Biodegradation of Synthetic Aliphatic-Aromatic Polyesters in Soils: Linking Chemical Structure to Biodegradability

Abstract

Biodegradable aliphatic-aromatic copolyesters are commercially important and used in diverse applications, including soil-biodegradable mulch films. This work investigates the effects of copolyester chemical structure on soil biodegradability using ¹³C-labeled variants of polybutylene adipate-co-terephthalate (PBAT) and polybutylene sebacate-co-terephthalate (PBSeT), as well as ¹³C-labeled cellulose as a positive biodegradation control. Biodegradation in soil was assessed by monitoring polyester and cellulose mineralization to ¹³CO₂ throughout multimonth incubations and quantifying the nonmineralized polyester in the soil after incubations. Mass balances on polyester- and cellulose-added ¹³C over the incubations were closed. The soil biodegradability of PBSeT was higher than that of PBAT for variants with a molar ratio of terephthalate (T) to total diacid of 50%. PBAT biodegradability substantially increased as its T content decreased from 50% to 47, 30, 20, and 0%. Increasing biodegradability with decreasing T content also resulted in preferential biodegradation of aliphatic-rich domains in the initial phase of the incubations. Polyesters undergoing extensive mineralization also showed substantial incorporation of polyester carbon into the soil microbial biomass. Differences in polyester soil biodegradability were rationalized based on differences in their enzymatic hydrolyzability. Qualitative chemical structure-biodegradability relationships can lead to tailoring polyester biodegradability to specific applications and the biodegradation potentials of receiving environments.

Introduction

Replacing conventional, environmentally persistent polymers such as polyethylene (PE) with environmentally biodegradable polymers in specific applications is a promising strategy to prevent plastic accumulation in the environment. − This is especially true for applications in which plastic products are directly used in the open environment and/or have a high probability of remaining (in part) in the open environment after use. An example is thin (<25 μm thickness) agricultural mulch films: conventional films are PE-based and, if not completely recollected after use, accumulate in agricultural soils over repeated application cycles to concentrations that negatively impact soil productivity. − Conversely, mulch films composed of biodegradable polymers undergo biodegradation by soil microorganisms after being plowed into soils after the application period. − Under oxic conditions, this process ultimately results in the complete metabolic utilization of polymer carbon to form both CO₂ (i.e., mineralization) and new microbial biomass. Polymer-derived carbon incorporated into microbial biomassas part of the soil organic matter poolalso undergoes biodegradation (possibly over longer time frames than the polymer itself), designating CO₂ as the ultimate end point of biodegradation. Therefore, biodegradation in soils is typically assessed by soil incubations coupled to respirometric analysis of the formation of polymer-derived CO₂ over time.

Among the biodegradable polymers with desirable material properties for mulch film applications, and at the same time documented biodegradability in soils, are aliphatic-aromatic copolyesters, including polybutylene adipate-co-terephthalate (PBAT) and polybutylene sebacate-co-terephthalate (PBSeT). As a copolyester synthesized from two aliphatic (i.e., the diol butanediol (B) and either the diacid adipate (A) or sebacate (Se)) and one aromatic (i.e., the diacid terephthalate (T)) monomer units, their biodegradability (as well as key material properties) can be adjusted by varying the ratio of the respective monomers. A first possibility, as documented for PBAT, is to alter the aromatic diacid T to aliphatic diacid A ratio: increasing the T-to-A ratio decreases PBAT enzymatic hydrolyzability, − as well as PBAT soil biodegradability, as determined indirectly by following the gravimetric weight loss of incubated polyester specimens over time. − Consistently, enzymatic hydrolysis of PBAT was also found to result in an increase in the T content of the residual (nonhydrolyzed) PBAT. The decrease in enzymatic hydrolyzability and biodegradability rates with increasing T content has been ascribed to a relative increase in T-enriched crystallites in PBAT, which undergo slower enzymatic hydrolysis as compared to A-enriched amorphous domains of the polyesters. Furthermore, transitional increases in bulk crystallinities and crystallite sizes of PBAT have been observed during biodegradation. , A second possibility to alter the biodegradability of polyesters is to alter the aliphatic diacid, for instance by using Se instead of A. While the biodegradability of polyesters depends on the chemical nature of the monomeric building blocks and not on their feedstock origin, it is noteworthy that Se is also available from biobased feedstock. The resulting partially biobased character of PBSeT is attractive in terms of reducing dependence on fossil feedstocks in a circular polymer economy.

While the effects of aliphatic-aromatic polyester chemical structure on biodegradability have long been recognized, , studies that determine chemical structure-soil biodegradability relationships for a larger set of these polyesters that systematically differ in their monomeric composition remain forthcoming. For such studies, it is desirable not only to selectively follow polyester mineralization to CO₂ over time but also to independently verify closed mass balances on initially added polyester carbon over the course of the soil incubations and to quantify potentially present residual biodegradable polyesters at the end of the soil incubations. These two additional measurements allow one to also estimate the amount of polyester carbon incorporated into microbial biomass as the difference between the total nonmineralized carbon and the amount of residual polyest carbon. Closing mass balances on polyester carbon is, however, difficult when using nonlabeled polyesters, as polyester carbon incorporated into biomass cannot be delineated from that of background soil organic matter.

We recently demonstrated the utility of ¹³C-labeled polyesters (with the exemplary polybutylene succinate (PBS)) for quantifying the fraction of polyester that was mineralized to ¹³CO₂ over the course of the soil incubations using online cavity ring-down spectroscopic (CRDS) detection and to close mass balances on the amount of nonmineralized ¹³C-label that remained in the soil matrix, either as residual polyester or incorporated into biomass/soil organic matter. Total nonmineralized polyester-added ¹³C was quantified by combusting small soil aliquots in an elemental analyzer coupled to isotope-ratio mass spectrometry (EA-IRMS), while solvent extraction of the soil at the end of the incubation coupled to quantitative ¹H-nuclear magnetic resonance (NMR) spectroscopy allowed us to quantify the residual PBS remaining in the soil. , When combined, these approaches provide detailed information on polyester biodegradation rates and extents and insights into how polyester carbon is microbially utilized in soil.

