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The pH optimum of Chath_Est1 was determined by following the hydrolysis of

228

bis-(2-benzoyloxy)ethyl) terephthalate (BaETaEBa) at different pHs. For this purpose, 10 mg

229

BaETaEBa were incubated for 23 h at 37 °C and 300 rpm with 6 µM Chath_Est1 in 2 ml

230

buffered solutions with 50 mM citrate phosphate at pH 4.0, pH 5.0 and pH 6.0, potassium

231

phosphate at pH 7.0, pH 8.0 and pH 9.0 and glycine-NaOH at pH 10.0. Samples were

232

withdrawn 18, and RP-HPLC analysis was performed 3 as previously described.

233

The hydrolysis of BaETaEBa was additionally followed over time. Samples containing 2 ml

234

50 mM potassium phosphate buffer pH 7.0, 10 mg BaETaEBa and 6 µM Chath_Est1 were

235

incubated at 37 °C and 300 rpm for 0, 1, 3, 5, 8, 20 and 24 h. Samples were withdrawn and

236

analyzed as described above.

237

238

Hydrolysis of the polyester PBAT and oligomeric PBAT model substrates

239

The ability of Chath_Est1 to hydrolyze PBAT and the model substrates BTaB and BTaBTaB

240

was tested by incubating 0.6 µM Chath_Est1 with 10 mg PBAT or 13.5 mM BTaB and

241

BTaBTaB respectively. The samples were incubated for 48 h and 72 h at 37 °C and 300 rpm

242

on a thermomixer (Thermomixer comfort, Eppendorf, Germany) in a total volume of 1 ml

243

50 mM potassium phosphate buffered solution pH 7.0. Reactions were stopped, methanol

244

precipitation was performed and hydrolysis products Ta, Ada, BTa and BTaB were quantified

245

via RP-HPLC as previously described 3. Control reactions were performed simultaneously

246

without addition of Chath_Est1.

247

248

Crystallization of Chath_Est1



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Crystallization experiments were performed by the sitting-drop vapor diffusion method using

250

different commercial crystallization screens (the Index Screen from Hampton Research and

251

the Morpheus Screen from Molecular Dimensions). Sitting drops were prepared by mixing

252

0.5 µl of the protein solution (at a concentration of 30 mg/ml) with an equal volume of mother

253

liquor, which were pipetted using an ORYX 8 pipetting robot from Douglas Instruments. The

254

trays were incubated at 20 °C. First crystal clusters were observed after approximately one

255

week. Well diffracting Chath_Est1 crystals were obtained with 1.0 M ammonium sulfate,

256

0.1 M HEPES, pH 7.0 and 0.5 % w/v Polyethylene glycol 8000 (Index Screen, condition

257

C11).

258

259

Data collection and processing

260

X-ray diffraction data were collected to a maximum resolution of 1.9 Å on beamline P11

261

(λ=1.033 Å) at the PETRA III/DESY Hamburg, Germany. The crystals were monoclinic

262

(space group P21) with unit-cell parameters a=62.05 Å, b=241.83 Å and c=83.58 Å, β=90.0°.

263

The data were processed using the XDS package

264

replacement using the structure of the thermostable carboxylesterase Est55 from Geobacillus

265

stearothermophilus (PDB code: 2OGS, 36 % sequence identity)

266

built using the mr-Rosetta module of PHENIX

267

the asymmetric unit consistent with the calculated Matthews coefficient

268

rebuilding and refinement were performed using the programs Coot 24 and PHENIX 22. Clear

269

electron density was observed for the majority of the amino acids except the first N-terminal

270

residue (Met) and 20 C-terminal residues in all four chains. Residual density was interpreted

271

as phosphate ions in all chains. The final structure was validated using MolProbity

272

Detailed statistics pertaining to data processing and structure refinement are summarized in

22

20

. The structure was solved by molecular

21

. The initial model was

yielding four molecules of Chath_Est1 in 23

. Structure

25

.


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supporting information Table S1. The atomic coordinates and structure factors have been

274

deposited in the Protein Data Bank under the code 5A2G.

