Technical Comments

Response to Comment on “A bacterium that degrades and assimilates poly(ethylene terephthalate)”

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Science  19 Aug 2016:
Vol. 353, Issue 6301, pp. 759
DOI: 10.1126/science.aaf8625


Yang et al. suggest that the use of low-crystallinity poly(ethylene terephthalate) (PET) exaggerates our results. However, the primary focus of our study was identifying an organism capable of the biological degradation and assimilation of PET, regardless of its crystallinity. We provide additional PET depolymerization data that further support several other lines of data showing PET assimilation by growing cells of Ideonella sakaiensis.

We appreciate the Comment by Yang et al. (1) and are grateful for this opportunity to explain the context of how our work builds on previous studies. The intent of our study was to isolate and describe a microorganism that can degrade and assimilate poly(ethylene terephthalate) (PET) (2). Therefore, we only cited the pioneering works that report specific microorganisms able to grow on PET (3, 4). In a previous study, Sharon and Sharon (5) confirmed microbial PET degradation by semi-isolated microorganisms, where the involvement of Nocardia species was implied by microscopic observation. However, the contribution of other microorganisms was not ruled out, nor was the possibility that the PET was degraded by the resting cells harboring PET-hydrolyzable enzymes. We identified the PET hydrolase (PETase) gene based on amino acid sequence identity with a known hydrolase from Thermobifida fusca (TfH) that exhibited PET-hydrolyzing activity (6), similar to other PET-hydrolyzing enzymes (710). We further referred cutinase from Humicola insolens (HiC) (11) and seven other enzymes in the supplementary materials [table S2 in (2)]. The functions of these enzymes, particularly TfH, LCC (9), and FsC (10), greatly contributed to the understanding of PETase enzymology.

With the intention of isolating a specific microorganism that can use PET for growth, even if the PET is mostly in the amorphous form, the extent of crystallinity is of secondary importance. We described the motivation for using low-crystallinity (1.9%) PET and the observation that the structure of crystallized PET hampered the enzymatic hydrolysis of its ester linkages (8, 12) in our Report.

Based on the proton nuclear magnetic resonance and gel permeation chromatography profiles of the degraded PET film, we concluded that the degradation by the consortium termed “no. 46” proceeded from the PET surface (2). To support this inference, we analyzed the surface of the PET film using x-ray photoelectron spectroscopy (XPS). The XPS spectrum for degraded PET film by no. 46 showed that a new peak appeared at 0.6 eV higher in binding energy than that observed for the O1s peak of C–O–C linkage (532.5 eV), indicating the formation of the C–O–H linkage by hydrolysis of the ester linkage, which should either be of hydroxyl or carboxyl groups. To accurately determine the surface functional groups, each group was labeled with heptafluorobutyryl chloride (for –OH) and 1,1-carbonyldiimidazole (for –COOH) (13) and analyzed by XPS. Consequently, the peaks of F1S (687.5 eV) and N1S (398.5 eV) after degradation dramatically increased (Fig. 1A). The values of surface hydroxyl and carboxyl groups for the degraded film were calculated to be 1.09 ± 0.03 × 1014 and 7.11 ± 0.92 × 1014 groups per cm2, respectively; those for the control film were 0.13 ± 0.11 × 1014 and 0.36 ± 0.10 × 1014 groups per cm2, respectively (Fig. 1B). The increase in the surface functional groups strongly suggests an endo-type scission of the polymer chain. In addition, the surface of the PET film cultured with Ideonella sakaiensis was qualitatively analyzed. Reactive Black 5 staining (14) revealed an increased number of –OH groups (Fig. 2A), while Alexa Fluor 488-hydrazide labeling revealed an increased number of –COOH groups (Fig. 2B). The surface was not stained with Coomassie brilliant blue R250 (Fig. 2C). These results indicate that the increase in these functional groups was due to PET hydrolysis in an endo-type manner and not due to protein contamination.

Fig. 1 XPS analyses of PET films degraded by consortium no. 46.

No. 46 was cultured with PET film (20 by 15 by 0.2 mm) in MLE (modified lettuce and egg) medium at 30°C for 22 days with a biweekly medium change. The degraded PET film was washed with 1% SDS, distilled water, and then ethanol, followed by air drying. The hydroxyl and carboxyl groups on the film surface were labeled with heptafluorobutyryl chloride and 1,1′-carbonyldiimidazole, respectively (13), and independently quantified by XPS on a JEOL JPS-9010MC/SP photoelectron spectrometer (Tokyo, Japan) with a 100 W MgKα x-ray source (λ 9.889 Å, 1253.5 eV) with take-off angle of 45°. (A) Narrow spectra of C1s, N1s, and F1s regions. The spectra were obtained by accumulation of 15 scans at intervals of 0.1 eV. (B) Density of surface functional groups. The atomic compositions were calculated on the software equipped. At the take-off angle, photoelectrons ~3 nm in depth were assumed to be detected. The surface functional density (groups per cm2) was calculated with PET density of 1.375 g/cm3.

Fig. 2 Qualitative surface analyses of PET films degraded by I. sakaiensis.

I. sakaiensis was cultured with PET film (20 by 15 by 0.2 mm) in YSV medium at 30°C for 7 days. Intact PET film (control) and PET film after cultivation (7 days) were washed with 1% SDS, distilled water, and then ethanol, followed by air drying. (A) Detection of the hydroxyl groups. The film was stained with Reactive Black 5 following the reported manuscript (14). (B) Detection of the carboxyl groups. The film was soaked in 0.18% (w/v) Alexa Fluor 488-hydrazide, and then 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide was added to the solution (final concentration, 0.25% w/v) for activating carboxyl groups. After 24 hours in the dark, the film was washed with distilled water and then ethanol, followed by air drying. The surface was observed by fluorescence microscope at λ excitation = 493 nm, λ emission = 517 nm. Scale bar, 100 μm. (C) Detection of the proteins. The film was stained with 0.025% (w/v) Coomassie brilliant blue R250 in 20% (v/v) acetic acid and then destained with 10% (v/v) acetic acid.

The isotopic carbon tracing experiments are a direct method for tracing carbon among nutrients. We demonstrated PET assimilation using multiple approaches instead, because isotope-labeled PET was commercially unavailable. As shown in figure S9 in (2), the growth of I. sakaiensis in the PET film–fed YSV (yeast extract–sodium carbonate–vitamins) medium reached an average optical density of 0.88 at 600 nm for 151 hours (the rightmost panel), whereas that in the absence of PET stopped at ~0.06 (the leftmost panel). During this period of time, the weight loss of the PET film was clearly observed [figure 1H and figure S8 in (2)]. These results indicate that a part of the degraded PET was assimilated by I. sakaiensis cells. Furthermore, in the culture fluid, the amount of enzymatically released compounds from the PET film—such as terephthalic acid (TPA) and mono(2-hydroxyethyl) terephthalic acid (MHET)—was markedly lower than that of the TPA or MHET units in the degraded PET, indicating that they were incorporated into the cells for growth. By culturing I. sakaiensis cells with the PET film, the expression of genes for PETase, MHET hydrolase (MHETase), and TPA degradation pathway, involved in the PET degradation, was found to be dramatically up-regulated, indicating that PET was actively metabolized for energy acquisition and cell proliferation.


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