The objective of the present study was to assess the effect of polyester chemical structure on soil biodegradability for a set of aromatic–aliphatic polyesters that systematically varied in their chemical composition. To this end, we conducted soil incubations with ¹³C-labeled variants of PBAT and PBSeT, both at the same molar ratio of T to total diacids of 50% (i.e., T/(T+A) and T/(T+Se))to assess the effect of the aliphatic diacid chemistry on biodegradabilityand a set of butanediol-¹³C₄ variants of PBAT that differed in their molar T to total diacid contents of 47%, 30%, 20%, or 0% (i.e., the aliphatic polyester PBA)to assess the effect of T content on polyester soil biodegradability. For the PBAT and PBSeT comparison, we tested a total of five labeled variants (i.e., three PBAT variants ¹³C-labeled either in the monomers B, T, or A, and two PBSeT variants ¹³C-labeled in the monomers B and T; a variant labeled in Se was not included as the respective labeled monomer was not commercially available) to also assess potential differences in the biodegradation dynamics of the individual monomeric building blocks in these polyesters. The polymerization was restricted to small-scale laboratory reactors (due to the costs of ¹³C-labeled monomers) and resulted in polyester molecular weights lower than those typically achieved by synthesis at the industrial scale and applied in commercial products, such as certified soil-biodegradable mulch films. At the same time, the molecular weight characteristics were consistent between the polyester variants tested herein, thereby allowing the assessment of the effect of polyester monomer composition on soil biodegradability. All polyesters were incubated in an agricultural soil for up to 425 days. During soil incubation, we continuously monitored the mineralization of polyester-¹³C to ¹³CO₂, followed by quantifying the nonmineralized polyester-added ¹³C, as well as solvent-extracting and quantifying residual polyesters in the soil. We additionally performed incubations of ¹³C-labeled cellulose as a positive biodegradation control in the same soil. Cellulose incubations were conducted at three different added masses (one lower, one matching, and one higher amount than that of the polyesters added) to test for potential decreases in biodegradation rates with increasing mass loadings, which could indicate nutrient limitations during biodegradation (e.g., insufficient nitrogen and phosphorus supply to cellulose-biodegrading soil microorganisms).

Materials and Methods

Materials

Chemicals

HPLC-grade chloroform (CHCl₃) and methanol (MeOH) were purchased from Fischer. Deuterated chloroform (CDCl₃; 99.8% atom D) was from Amar, and 1,4-dimethoxybenzene (DMB; >99%) was from TCI. Butane-1,4-diol (B), ¹³C₄-butane-1,4-diol, adipic acid (A) (99 atom % ¹³C), 1,6-¹³C₂-adipic acid, sebacic acid (Se) (99 atom % ¹³C), terephthalic acid (T), and 1-¹³C₁-terephthalic acid (99 atom % ¹³C) were from Sigma. CHCl₃ and CDCl₃ were stored with 4 Å molecular sieves to remove water. All other chemicals were used as received.

Polyesters

The ¹³C-labeled PBA_100‑X T _X and PBSe₅₀T₅₀ variants were synthesized from the respective monomers according to established polycondensation protocols. Herein, X refers to the percent contribution of T to the total diacid monomeric units in the polyester (Figure ). The first set of polyester variants were ¹³C-labeled either in B (i.e., P­(¹³C₄-B)­A₅₀T₅₀ and P­(¹³C₄-B)­Se₅₀T₅₀), in T (i.e., PBA₅₀(1-¹³C₁-T)₅₀ and PBSe₅₀(1-¹³C₁-T)₅₀), or in A (i.e., PB­(1,6-¹³C₂-A)₅₀T₅₀). In the second PBAT series, which varied in T content (and thus X), all variants were ¹³C-labeled in B (i.e., P­(¹³C₄-B)­A₁₀₀T₀, P­(¹³C₄-B)­A₈₀T₂₀), P­(¹³C₄-B)­A₇₀T₃₀, and P­(¹³C₄-B)­A₅₃T₄₇). Based on the fraction of carbon in the respective monomeric unit selected for labeling to the total carbon in the polyester and the abundance of ¹³C to total C in the labeled monomer used in the synthesis, the ratio of ¹³C-labeled to nonlabeled monomers was adjusted in the synthesis to ensure that the overall ¹³C content in all polyesters was comparable and in the range of 3.5–4.0 atom %. These values and other key physicochemical properties of the polyesters are provided in Section S1. Consistent conditions during the small-scale synthesis of the polyester variants resulted in similar molecular weight characteristics among the variants. We note, however, that during the syntheses, increases in the viscosity of the polyester melts ultimately caused the magnetic stirring in the small reactors to stop, resulting in the polyesters having lower average molecular weights and broader molecular weight distributions than polyesters synthesized at industrial scale and commonly used in commercial products such as certified soil-biodegradable mulch films. Following synthesis, each polyester variant was cryomilled to a powder, which was sieved to collect the fraction with particle diameters between 100 and 300 μm for soil incubations.

1.

Chemical structures of ¹³C-labeled polyesters and cellulose used in soil incubations. Seven variants of monomer-specific ¹³C-labeled polybutylene adipate-co-terephthalate (PBAT) and two variants of polybutylene sebacate-co-terephthalate (PBSeT) were used, as shown. Uniformly and fully (≥97%) ¹³C-labeled celluloses were mixed with varying amounts of nonlabeled cellulose (see Materials and Methods text for details).

Cellulose

Nonlabeled cellulose (Sigma-Aldrich) and uniformly ¹³C-labeled cellulose (U-¹³C-cellulose; labeling extent of ≥97%; IsoLife, Netherlands) were added at defined ratios to Eppendorf tubes, followed by homogenization of the cellulose mixtures using bead beating with zirconia beads.

Soil

The soil for incubations was collected from the Agricultural Center Limburgerhof (Germany) and was used as previously described (i.e., the soil was air-dried, sieved to 2 mm, stored at 4 °C, and, prior to use in the incubations, the water content was adjusted to 45% of its maximum water-holding capacity (WHC_max = 37.3 g H₂O 100 g^–1 dry soil) with Milli-Q water (resistivity ≥ 18.2 MΩ cm, TOC ≤ 5 ppb)). The soil was a sandy clay loam according to the USDA soil texture classification system (i.e., particle size distribution by mass of 54.9% in the sand (diameters 50–2 mm), 12.3% in the silt (i.e., 2–50 μm), and 30.8% in the clay fraction (<2 μm)). The organic carbon and total nitrogen contents were 1.14% and 0.11% by weight, respectively.