275

276

Modelling

277

To build the model of the covalent tetrahedral intermediate compound a molecular model of

278

methyl acetate was used in the first step to fit the molecule into the proposed active site of the

279

Chath_Est1 structure. The substrate was prepared and optimized using YASARA

280

carboxylic oxygen of the substrate in its anionic form was fit into the proposed oxyanion hole

281

and the methyl acetate was covalently bond to the active site serine oxygen (Ser-200)

282

mimicking the covalent tetrahedral intermediate. The protonation and tautomerisation states

283

of His residues were chosen according to hydrogen bonding networks. Asp, Glu, Arg and Lys

284

residues were treated as charged. The active site His-416 was modeled to be protonated. This

285

initial complex was subjected to a molecular mechanics optimization using YASARA using

286

the AMBER03 force field

287

energy-minimized complex was used to build and extend the tetrahedral intermediate to the

288

BTaB-substrate, and was also subjected to a molecular mechanics optimization. The final

289

polymer complex was built by successive adding new functional groups and polymer subunits

290

keeping the extensions as close as possible to the surface of the protein by also keeping

291

reasonable bond angles and dihedrals. Between all addition steps, energy minimization of the

292

complex structure was performed and water was deleted where it clashed with the added

293

functional groups. This model was then used for a final energy minimization step, resulting in

294

the proposed model of the active site complex. Figures were prepared using PyMOL (DeLano

295

WL (2002) The PyMOL Molecular Graphics System, Version 1.5.0.4 Schrödinger, LLC) and

296

surface hydrophobicity was generated using VASCo 28, 29.

27

26

. The

applying the standard optimization protocol. This



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Results and Discussion

298

Hydrolysis products in anaerobic batch cultures

299

Biodegradable polyesters like PBAT are used e.g. in organic waste bags or food packaging.

300

So far, kitchen- and organic waste was usually biologically treated in composting plants.

301

Nevertheless, the trend of the recent years is to recover energy from organic waste during

302

anaerobic digestion, often in combination with post-composting to degrade residual organic

303

waste

304

(composting)

305

biodegradation of PBAT. Consequently, the hydrolysis of PBAT by “anaerobic” enzymes was

306

investigated in this study.

307

In a first step, the general degradation ability of PBAT in biogas sludge was investigated.

308

SEM analysis of oPBAT, and PBAT pellets that were incubated in biogas sludge showed

309

formation of biofilm on the surface consisting both of cocci and rod-shaped bacteria.

310

(Figure 1A and supporting information Figure S2). RP-HPLC analysis of the control batch

311

that was spiked with 6.5 mM Ada and Ta revealed that Ada is metabolized in the sludge quite

312

efficiently. After 21 days the concentration of Ada in the spiked control batch was beneath the

313

detection limit of 0.5 mM (data not shown). In the PBAT sample batches (oPBAT+, PBAT+,

314

PBAT-) no Ada could be detected due to immediate metabolization when released from the

315

polymer. Generally, aliphatic compounds are easier biodegraded than aromatic ones. Sotto et

316

al. observed relatively fast biodegradation rates for different aliphatic monomers including

317

Ada 31. Consequently, the analysis of released molecules in the batch tests was focused on Ta.

318

In all PBAT batches, the Ta concentration raised within the first 14 days of incubation

319

indicating a certain degree of oPBAT and PBAT hydrolysis (Figure 1B). Interestingly, Ta

320

concentration started to decrease significantly after day 21 in all batches including the Ta

30

. The biodegradability of PBAT was proven and studied under aerobic conditions 1, 2

. However, there is considerably less known about the anaerobic


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spiked control batch. After 35 days, Ta was no longer detectable in the sample batches and

322

after 60 days only 6 % of initial Ta concentration was detected in the spiked control batch. In

323

abiotic control batches no hydrolysis products were detected. Ta extraction with DMSO that

324

was performed after 133 days did not regain Ta from any of the biogas batches including the

325

control batch. This led to the assumption that Ta was indeed metabolized by action of

326

microorganisms in all biogas batches. Several other research groups have already reported the

327

biodegradation of Ta by different aerobic microorganisms like species from Bacillus

328

Pseudomonas

329

biodegradation potential 35, 36 and were analyzed to identify the key organisms responsible for

330

Ta biodegradation

331

and starts to metabolize Ta after 3 weeks when alternative C-sources are limited.

332

Kleerebezem et al. 36 reported Ta degradation in a first stage reactor after 300 days and in the

333

second stage reactor from day 1.

334

One parameter that influences the hydrolysis of PBAT seems to be the size of the polymer as

335

the lower molecular weight model substrate oPBAT was hydrolyzed considerable faster.

336

Melting temperatures (Tm) of PBAT and oPBAT lay around 125 °C, however the melting

337

range of oPBAT was broader and started at lower temperatures. PBAT batch that was spiked

338

with the naturally occurring polyesters PHB with the aim to induce esterase expression.

339

(PBAT+) reached a maximum Ta concentration of about 20.0 µM after 10 days, whereas in

340

the PBAT batch without inducing PHB (PBAT-) the maximum Ta concentration was detected

341

to be 14.4 µM after 21 days. The maximum Ta concentration in PBAT+ was higher and

342

reached faster when compared to PBAT-. This indicates that PHB (i.e. hydrolysis products)

343

may induce secretion of enzymes that are capable of degrading PBAT.