Measurement of Polyester- and Cellulose-Derived ¹³CO₂, ¹³C_mineralized

Polyester and cellulose mineralization to ¹³CO₂ was quantified using an automated soil incubation system described previously with up to 36 incubation bottles (250 mL), each holding 100 g of soil (dry weight equivalent) at 25.0 ± 0.2 °C. The headspace gas in each incubation bottle was continuously exchanged during respirometric analysis at a volumetric flow rate of 24 mL min^–1. For equilibration, bottles containing soils were run on the system for at least 7 days prior to polyester or cellulose additions. For polyester and cellulose addition, we mixed either ¹³C-labeled polyesters (100 mg per bottle; resulting in an initial polyester concentration of 1 mg polyester g^–1 dry weight soil) or varying amounts of cellulose, each containing 10 mg of U-¹³C-cellulose (i.e., either only 10 mg ¹³C-cellulose or 10 mg ¹³C-cellulose with either 90 or 490 mg of nonlabeled cellulose; resulting in total added cellulose masses of 10, 100, and 500 mg, respectively). The polyester and cellulose particles were added stepwise to the top of the soil in the incubation bottle, followed by carefully mixing in the particles with a metal spatula to achieve an even distribution in the soil. We note that the experimental data for 100 mg cellulose has already been published in a previous study.

Triplicates of monomer-specific ¹³C-labeled PBAT and PBSeT were incubated in soils for 319 days, at which time we removed one replicate per variant for analyses of nonmineralized polyester carbon, while the remaining duplicate bottles were incubated up to 425 days. Similarly, cellulose incubations were run in triplicate for 139 days, when one bottle was removed, and incubation of the remaining two bottles continued up to 254 days. Incubations of PBAT variants with different T contents were run in triplicate for 145 days. The termination times for the incubations described above were chosen by balancing the amount of data collected vs freeing capacity on the incubation systems. In all cases, the formation of polyester-derived ¹³CO₂ was still observed when stopping the incubations. The water content of the bottles was monitored gravimetrically and was periodically readjusted using double-deionized water, thereby ensuring that water contents were maintained between 42 and 45% of WHC_max.

The efflux gas from one incubation bottle at a time was directed to an isotope-specific cavity ring-down spectroscopy analyzer (CRDS; Picarro model G2201-i) for the quantification of formed ¹³CO₂ and ¹²CO₂. Incubation bottles were measured more frequently in the beginning of the incubations to capture the faster initial mineralization rates. Afterward, bottles were periodically analyzed but stored closed at 25 °C in the dark. We periodically also ran three synthetic air standards containing known concentrations of ¹²CO₂ and ¹³CO₂ (Pangas; [CO₂]_total = 400, 500, 700 ppm (±1% relative to concentration) and δ¹³CO₂ = −5.27, −10.76, −10.21 ‰ (respectively, as quantified by gas-chromatography isotope-ratio mass spectrometry (GC-IRMS)) to the CRDS to correct for minor instrument drift. Each measurement period of a given incubation bottle was 10 min, with a measurement frequency of about 1 s^–1. Data from the final 3 min in this interval were time-averaged to obtain ¹²CO₂ and ¹³CO₂ concentrations per measurement period. Calculations used to determine polyester and cellulose mineralization rates and extents are given in Section S2. The mineralization extents, ¹³C_mineralized, are expressed as polyester- and cellulose-derived ¹³CO₂ formed as a percent of polyester- and cellulose-¹³C added to the soil at the onset of the incubation.

Measurement of Total Nonmineralized Polyester- and Cellulose-Added ¹³C Remaining in Soils, ¹³C_{nonmineralized}

After the incubations were terminated, all bottles were transferred to a −20 °C freezer, followed by lyophilization at 0.01 mbar for at least 24 h. The dried soils were passed through a 2 mm sieve, followed by milling approximately half of each dried soil in a vibratory disk mill (RS1, Retsch). To finely disperse residual polyester in soil prior to EA-IRMS analysis, we treated a 5 g subsample of each milled soil with chloroform-sonication. A polyester-free control soil was treated identically. Details are provided in Section S3-1, including chloroform removal. The chloroform-sonication step was omitted for the soils with cellulose, as it is not soluble in chloroform. A 10 mg subaliquot of each milled and solvent-treated soil for polyester-containing and polyester-free incubations and of the milled soils for cellulose was then weighed into a tin capsule for EA-IRMS analysis.

The tin capsules containing samples were placed on an elemental analyzer (EA) (Thermo Fisher FlashEA 1112) coupled to a continuous flow interface (Thermo Fisher Conflo IV) and an isotope-ratio mass spectrometer (IRMS) (Thermo Fisher Delta V Plus). The EA-IRMS was operated and calibrated as described previously. We calculated the carbon contents (%C) and δ¹³C values of the combusted soil samples using calibration curves constructed from measurements of various organic compounds with known carbon contents and δ¹³C values, as well as of ¹³C-enriched glucose standards (details in Section S3-2). We converted the determined sample δ¹³C values to ¹³C atom% (see Section S1). The nonmineralized polyester- and cellulose-added ¹³C amounts, ¹³C_{nonmineralized}, were calculated as detailed in Section S3 and are expressed as a percent of polyester- or cellulose-¹³C added to the soil at the onset of the incubation.

Measurement of Residual Polyester in Soils, ¹³C_{polymer residual}

Analyses followed extraction procedures previously published. , In brief, 3 g of dried and sieved soil from each polyester incubation were transferred into cellulose thimbles, which were placed in Soxhlet extractors (ChemGlass; body volume = 6 mL). The soils were extracted with a 90:10 vol % chloroform-methanol (CHCl₃:MeOH; total of 10 mL) solvent mixture for 8 h. The solvent was subsequently removed under a stream of compressed air, followed by reconstitution of the dried extract in 2 mL of CDCl₃ containing a known amount of DMB as an internal standard. The samples were analyzed on a Bruker Avance III 400 MHz NMR spectrometer equipped with a 5 mm BBFO Z-Gradient probe, with settings specified in Section S4. We calculated the extracted polyester mass in each sample as detailed in Section S4, accounting, if occurring, for changes in its chemical composition (i.e., a higher ratio of T to total diacid than for the initially added polyester). We multiplied the extracted polyester mass by the mass ratio of soil in the incubation to the soil extracted and then expressed the residual polyester mass in the soil as a percent of the initially added polyester mass, ¹³C_{polymer residual}. Exemplary ¹H NMR spectra of different polyester chemical structures and DMB, along with details and results of spike-recovery experiments to validate the extraction procedure, are given in Section S4.