33

and Arthrobacter

32

,

34

. Moreover, also anaerobic consortia exhibited Ta

37-39

. It might be that the biogas sludge needs a certain time to adapt to Ta

344

345 15 ACS Paragon Plus Environment


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346 347

Figure 1. (A) SEM image of oPBAT after incubation in biogas plant sludge inoculated anaerobic batch cultures

348

for 86 days. Additional images can be found in the supporting information Figure S2. (B) Liberation of

349

terephthalic acid (Ta) from PBAT and oPBAT incubated in biogas plant sludge inoculated anaerobic batch

350

cultures in presence (+) and absence (-) of PHB as inductor. As a control a batch culture was spiked with 6.5 mM

351

Ta. Each data point represents the average of three independent samples; displayed error bars indicate respective

352

minimum and maximum values. An example of an HPLC profile can be found in the supporting information

353

Figure S3.

354

355

In-silico search for anaerobic polysterases

356

Based on the above indication for anaerobic degradation of PBAT, the next step was

357

dedicated to identification of anaerobic organisms and enzymes responsible for PBAT

358

hydrolysis. Consequently, an in-silico screening for enzymes from anaerobic organisms with

359

activity on synthetic polyesters was performed in order to identify active microorganisms and

360

their respective polyesterases. Clostridium species belonging to the phylum of Firmicutes are

361

described to be omnipresent in biogas sludge

362

to biodegrade Ta 39. Several proteins extracted from a Ta degrading biofilm were annotated to

40, 41

including an anaerobic consortium known


Page 17 of 28


37

363

Clostridium

. Typically, Clostridium species are described to be important for degradation

364

of cellulosic material

365

poly(β-caprolactone) (PCL) and poly(β-hydroxybutyrate) (PHB)

366

identified Clostridium strains with significant activity on PCL did not hydrolyze

367

poly(butylene terephthalate-co-butylene adipate) 44.

368

An extensive search of the NCBI protein database revealed that the risk 1 strain C. hathewayi

369

(also Hungatella hathewayi) carries the gene for a putative, partial esterase which has been

370

completed according to the sequences of highly homologous proteins (Supporting information

371

Figure S1) and named Chath_Est1. Based on our own metagenomics based identification of

372

hydrolytic Clostridia and of others who found the closely related C. saccharolyticum WM1 in

373

biogas sludge

374

investigation since this strain attractive for bioaugmentation anaerobic communities to

375

enhance plastics degradation.

42

but they are also reported to hydrolyze the polyesters 43

. Nevertheless, the so far

45

, we have specifically selected the risk 1 strain C. hathewayi for further

376

377

Hydrolysis of PBAT by C. hathewayi cultures

378

To analyze the ability of C. hathewayi to hydrolyze PBAT, resting cells were pre-incubated in

379

PYX medium supplemented with PBAT and oPBAT. One sample supplemented with glucose

380

and one sample without further addition was used as control. C. hathewayi supernatants

381

obtained from these three pre-incubations were then incubated with PBAT. Degradation

382

products of PBAT were found, even though the amount of monomers differed. The

383

supernatants from the cultures that were pre-incubated with PBAT and oPBAT to stimulate

384

the expression of polyesterases showed an elevated level of released Ta and BTa when

385

compared to the cultures that were pre-incubated in PYX medium or PYX medium

386

supplemented with glucose (Figure 2). 17 ACS Paragon Plus Environment


Page 18 of 28

387

Ta

Total released molecules [µM]

70

BTa

BTaB

60 50 40 30 20 10 0 PYX

+ PYX

ose Gluc

388

PYX

+

T+o P BA

T PBA

389

Figure 2. Total released molecules after incubation of C. hathewayi cell supernatants with PBAT for 6 days at

390

37 °C and 100 rpm. Released molecules are shown in µM and were quantified by means of HPLC analysis. Ta:

391

terephthalic acid; BTa: mono(4-hydroxybutyl) terephthalate; BTaB: bis(4-hydroxybutyl) terephthalate. Each bar

392

represents the average of three independent samples; error bars indicate the standard deviation. C. hathewayi was

393

pre-incubated in PYX in absence or presence of glucose or PBAT and oPBAT and of the respective culture

394

supernatants were analyzed. Hydrolysis products quantified in controls incubated under the identical conditions

395

(i.e. culture supernatants without PBAT, PYX medium alone, PYX with PBAT) and were subtracted. Examples

396

for RP-HPLC profiles can be found in the Supporting information Figure S4.

397

398

Biochemical properties of Chath_Est1

399

Expression of the codon-optimized Chath_Est1 gene carrying the C-terminal 6×HisTag

400

resulted in production of soluble protein but also in significant amounts of inclusion bodies.


Page 19 of 28


401

Highest protein yields were obtained after 20 h of induction at 20 °C and 160 rpm (Supporting

402

information Figure S5).

403

The temperature stability of Chath_Est1 was tested at pH 7.0 at 25 °C, 37 °C and 50 °C with

404

the soluble substrate pNPB. Chath_Est1 exhibited the highest stability at 25 °C with 85 %

405

activity remaining after 35 days. The activity half-lives at 37 °C and 50 °C were determined

406

to be 10 days and 20 min, respectively. Chath_Est1 shows significant active in a pH range of

407

6.0 to 8.0 (Supporting information Figure S6). For this reason the neutral pH 7.0 was chosen

408

for further characterizations of the esterase.

409 410

Table 1. Kinetic parameters of Chath_Est1 on the soluble esterase standard substrates pNPA and pNPB.

pNPA pNPB

Vmax [µmol min-1 mg-1]

Km [mM]

kcat [sec-1]

kcat/Km [sec-1 mM-1]

2.50 ± 0.20 7.00 ± 0.20

2.00 ± 0.20 2.00 ± 0.20

2.47 6.91

1.23 3.45

411 412

The water-soluble esterase substrates pNPA and pNPB were utilized to determine the kinetic

413

parameters of Chath_Est1. Vmax and kcat of Chath_Est1 were more than 2 times higher for

414

pNPB compared to pNPA whereas Km values were in the same range for both standard

415

substrates.

416

417

Hydrolysis of PBAT and model substrates by Chath_Est1

418

During the incubation of BaETaEBa with Chath_Est1 the release of the hydrolysis products

419

ETa, Ta and Ba were quantified by RP-HPLC (Figure 3). ETaE and BaE were not found

420

leading to the assumption that these esters were fast hydrolyzed to their constituents and

421

therefore not accumulated in the reaction mixture. Taking into account the structure of the 19 ACS Paragon Plus Environment


Page 20 of 28

422

model substrate, Chath_Est1 seems to release the Ta subunits more efficiently than the Ba

423

subunits. The concentrations of released Ta and Ba are in the same range. However, two Ba

424

subunits but just one Ta would be available in the BaETaEBa molecule. This is in contrast to

425

the recently investigated esterases Cbotu_EstA and Cbotu_EstB from Clostridium botulinum,

426

which preferably released Ba 18.

427

428 429

Figure 3. Chemical structure of BaETaEBa and released molecules after hydrolysis with 6 µM Chath_Est1 at

430

37 °C and 300 rpm in 50 mM potassium phosphate buffer pH 7.0. Release molecules are shown in µM, ETaE:

431

bis(2-hydroxyethyl) terephthalate, ETa: mono(2-hydroxyethyl) terephthalate, Ta: terephthalic acid, BaE:

432

2-hydroxyethyl benzoate and Ba: benzoic acid. Quantification was performed with RP-HPLC. Each bar

433

represents the average of three independent samples; error bars indicate the standard deviation.

434 435

Chath_Est1 hydrolyzed PBAT as well as the oligomeric model substrates BTaB and

436

BTaBTaB that are possible fragments of the PBAT polymer. After 72 h the highest

437

concentration of released molecules was observed with the substrate BTaB followed by


Page 21 of 28


438

BTaBTaB and PBAT which was hydrolyzed slower by two orders of magnitude (Supporting

439

information Figure S7).

440

441

Overall structure of Chath_Est1

442

The X-ray crystal structure of Chath_Est1 was determined at 1.90 Å resolution (Supporting

443

information Table S1). The crystal belongs to the monoclinic space group P21) and contains

444

four esterase molecules in the asymmetric unit. According to an analysis of intermolecular

445

interactions in the crystal using the PDBePISA server

446

dimer in solution. The overall structure consists of a central β-sheet surrounded by α-helices

447

indicating that Chath_Est1 belongs to the family of α/β-hydrolases (Supporting information

448

Figure S8). The structures of the four crystallographically independent esterase molecules are

449

very similar with pairwise root-mean-square-deviations (rmsd) ranging from 0.208 to 0.220

450

calculated using the program PyMOL (DeLano WL (2002) The PyMOL Molecular Graphics

451

System, Version 1.5.0.4 Schrödinger, LLC).

452

Based on sequence and structure comparisons, Chath_Est1 contains a catalytic triad formed

453

by Ser-200, His-416 and Glu-323 (Figure 4). The entrance to the active site is formed by

454

loops and helices. Residues 67-74 and 430-434 form one side of the entrance whereas

455

residues 329-333 and 274-282 form the rest of the entrance. The active site is located at the

456

base of the open cavity (Figure 6) which was also observed in pNB esterase (PDB

457

code: 1C7I)

458

stearothermophilus (PDB code: 2OGS) 21. The presence of Glu instead of Asp in the catalytic

459

triad is similar to the situation in acetylcholine esterase

460

Chath_Est1 compared to carboxylesterase Est55 is about 36 % and about 34 % to

47

46

Chath_Est1 is most likely present as

and in the thermostable carboxylesterase Est55 from Geobacillus

48

. The sequence identity of



Page 22 of 28

461

pNB esterase, respectively 47 (Supporting information Figure S9). The active site residues are

462

in similar positions as in the other two structures.