Results and Discussion

Mineralization of ¹³C-Labeled Polyesters and Cellulose in Soil

PBA₅₀T₅₀ and PBSe₅₀T₅₀

Figure shows ¹³C-mineralization rates (panels a, c) and corresponding calculated cumulative mineralization extents (panels b, d) for the three monomer-specific ¹³C-labeled PBA₅₀T₅₀ (panels a, b) and the two monomer-specific ¹³C-labeled PBSe₅₀T₅₀ variants (panels c, d). Within a few hours of being added to the soil, the PBA₅₀T₅₀ and PBSe₅₀T₅₀ variants were mineralized to ¹³CO₂ (Figure a, c; insets). The three variants labeled in B and A (i.e., P­(¹³C₄-B)­A₅₀T₅₀, P­(¹³C₄-B)­Se₅₀T₅₀, and PB­(1,6-¹³C₂-A)₅₀T₅₀) exhibited first maxima in ¹³C-mineralization rates at around six h of incubation. The initial rate maxima of the two variants labeled in T (i.e., PBA₅₀(1-¹³C₁-T)₅₀ and PBSe₅₀(1-¹³C₁-T)₅₀) were slightly delayed to 18 h of incubation. Following these initial maxima, mineralization rates of all variants decreased to low values after approximately 24 h. At this time, ¹³C_mineralized amounted to only small values of 0.5 to 1.25% of the initially added polyester-¹³C (Figure b, d; insets). We ascribe these initial mineralization maxima to the microbial utilization of residual labeled monomers and short (possibly cyclic) labeled oligomers that readily diffused out of the bulk at the onset of the incubation and that were sufficiently small to not require enzymatic hydrolysis before being taken up and metabolized by microbes. The presence of such monomers and oligomers at low mass percentages in polyesters synthesized through polycondensation is well established. This explanation is consistent with the differences in the occurrence of the initial rate maxima, as these matched the differences in mineralization dynamics of the labeled monomers (i.e., ¹³C₄-B, 1,6-¹³C₂-A and 1-¹³C₁-T) when added directly to soil, as previously reported.

2.

Mineralization rates and cumulative extents of ¹³C-labeled polybutylene adipate-co-terephthalate (PBAT) and polybutylene sebacate-co-terephthalate (PBSeT) variants and of ¹³C-labeled cellulose in soil. Panels (a, b): monomer-specific ¹³C-labeled PBA₅₀T₅₀ variants. Panels (c,d) monomer-specific ¹³C-labeled PBSe₅₀T₅₀ variants. Panels (e, f): butanediol-¹³C-labeled PBA_100‑X T _X variants with varying molar terephthalate (T) to total diacid contents (i.e., X %). All polyester incubations contained 100 mg of the respective variant in 100 g of soil (dry weight equivalents). Panels (g, h): ¹³C-labeled cellulose added in different total amounts of 10, 100, and 500 mg, by mixing 10 mg labeled cellulose with different amounts of nonlabeled cellulose, each in 100 g of soil. Panels b, d, f, and h show cumulative polymer-¹³C mineralization extents (calculated by integration of mineralization rates in panels a, c, e, and g, respectively) expressed as mass percent of polyester- or cellulose-¹³C initially added to soil. Samples marked with an asterisk (*) correspond to those with higher mineralization extents compared with the other replicates of the same polyester variant. For panels (a–d): triplicate soil incubations were run for each polyester variant up to 319 days, when one incubation bottle per variant was terminated, whereas the incubation of the remaining duplicate bottles was continued up to 425 days. For panels (e, f): triplicate incubations were run for each variant for 145 days. For panels (g, h): triplicate incubations were run for each cellulose loading up to 139 days, when one incubation bottle at each mass loading was terminated, and the incubation of the remaining duplicate bottles at each mass loading was continued up to 254 days. Measured points are plotted for each individual replicate, and dotted lines represent linear interpolations between measurement points for those timeframes when bottles continued to be incubated under the same conditions but were not connected to the ¹³CO₂ detection unit. The cellulose mineralization data at 100 mg has been replotted here from an earlier publication for comparison.

Following this initial mineralization phase, mineralization rates slowly reincreased to second maxima of around 0.05 to 0.11 μg ¹³C h^–1 between 100 and 150 days for PBA₅₀T₅₀ variants and 0.35 to 0.38 μg ¹³C h^–1 between 15 and 40 days for the PBSe₅₀T₅₀ variants. Following these maxima, polyester mineralization rates decreased in all incubations to low but above-background values determined from control bottles with only soil until the incubations were terminated. Integration of the ¹³C-mineralization rates of duplicates incubated up to 425 days yielded cumulative mineralization extents of ¹³C_mineralized = 26 (±1)%, 25 (±2)%, and 18 (±1)% for PBA₅₀(1-¹³C₁-T)₅₀, PB­(1,6-¹³C₂-A)₅₀T₅₀, and P­(¹³C₄-B)­A₅₀T₅₀, respectively, and 58 (±2)% and 52 (±1)% for P­(¹³C₄-B)­Se₅₀T₅₀ and PBSe₅₀(1-¹³C₁-T)₅₀, respectively (n = 2 per polyester variant, average ± range of two bottles each; note that one bottle of each set of triplicate bottles was already removed after 319 days for quantifying the residual polyester amount and determining the ¹³C mass balance at this intermediate incubation time). This continuous mineralization up to 425 days with a second rate maxima between 100 and 150 days corresponded to the biodegradation of the bulk polyester.

One incubation of each of the above polyester variants was terminated early, after 319 days, for intermediate mass balance assessment and quantification of residual polyester mass (see below). Among these was one incubation bottle of PB­(1,6-¹³C₂-A)₅₀T₅₀ and one of P­(¹³C₄-B)­Se₅₀T₅₀ with “sudden” enhanced mineralization rates at about 80 days of incubation (marked with asterisks (*) in Figure a–d). These two bottles exhibited higher final mineralization extents of ¹³C_mineralized = 86% and 67%, respectively, compared to the other two bottles of the same variant (Figure b,d). Potential causation for this enhanced biodegradation will be discussed below.

PBA_100‑X T _X with Varying T Content

The mineralization rates and extents of the ¹³C-B-labeled PBA_100‑X T _X variants with varying T content are shown in Figure e,f. The variant with the highest tested T content, PBA₅₃T₄₇, showed the lowest mineralization rates and final extents (i.e., 13 (±1) % (n = 3) after 145 days of incubation; a zoom-in on the data better showing low mineralization rates (≤0.3 μg ¹³C h^–1) is provided in Section S5). The mineralization rate and extent of the PBA₅₃T₄₇ variant were higher than those of PBA₅₀T₅₀ discussed above (i.e., 7 (±1) % after 145 days (n = 3); Figure b). This finding suggests that PBAT soil biodegradability is highly dependent on the T content at around 50%. Consistently, the PBA_100‑X T _X variants with lower T contents showed higher mineralization rates and extentswhich increased with decreasing T contentafter 145 days of incubation: 75 (±1) %, 78 (±6) %, and 82 (±2) % for PBA₇₀T₃₀, PBA₈₀T₂₀, and PBA₁₀₀T₀, respectively (n = 3 per polyester variant). While decreases in the T contents from 30 to 0% thus increased soil biodegradability, the comparatively large difference in mineralization between the PBA₇₀T₃₀ and PBA₅₃T₄₇ variants supports that the soil biodegradability of PBAT is strongly dependent on T contents between 30 and 47%. As previously reported in the literature, in this T content range, the crystalline domains are formed from B-T units, while for PBAT with lower T contents, these domains are formed from B-A segments.

Cellulose

Cellulose mineralization rates increased continuously from the onset of the incubations up to approximately 2 to 3 days, when rates reached maxima of 21 (±3), 26 (±3), and 35 (±8) μg ¹³C h^–1 in bottles with 10, 100, and 500 mg cellulose, respectively (n = 3 per cellulose amount; see Figure g, inset). This increase in maximum ¹³C-mineralization rates scaled linearly with the total cellulose ¹³C added (i.e., 10 mg of U-¹³C-labeled cellulose plus approximately 1.1% natural abundance of ¹³C in the 90 and 490 mg of nonlabeled cellulose added) (see Section S6), suggesting similar mineralization dynamics of the labeled and nonlabeled celluloses. Following the initial maxima, the mineralization rates substantially decreased over the next 4 weeks to relatively low values. The secondary, smaller peaks in mineralization rates in the bottles with 500 mg of cellulose after about 50 days of incubation coincided with the addition of water to these soils to maintain a stable soil water content throughout the incubation. The data suggest that the sensitivity of cellulose mineralization rates to soil water content was higher with increasing amounts of cellulose in the soil.

The cumulative mineralization extents showed biphasic behaviors comparable to those for each cellulose treatment (Figure h). The first phase corresponded to mineralization of approximately 60% of the added cellulose-¹³C over 10 days, and the second phase was characterized by a flattened slope in the mineralization extent curves. After 254 days, the final mineralization extents were ¹³C_mineralized = 78 (±1) %, 75 (±1) %, and 82 (±1) % for incubations with 10, 100, and 500 mg of cellulose added, respectively (n = 2, average ± range of two bottles each; note that one bottle of each set of triplicate bottles was already removed after 139 days for determining the cellulose ¹³C mass balance at an intermediate incubation time). Biphasic mineralization of cellulose in soil is consistent with previous reports. ,

We ascribe the initial phase with high mineralization rates to fast and extensive microbial utilization of cellulose both for energy generation (respiratory production of CO₂) and for biomass buildup. The onset of the second phase then occurred following consumption of most of the added cellulose and was due to slower turnover of microbial bio­(necro)­mass containing cellulose-derived ¹³C. In this case, soil microorganisms had comparable metabolic utilization patterns of the cellulose at the three mass loadings, with approximately 60% of the cellulose carbon being mineralized to CO₂ and 40% of the carbon being incorporated. This carbon use efficiency of approximately 40% is fully consistent with previous studies and theoretical considerations, although not experimentally proven in our experiments due to solvent extraction and thus subsequent quantification of residual cellulose being impossible. − More importantly, comparable mineralization dynamics at the three different cellulose mass loadings strongly suggest that substrate concentrations used in our incubations were sufficiently low to ensure that microbial biodegradation of cellulose and polyesters was not limited by the supply of nitrogen or phosphorus to the cells (which would have slowed biodegradation at high mass loadings). The possibility to run incubations at such low polyester and cellulose concentrations (due to high signal-to-noise when quantifying ¹³C) is a unique advantage of using ¹³C-isotopically labeled instead of nonlabeled polymers.

Closing Mass Balances on Polyester- and Cellulose-Added ¹³C over the Course of the Incubations

Figure (left side) shows the total amount of nonmineralized polyester- and cellulose-added ¹³C remaining in the soil at the end of the incubations (¹³C_{nonmineralized;} expressed in % of polyester- or cellulose-¹³C initially added to the soils), as quantified by EA-IRMS on small soil aliquots. Figure (right side) also shows the corresponding final mineralization extents (¹³C_mineralized) replotted from Figure . Exact ¹³C_{nonmineralized} and ¹³C_mineralized values are also provided in Section S7.

3.

Quantification and characterization of nonmineralized polymer-added ¹³C remaining in soils after incubation. Bars on the left represent polyester- and cellulose-added ¹³C that did not mineralize over the course of the incubation and thus remained in the soils, ¹³C_{nonmineralized} (expressed as a percent of polyester- and cellulose-added ¹³C initially added to the soil). Bars on the right represent cumulative final amounts of polyester-¹³C mineralized (¹³C_mineralized; expressed as a percent of polyester- and cellulose-added ¹³C initially added to the soil) in each bottle at the end of the incubations (respective incubation times are indicated to the left of the box; this data corresponds to the final mineralized amounts of polyesters and cellulose shown in Figure ; samples marked with an asterisk (*) correspond to those with higher mineralization extents compared to the other replicates of the same polyester variant). The mass balance on both polyester- and cellulose-added ¹³C was closed, as indicated by the good agreement between the sum of ¹³C_{nonmineralized} and ¹³C_mineralized with the initially added amounts of ¹³C (mass balances in percent of polyester- and cellulose-added ¹³C initially added to the soil to the right of the plot (average ± standard deviation of triplicate incubation bottles)). The orange vertical lines plotted on the left side for each incubation bottle represent the quantified amounts of polyester carbon that remained in the soil, ¹³C_{polymer residual}, as determined by Soxhlet extraction of soil aliquots followed by ¹H NMR spectroscopy analysis (also expressed as a mass % of added polyester carbon, converted to polyester-¹³C added to the soil at the onset of the incubation). The data for 100 mg of cellulose has been reported previously.