463

464 465

Figure 4. Catalytic triad of Chath_Est1

466 467

We modeled a structure mimicking the tetrahedral intermediate formed during the hydrolysis

468

of the substrate (Figure 5 and supporting information Figure S10). In that model, the active

469

site can be divided into two regions by a plane defined by the hydroxyl oxygen atom of

470

Ser-200 as well as the central carbon atom and the oxyanion of the tetrahedral substrate

471

intermediate. The active site His-416 is located on one side of this plane and is correctly

472

positioned to protonate the alcoholic leaving group. The oxyanion hole is formed by the helix,

473

at the N-terminal end of which the active site Ser-200 is located, and by the main chain amide

474

groups of Gly-114, Ala-115 and Gly-201.

475 476


Page 23 of 28


477 478

Figure 5. Left: Model of the Chath_Est1 dimer structure with the open active site in complex with the poly

479

substrate (shown in stick representation). The surface is colored by hydrophobicity values (blue hydrophilic to

480

red hydrophobic). Right: Proposed covalent tetrahedral intermediate of the poly-substrate. The oxyanion hole

481

and the active site residues are shown in a sticks representation. The covalently bound poly-substrate is shown in

482

yellow. The surface is shown transparent and colored by hydrophobicity values (blue hydrophilic to red

483

hydrophobic).

484 485

In conclusion, with the here described results we tried to portray anaerobic biodegradation of

486

PBAT from the crude sludge via an anaerobic organism through to a mechanistic focus on

487

polyesterases that are responsible for the initial enzymatic hydrolysis. We have shown that

488

typical anaerobic sludges are able to hydrolyze PBAT to a certain degree and the released

489

monomers Ada and Ta are not detectable anymore. Nevertheless, PBAT hydrolysis rates are

490

very low. Based on the presented results it is impossible to say if PBAT is hydrolyzed

491

completely and the efficient degradation of PBAT in industrial biogas plants with retention

492

times of about 3 weeks is not feasible so far. The application of polyesterase inducing

493

naturally occurring polyesters like PHB might be an option to stimulate the growth of

494

microorganisms expressing hydrolytic enzymes with activity on PBAT for improved PBAT

495

hydrolysis in anaerobic environments. In addition, we have cloned and heterologously

496

expressed the esterase Chath_Est1 from the anaerobic risk 1 strain C. hathewayi that

497

hydrolyzes the synthetic polyester PBAT. The crystal structure of Chath_Est1 was solved and 23 ACS Paragon Plus Environment


Page 24 of 28

498

will help to improve the knowledge about polyesterases in general. To close the circle, our

499

experiments indicate that C. hathewayi cells (i.e. their enzymes in the supernatants) are

500

hydrolyzing PBAT.

501

The present study gives a hint about how and by which kind of microorganisms these

502

manmade plastics could be biodegraded in anaerobic environments. The knowledge about

503

microorganisms that are hydrolyzing polyesters like PBAT could also open the way for new

504

application like addition of cell or spore suspension to biogas plants in order to enhance the

505

anaerobic biodegradation of biodegradable polymers that are e.g. used in food packaging or

506

organic waste bags.

507

508

Associated content

509

Supporting information. Additional information and data on crystographic paramenters and

510

crystal structures, amino acid sequence alignments, SEM images, protein expression, pH

511

optimum determination and hydrolysis experiments. This material is available free of charge

512

via the Internet at http://pubs.acs.org.

513

514

Author information

515

Corresponding Author: *Email: [email protected]

516

517

518 24 ACS Paragon Plus Environment

Page 25 of 28


519

Acknowledgment

520

This work has been supported by the Federal Ministry of Science, Research and Economy

521

(BMWFW), the Federal Ministry of Traffic, Innovation and Technology (bmvit), the Styrian

522

Business Promotion Agency SFG, the Standortagentur Tirol, the Government of Lower

523

Austria and ZIT - Technology Agency of the City of Vienna through the COMET-Funding

524

Program managed by the Austrian Research Promotion Agency FFG.