As expected, ¹³C_{nonmineralized} of both the polyester- and cellulose-added ¹³C in the soil decreased with increasing mineralization extents for the respective incubation. More importantly, the sum of ¹³C_{nonmineralized} and ¹³C_mineralized corresponded to, on average, 97–105% of the initially added ¹³C (averages of triplicate incubations per polyester and cellulose variant). This finding implies closed mass balances on polyester- and cellulose-added ¹³C over the course of the incubations, including for the two incubation bottles of PB­(1,6-¹³C₂-A)­T and P­(¹³C₄-B)­SeT with higher mineralization extents (marked with asterisks (*) in Figure ), thereby confirming that these incubations indeed showed enhanced biodegradation. Accurate quantification and tracking of polymer-added ¹³C and the capability to close mass balances on polymer-¹³C over the course of long-term (soil) incubations is another advantage of using ¹³C-labeled polymers as compared to using nonlabeled polymers. For the latter, closing mass balances are impaired as polymer-added carbon remaining in the soil cannot readily be delineated from carbon in soil organic matter.

Assessing the Contribution of Residual Polyester to the Nonmineralized Polyester-Added ¹³C

The amounts of residual PBAT and PBSeT, ¹³C_{polymer residual} (expressed as the percent of initially added polyester C extracted, converted to polyester-¹³C added to the soil at the onset of the incubation) that remained in the soils at the end of the incubations are plotted as vertical orange lines on the left side. This assessment could not be conducted for cellulose as it is not solvent-extractable. It is important to note here that incubation experiments were terminated prior to ¹³CO₂ formation plateauing, implying that PBAT and PBSeT biodegradation was still ongoing. After 425 days of incubation, the three different monomer-specific ¹³C-labeled PBA₅₀T₅₀ variants had similar ¹³C_{polymer residual} values of 67 (±1) %, 71 (±7) %, and 74 (±4) % for PBA₅₀(1-¹³C₁-T)₅₀, PB­(1,6-¹³C₂-A)₅₀T₅₀, and P­(¹³C₄-B)­A₅₀T₅₀, respectively (average ± range for duplicate samples per polyester variant). By comparison, ¹³C_{polymer residual} after 425 days was lower for the two ¹³C-labeled PBSe₅₀T₅₀ variants at 29 (±7) % and 30 (±1) % (n = 2) for P­(¹³C₄-B)­Se₅₀T₅₀ and PBSe₅₀(1-¹³C₁-T)₅₀, respectively, consistent with their higher extents of mineralization. Finally, ¹³C_{polymer residual} of the PBA_100‑X T _X series with varying T content was 3 (±1) %, 2 (±1) %, 10 (±1) %, and 91 (±5) %, (n = 3) for PBA₁₀₀T₀, PBA₈₀T₂₀, PBA₇₀T₃₀, and PBA₅₃T₄₇, respectively. We note that the few cases in which calculated ¹³C_{polymer residual} exceeded ¹³C_{nonmineralized} likely resulted from the 3 g of soil being extracted containing a nonrepresentative, slightly higher amount of polyester than expected for a representative subsample taken from the 100 g of bulk soil. This finding implies that downsizing soil aggregates by sieving the entire soil prior to subsampling 3 g for Soxhlet extraction did not completely remove the heterogeneity in polyester distribution in the soil. This explanation is favored over ¹³C_{nonmineralized} values being inaccurate, given that ¹³C_{nonmineralized} is indirectly validated by closed mass balances when combined with ¹³C_mineralized. Despite potential small uncertainties caused by extracting soil aliquots, ¹³C_{polymer residual} for all polyester variants decreased with decreasing ¹³C_{nonmineralized}, as expected (see Section S8).

Assessment of Chemical Structure-Soil Biodegradability Relationships of Biodegradable Polyesters

The use of ¹³C-labeled polyester variants differing in their chemical composition and the combined determination of ¹³C_mineralized, ¹³C_{nonmineralized}, and ¹³C_{polymer residual} allows for a detailed analysis of the polyester chemical structure-biodegradability dependencies. The observed differences between variants must result from the polyester chemical structure, as the same soil and incubation conditions were used in all experiments.