525

526

References

527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554 555 556 557

1. Witt, U.; Einig, T.; Yamamoto, M.; Kleeberg, I.; Deckwer, W. D.; Müller, R. J., Biodegradation of aliphatic–aromatic copolyesters: evaluation of the final biodegradability and ecotoxicological impact of degradation intermediates. Chemosphere 2001, 44, (2), 289-299. 2. Kijchavengkul, T.; Auras, R.; Rubino, M.; Selke, S.; Ngouajio, M.; Fernandez, R. T., Biodegradation and hydrolysis rate of aliphatic aromatic polyester. Polymer Degradation and Stability 2010, 95, (12), 2641-2647. 3. Perz, V.; Bleymaier, K.; Sinkel, C.; Kueper, U.; Bonnekessel, M.; Ribitsch, D.; Guebitz, G. M., Substrate specificities of cutinases on aliphatic–aromatic polyesters and on their model substrates. New Biotechnology 2016, 33, (2), 295-304. 4. Kawai, F.; Oda, M.; Tamashiro, T.; Waku, T.; Tanaka, N.; Yamamoto, M.; Mizushima, H.; Miyakawa, T.; Tanokura, M., A novel Ca2+-activated, thermostabilized polyesterase capable of hydrolyzing polyethylene terephthalate from Saccharomonospora viridis AHK190. Applied Microbiology and Biotechnology 2014, 98, (24), 10053-10064. 5. Shingo Kawai*, M. N. H. O., Degradation mechanisms of a nonphenolic -O-4 lignin model dimer by Trametes versicolor laccase in the presence of 1-hydroxybenzotriazole. Enzyme and microbial technology 2002, 30, 482-489. 6. Marten, E.; Müller, R.-J.; Deckwer, W.-D., Studies on the enzymatic hydrolysis of polyesters. II. Aliphatic–aromatic copolyesters. Polymer Degradation and Stability 2005, 88, (3), 371-381. 7. Kleeberg, I.; Welzel, K.; VandenHeuvel, J.; Mueller, R. J.; Deckwer, W. D., Characterization of a New Extracellular Hydrolase from Thermobifida fusca Degrading Aliphatic−AromaHc Copolyesters. Biomacromolecules 2005, 6, (1), 262-270. 8. Steer, T.; Collins, M. D.; Gibson, G. R.; Hippe, H.; Lawson, P. A., Clostridium hathewayi sp. nov., from Human Faeces. Systematic and Applied Microbiology 2001, 24, (3), 353-357. 9. Kaur, S.; Yawar, M.; Kumar, P. A.; Suresh, K., Hungatella effluvii gen. nov., sp. nov., an obligately anaerobic bacterium isolated from an effluent treatment plant, and reclassification of Clostridium hathewayi as Hungatella hathewayi gen. nov., comb. nov. Int J Syst Evol Microbiol 2014, 64, (Pt 3), 710-8. 10. Heumann, S.; Eberl, A.; Pobeheim, H.; Liebminger, S.; Fischer-Colbrie, G.; Almansa, E.; Cavaco-Paulo, A.; Gubitz, G. M., New model substrates for enzymes hydrolysing polyethyleneterephthalate and polyamide fibres. Journal of Biochemical and Biophysical Methods 2006, 69, (1-2), 89-99. 25 ACS Paragon Plus Environment


558 559 560 561 562 563 564 565 566 567 568 569 570 571 572 573 574 575 576 577 578 579 580 581 582 583 584 585 586 587 588 589 590 591 592 593 594 595 596 597 598 599 600 601 602 603 604 605 606 607 608 609

Page 26 of 28

11. Perz, V.; Bleymaier, K.; Sinkel, C.; Kueper, U.; Bonnekessel, M.; Ribitsch, D.; Guebitz, G. M., Data on synthesis of oligomeric and polymeric poly(butylene adipate-co-butylene terephthalate) model substrates for the investigation of enzymatic hydrolysis. Data in Brief 2016, submitted. 12. Schlegel, H. G., Allgemeine Mikrobiologie. Georg Thieme Verlag: Stuttgart - New York, 1992; Vol. 7, p 634. 13. Carrez, C., Estimation of lactose in Milk. Ann. Chim. Anal. 1908, 13, 17-22. 14. Culhaoglu, T.; Zheng, D.; Mechin, V.; Baumberger, S., Adaptation of the Carrez procedure for the purification of ferulic and p-coumaric acids released from lignocellulosic biomass prior to LC/MS analysis. J Chromatogr B Analyt Technol Biomed Life Sci 2011, 879, (28), 3017-22. 15. Weiss, S.; Lebuhn, M.; Andrade, D.; Zankel, A.; Cardinale, M.; Birner-Gruenberger, R.; Somitsch, W.; Ueberbacher, B. J.; Guebitz, G. M., Activated zeolite--suitable carriers for microorganisms in anaerobic digestion processes? Appl Microbiol Biotechnol 2013, 97, (7), 3225-38. 16. Goldstein, J. I.; Newbury, D. E.; Echlin, P.; Joy, D.; Lyman, C. E.; Lifshin, E.; Sawyer, L.; Michael, J., Scanning Electron Microscopy and X-Ray Microanalysis. Kluwer Academic/Plenum Publisher: New York, 2003; p 690. 17. Sambrook, J. E.; Fritsch, E. F.; Maniatis, T., Molecular Cloning: A Laboratory Manual. Cold Spring Harbor: New York, 1989; Vol. 2nd ed. 18. Perz, V.; Baumschlager, A.; Bleymaier, K.; Zitzenbacher, S.; Hromic, A.; Steinkellner, G.; Pairitsch, A.; Łyskowski, A.; Gruber, K.; Sinkel, C.; Küper, U.; Ribitsch, D.; Guebitz, G. M., Hydrolysis of synthetic polyesters by Clostridium botulinum esterases. Biotechnology and Bioengineering 2015, in press. 19. Laemmli, U. K., Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 1970, 227, (5259), 680-5. 20. Kabsch, W., XDS. Acta Crystallogr D Biol Crystallogr 2010, 66, (Pt 2), 125-32. 21. Liu, P.; Ewis, H. E.; Tai, P. C.; Lu, C. D.; Weber, I. T., Crystal structure of the Geobacillus stearothermophilus carboxylesterase Est55 and its activation of prodrug CPT-11. In J Mol Biol, England, 2007; Vol. 367, pp 212-23. 22. Adams, P. D.; Afonine, P. V.; Bunkoczi, G.; Chen, V. B.; Davis, I. W.; Echols, N.; Headd, J. J.; Hung, L. W.; Kapral, G. J.; Grosse-Kunstleve, R. W.; McCoy, A. J.; Moriarty, N. W.; Oeffner, R.; Read, R. J.; Richardson, D. C.; Richardson, J. S.; Terwilliger, T. C.; Zwart, P. H., PHENIX: a comprehensive Python-based system for macromolecular structure solution. Acta Crystallogr D Biol Crystallogr 2010, 66, (Pt 2), 213-21. 23. Winn, M. D.; Ballard, C. C.; Cowtan, K. D.; Dodson, E. J.; Emsley, P.; Evans, P. R.; Keegan, R. M.; Krissinel, E. B.; Leslie, A. G.; McCoy, A.; McNicholas, S. J.; Murshudov, G. N.; Pannu, N. S.; Potterton, E. A.; Powell, H. R.; Read, R. J.; Vagin, A.; Wilson, K. S., Overview of the CCP4 suite and current developments. Acta Crystallogr D Biol Crystallogr 2011, 67, (Pt 4), 235-42. 24. Emsley, P.; Lohkamp, B.; Scott, W. G.; Cowtan, K., Features and development of Coot. Acta Crystallogr D Biol Crystallogr 2010, 66, (Pt 4), 486-501. 25. Chen, V. B.; Arendall, W. B., 3rd; Headd, J. J.; Keedy, D. A.; Immormino, R. M.; Kapral, G. J.; Murray, L. W.; Richardson, J. S.; Richardson, D. C., MolProbity: all-atom structure validation for macromolecular crystallography. Acta Crystallogr D Biol Crystallogr 2010, 66, (Pt 1), 12-21. 26. Krieger, E.; Koraimann, G.; Vriend, G., Increasing the precision of comparative models with YASARA NOVA - a self-parameterizing force field. Proteins: Structure, Function, and Bioinformatics 2002, 47, (3), 393-402. 27. Duan, Y.; Wu, C.; Chowdhury, S.; Lee, M. C.; Xiong, G.; Zhang, W.; Yang, R.; Cieplak, P.; Luo, R.; Lee, T.; Caldwell, J.; Wang, J.; Kollman, P., A Point-Charge Force Field for Molecular Mechanics Simulations of Proteins Based on Condensed-Phase Quantum Mechanical Calculations. J. Comput. Chem. 2003, 24, (16), 1999-2012. 28. Steinkellner, G.; Rader, R.; Thallinger, G.; Kratky, C.; Gruber, K., VASCo: computation and visualization of annotated protein surface contacts. BMC Bioinformatics 2009, 10, (1), 32. 29. Sanner, M. F.; Olson, A. J.; Spehner, J. C., Reduced surface: An efficient way to compute molecular surfaces. Biopolymers 1996, 38, (3), 305-320. 26 ACS Paragon Plus Environment