Variations in the Aliphatic Diacid

Mineralization extents after 425 days were much higher for PBSe₅₀T₅₀ (i.e., ¹³C_{nonmineralized} from 52 to 59%) than for PBA₅₀T₅₀ (from 17 to 26%), implying that variations in the aliphatic diacid in aliphatic-aromatic copolyesters strongly affect polyester biodegradation. By subtracting the amounts of extracted residual polyesters, ¹³C_{polymer residual}, from the amounts of total polyester-added ¹³C left nonmineralized in the soils, ¹³C_{nonmineralized}, we could estimate the amounts of polyester-¹³C that were incorporated into microbial biomass, ¹³C_biomass (note, however, that this analysis calls for a careful interpretation, particularly for polyesters with low mineralization extents, due to several confounding factors addressed in Section S9). For PBSe₅₀T₅₀, we estimated ¹³C_biomass to be 19 (±6) % across all incubations (n = 6) (Figure a), revealing that considering only mineralization extents, despite them being high, would result in substantially underestimating total biodegradation extents of this polyester. While biomass incorporation also seems to have occurred for the PBA₅₀T₅₀ and PBA₅₃T₄₇ variants, its extent was much smaller, reflecting the overall lower mineralization extents (Figure a) (i.e., the PBA₅₀T₅₀ and PBA₅₃T₄₇ variants exhibited low carbon incorporation into biomass of ¹³C_biomass 3 (±9) % (n = 12)). The high uncertainty in ¹³C_biomass results from ¹³C_{polymer residual} in some bottles exceeding ¹³C_{nonmineralized} (see above discussion), resulting in tentatively negative biomass incorporation (see Figure a). The low incorporation of ¹³C into biomass for the one PB­(1,6-¹³C₂-A)₅₀T₅₀ incubation that exhibited a high mineralization extent (see sample marked with an asterisk (*) in Figure a) likely reflects extensive decarboxylation of the ¹³C in the carboxylate carbons of A into ¹³CO₂, while the four nonlabeled inner carbons of A of this variant were relatively more extensively incorporated into biomass. This pattern of carbon position-specific preferential mineralization or biomass incorporation has been previously demonstrated and discussed in detail for soil incubations of position-specific ¹³C-labeled succinate (S; i.e., 1,4-¹³C₂-S and 2,3-¹³C₂-S) and the corresponding position-specific ¹³C-labeled variants of the S-containing polyester polybutylene succinate (PBS). The monomer-specific labeling of PBA₅₀T₅₀ and PBSe₅₀T₅₀ further allowed demonstrating that carbon from all labeled monomeric units (i.e., A, B, and T) biodegraded to similar extentsthereby ruling out that any of these building blocks underwent delayed biodegradation. This finding is consistent with previous work demonstrating rapid biodegradation of ¹³C-labeled B, A, and T when added as monomers to soil, suggesting that low-molecular-weight monomeric and oligomeric polymer breakdown products do not accumulate in soils. We note that ¹³C-labeled Se was not commercially available at the time of synthesis. Overall, faster and more extensive biodegradation of PBSe₅₀T₅₀ than PBA₅₀T₅₀ in the tested soil is consistent with previously reported differences in PBAT and PBSeT degradation in soil and compost assessed indirectly by gravimetric weight loss measurements. ,,

4.

Dependence of polyester biodegradation on chemical structure. (a) Microbial utilization of polyester carbon in soil incubations, shown as the amounts of polyester-¹³C incorporated into microbial biomass (¹³C_biomass) vs the extents of polyester-¹³C mineralized (¹³C_mineralized) at the end of incubations (¹³C_mineralized values are replotted from Figure ). Small, faint symbols represent values for individual incubations; large squares and error bars represent averages and standard deviations, respectively, for the indicated polyester variants (note that the inverted triangle marked with an asterisk (*) represents one replicate of the PB­(1,6-¹³C₂-A)₅₀T₅₀ variant that mineralized to an exceptionally high extent; therefore, its value was excluded from the average). The dashed black line corresponds to states of complete polyester biodegradation (i.e., ¹³C_mineralized + ¹³C_biomass = 100%). Light gray lines represent different theoretical values of the carbon use efficiencies (CUE, i.e., ¹³C_biomass/(¹³C_biomass+¹³C_mineralized); value indicated by the respective label in the plot). (b) Terephthalate (T) contents in residual polyesters extracted from soils at the end of incubations (symbols, “residual”; data points correspond to three replicate incubations) relative to those of the starting polyesters (light horizontal lines, “initial”), as determined using proton nuclear magnetic resonance spectroscopy (¹H NMR; see Materials and Methods for details on measurements and calculations).

While the cause for enhanced mineralization in one of the PB­(1,6-¹³C₂-A)₅₀T₅₀ and P­(¹³C₄-B)­Se₅₀T₅₀ replicate incubations remains unidentified, two explanations seem plausible. First, these two incubation bottles may have contained a microbial strain that is particularly competent in biodegrading PBAT and PBSeT and began to proliferate and degrade these polyesters after approximately 80 days of incubation. Second, a random adaptation of one (or more) microbial strain in these two bottles may have enhanced their potential to biodegrade the two polyesters. These adaptations may have involved the upregulation of the production and exudation of highly active extracellular esterases, as enzymatic ester bond hydrolysis is generally thought to control the overall biodegradation rates of PBAT and PBSeT in soil. Irrespective of which explanation holds true, the finding of sudden enhanced biodegradation highlights that soil microbial communities can become more adept, through either adaptation or selection of specialist microbial strains, and that repeated inputs of biodegradable polyesters to soils could enhance biodegradation rates.

Variations in the T Content of PBAT

As compared to the PBA₅₀T₅₀ and PBA₅₃T₄₇ variants, the PBA₇₀T₃₀, PBA₈₀T₂₀, and PBA₁₀₀T₀ variants not only underwent much more extensive mineralization but also showed substantial incorporation of polyester carbon into microbial biomass (i.e., ¹³C_biomass= 17 (±1) % (n = 9); Figure a). More extensive biomass incorporation is consistent with PBSe₅₀T₅₀ biodegradation, as this polyester also underwent fast and extensive mineralization in soils. For such polyesters, incubation experiments determining only ¹³C_mineralized (x-axis in Figure a) underestimate true biodegradation by the extent of ¹³C_biomass (y-axis in Figure a; the corresponding carbon use efficiencies (CUEs) are shown as gray lines). In this figure, the “true” extent of biodegradation corresponds to the vertical offset between the data points and the dashed black line representing “complete biodegradation” (i.e., ¹³C_mineralized + ¹³C_biomass = 100%) (and not the vertical offset between the dashed line and the x-axis). The finding of low ¹³C_biomass for polyesters which underwent slower mineralization may not necessarily imply that there was no incorporation but that cycling of polyester-derived ¹³C through the microbial biomass pool into ¹³CO₂ occurred at rates comparable to (or faster than) the rate by which ¹³C was incorporated into microbial biomass. In such cases, the steady-state ¹³C_biomass would be small (as shown here for PBAT with T contents ≥ 47%), which aligns with modeled biodegradation dynamics of PBS in soil.

We ascribe the pronounced decrease in soil biodegradability from the PBA₁₀₀T₀, PBA₈₀T₂₀, and PBA₇₀T₃₀ variants to the PBA₅₃T₄₇ (and PBA₅₀T₅₀) variant to an underlying decrease in the enzymatic hydrolyzability of these polyesters. , Higher T contents lead to increased contents of T-enriched microcrystalline domains, which undergo slower enzymatic hydrolysis. This explanation is directly supported by observed changes in the chemical composition of the extracted residual PBAT and PBSeT at the end of the incubations based on their ¹H NMR spectra. Both the extracted residual P­(¹³C₄-B)­A₇₀T₃₀ and P­(¹³C₄-B)­A₈₀T₂₀ variants showed increased T contents of 35 (±1) % and 29 (±2) % (n = 3 per variant), respectively, as compared to the T contents of the initially added polyesters of 30 and 20%, respectively. Similarly, the residual PBSeT extracted from the soils also had elevated T contents of 59 (±2) % (n = 6) as compared to 50% in the initially added PBSe₅₀T₅₀. An increased T content of 56% (from initially 50%) was also found for the residual PB­(1,6-¹³C₂-A)₅₀T₅₀ in the one incubation bottle that showed enhanced biodegradation (inverted triangle marked with an asterisk (*) in Figure b), demonstrating that the T-enrichment also affected higher T content PBAT variants when undergoing extensive biodegradation. Our finding that extensive biodegradation results in increased T contents in the residual polyesters is in agreement with previous reports of relative increases in T contents in aromatic-aliphatic polyesters upon both abiotic hydrolysis , as well as incubations with isolated soil microorganisms or in bulk soils. ,, It is important to emphasize that while there is enrichment in T during the biodegradation, the terephthalate units are also subject to mineralization. This is seen in the two variants that were ¹³C-labeled in T (i.e., PBA₅₀(1-¹³C₁-T)₅₀ and PBSe₅₀(1-¹³C₁-T)₅₀), both of which showed production of ¹³CO₂, unequivocally demonstrating that biodegradation of PBA_100‑X T _X and PBSe_100‑X T _X also involves their terephthalate components.

Implications

This work established that changes in the monomer composition of aliphatic-aromatic copolyesters can strongly affect their biodegradation rates and extents in soils. Compared to polybutylene adipate-co-terephthalate with (close to) equimolar contents of T and A, mineralization rates and biodegradation extents in soil increased largely when replacing the diacid A by Se (i.e., increasing the length of the aliphatic diacid from six to ten carbon atoms) and when decreasing the T content while increasing the A content. These results highlight a unique potential of the modular composition of aliphatic-aromatic polyesters: the possibility of tuning their biodegradation rates through alteration of their chemical composition. Polyester chemical composition may thus present a means to establish a desired polyester biodegradability for not only soils but also other natural (e.g., freshwater and marine) or engineered (e.g., compost) receiving environments. For instance, it is conceivable that a set of commercially certified soil-biodegradable products composed of aliphatic-aromatic polyesters, such as thin mulch films, could be marketed that vary in their monomeric polyester compositions to match the biodegradation potential of specific receiving soils. Clearly, such an approach requires also a more detailed understanding of the factors driving variations in the biodegradation potential of soils , (or other receiving systems). ,, It is noteworthy, however, that the rate of biodegradation can also be adjusted by other approaches, such as compounding with other polymers or modifying the crystallinity by altering the production process. Furthermore, we note that the average molecular weights of the PBAT variants synthesized in the reported small-scale laboratory reactors were lower than those in PBAT (and other polyesters) commonly used in commercial products (e.g., PBAT used in certified soil-biodegradable mulch films). Considering that polyester biodegradation rates may decrease with increasing molecular weight, biodegradation rates determined herein may be higher than those of polyesters with the same or comparable chemical composition in commercial products.

Solvent extraction of residual polyesters from soils followed by ¹H NMR analysis is shown to be useful not only for accurately quantifying the true extent of polyester biodegradation (which may be underestimated by mineralization measurements alone) but also for determining changes in the monomeric compositions of a polyester during its biodegradation. The observed relative increase in T contents in the residual polyesters can be rationalized based on the established faster biodegradation of A- and Se-enriched amorphous domains and the slower biodegradation of T-enriched crystalline domains. This finding thus highlights how chemical structure-soil biodegradability assessments can be linked to a molecular-level enzymatic hydrolysis reaction. This finding also supports that the assessment of polyester hydrolyzability by natural extracellular esterases has predictive power for their environmental biodegradability. Our work further supports the use of solvent extraction of residual polyesters coupled to their quantification and chemical characterization in future biodegradation studies. This is especially interesting for studies following polyester (or products composed thereof) biodegradation in the open environment (e.g., field soils) in which mineralization measurements are not feasible (or even impossible). In such studies, changes in the monomer composition of extracted, residual polyester can be used as an independent indicator (besides decreasing polyester concentrations) that biodegradation has occurred. It remains to be established how such transient changes in the chemical composition of residual polyesters during biodegradation affect their subsequent biodegradation rates. It may be possible to counteract potential slowdown in subsequent biodegradation by other chemical modifications of the polyester structure or by using polymer blends that lead to more rapid or synergistic biodegradation.

Quantification of the residual polyesters, combined with mineralization extents, revealed a more extensive incorporation of polyester carbon into microbial biomass for polyesters undergoing fast biodegradation. Polyesters undergoing fast and extensive mineralization may have substantial polyester carbon assimilated into the microbial biomass. By comparison, for slowly mineralizing polyesters, little of the nonmineralized polyester carbon may be incorporated in microbial biomass. Quantification of C_{polymer residual} does not require ¹³C-labeling and thus is a valuable end point measurement also for standard incubations following polyester mineralization: in combination, quantification of C_mineralized and C_{polymer residual} can disclose the true extent of polyester biodegradation and of polyester-C incorporation into microbial biomass.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.est.5c03099.

• Physicochemical properties of polyesters (Section S1); calculations of polymer mineralization rates and extents (Section S2); details on quantifying total nonmineralized polyester- and cellulose-added ¹³C remaining in soils, ¹³C_{nonmineralized} (Section S3); details on quantifying residual polyester in soils, C_{polymer residual} (Section S4); mineralization dynamics of P­(¹³C₄-B)­A_100‑X T _X in soil (Section S5); mineralization rates of cellulose-¹³C vs total cellulose-¹³C added to soil (Section S6); compilation of ¹³C_mineralized, ¹³C_{nonmineralized}, and ¹³C_{polymer residual} data for all incubations (Section S7); correlation between ¹³C_{polymer residual} and ¹³C_{nonmineralized} for polyesters at the end of soil incubations (Section S8); interpretation of ¹³C_biomass (Section S9) (PDF)

The authors declare no competing financial interest.