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30. Chang, N.-B.; Pires, A., Sustainable Solid Waste Management: A Systems Engineering Approach. John Wiley & Sons Inc.: Hoboken, New Jersey, 2015; p 936. 31. Siotto, M.; Sezenna, E.; Saponaro, S.; Innocenti, F. D.; Tosin, M.; Bonomo, L.; Mezzanotte, V., Kinetics of monomer biodegradation in soil. Journal of Environmental Management 2012, 93, (1), 3137. 32. Karegoudar, T. B.; Pujar, B. G., Degradation of terephthalic acid by a Bacillus species. FEMS Microbiology Letters 1985, 30, (1-2), 217-220. 33. Wang, Z.-J.; Teng, L.-h.; Zhang, J.; Huang, X.-L.; Zhang, J.-F., Study on optimal biodegradation of terephthalic acid by an isolated Pseudomonas sp. African Journal of Biotechnology 2011, 10, (16), 3143-3148. 34. Zhang, Y.-M.; Sun, Y.-Q.; Wang, Z.-J.; Zhang, J., Degradation of terephthalic acid by a newly isolated strain of Arthrobacter sp.0574. South African Journal of Science 2013, 109, (7/8), 1-4. 35. Kleerebezem, R.; Pol, L. W.; Lettinga, G., Anaerobic biodegradability of phthalic acid isomers and related compounds. Biodegradation 1999, 10, (1), 63-73. 36. Kleerebezem, R.; Beckers, J.; Hulshoff Pol, L. W.; Lettinga, G., High rate treatment of terephthalic acid production wastewater in a two-stage anaerobic bioreactor. Biotechnol Bioeng 2005, 91, (2), 169-79. 37. Wu, J. H.; Wu, F. Y.; Chuang, H. P.; Chen, W. Y.; Huang, H. J.; Chen, S. H.; Liu, W. T., Community and proteomic analysis of methanogenic consortia degrading terephthalate. Appl Environ Microbiol 2013, 79, (1), 105-12. 38. Wu, J. H.; Liu, W. T.; Tseng, I. C.; Cheng, S. S., Characterization of microbial consortia in a terephthalate-degrading anaerobic granular sludge system. Microbiology 2001, 147, (Pt 2), 373-82. 39. Lykidis, A.; Chen, C. L.; Tringe, S. G.; McHardy, A. C.; Copeland, A.; Kyrpides, N. C.; Hugenholtz, P.; Macarie, H.; Olmos, A.; Monroy, O.; Liu, W. T., Multiple syntrophic interactions in a terephthalate-degrading methanogenic consortium. ISME J 2011, 5, (1), 122-30. 40. Krause, L.; Diaz, N. N.; Edwards, R. A.; Gartemann, K.-H.; Krömeke, H.; Neuweger, H.; Pühler, A.; Runte, K. J.; Schlüter, A.; Stoye, J.; Szczepanowski, R.; Tauch, A.; Goesmann, A., Taxonomic composition and gene content of a methane-producing microbial community isolated from a biogas reactor. Journal of Biotechnology 2008, 136, (1–2), 91-101. 41. Weiland, P., Biogas production: current state and perspectives. Applied Microbiology and Biotechnology 2010, 85, (4), 849-860. 42. Schluter, A.; Bekel, T.; Diaz, N. N.; Dondrup, M.; Eichenlaub, R.; Gartemann, K. H.; Krahn, I.; Krause, L.; Kromeke, H.; Kruse, O.; Mussgnug, J. H.; Neuweger, H.; Niehaus, K.; Puhler, A.; Runte, K. J.; Szczepanowski, R.; Tauch, A.; Tilker, A.; Viehover, P.; Goesmann, A., The metagenome of a biogasproducing microbial community of a production-scale biogas plant fermenter analysed by the 454pyrosequencing technology. J Biotechnol 2008, 136, (1-2), 77-90. 43. Abou-Zeid, D. M.; Muller, R. J.; Deckwer, W. D., Degradation of natural and synthetic polyesters under anaerobic conditions. Journal of Biotechnology 2001, 86, (2), 113-126. 44. Abou-Zeid, D. M.; Muller, R. J.; Deckwer, W. D., Biodegradation of aliphatic homopolyesters and aliphatic-aromatic copolyesters by anaerobic microorganisms. Biomacromolecules 2004, 5, (5), 1687-97. 45. Murray, W. D.; Khan, A. W.; van den Berg, L., Clostridium saccharolyticum sp. nov., a Saccharolytic Species from Sewage Sludge. International Journal of Systematic and Evolutionary Microbiology 1982, 32, (1), 132-135. 46. Krissinel, E.; Henrick, K., Inference of macromolecular assemblies from crystalline state. J Mol Biol 2007, 372, (3), 774-97. 47. Spiller, B.; Gershenson, A.; Arnold, F. H.; Stevens, R. C., A structural view of evolutionary divergence. Proceedings of the National Academy of Sciences 1999, 96, (22), 12305-12310. 48. Sussman, J. L.; Harel, M.; Frolow, F.; Oefner, C.; Goldman, A.; Toker, L.; Silman, I., Atomic structure of acetylcholinesterase from Torpedo californica: a prototypic acetylcholine-binding protein. Science 1991, 253, (5022), 872-9.

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An Esterase from Anaerobic Clostridium hathewayi Can Hydrolyze

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