# Enhancing the biodegradation of PBS, PBAT, and their biocomposites under decentralized mesophilic composting conditions through bioaugmentation

Jules Bellon, Feriel Bacoup, Richard Gattin

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Jules Bellon, Feriel Bacoup, Richard Gattin. Enhancing the biodegradation of PBS, PBAT, and their biocomposites under decentralized mesophilic composting conditions through bioaugmentation. Journal of Environmental Chemical Engineering, 2026, 121600, 14 (2), pp.121600. 10.1016/j.jece.2026.121600. hal-05511083

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# Enhancing the biodegradation of PBS, PBAT, and their biocomposites under decentralized mesophilic composting conditions through bioaugmentation

Jules Bellon* ID, Feriel Bacoup ID, Richard Gattin

UniLaSalle, Transformation and Agro-Resources Research Unit (ULR 7519 UniLaSalle–Université d'Artois), Mont-Saint-Aignan 76130, France

## ARTICLE INFO

### Keywords:

- Biodegradable plastics

- Mesophilic composting

- Bioaugmentation

- PBS

- PBAT

- Biocomposites

## ABSTRACT

The limited biodegradability of polyester-based bioplastics under mesophilic conditions remains a major constraint for decentralized end-of-life pathways, including home and community-scale composting. This study assesses whether bioaugmentation can improve the mesophilic compostability of two biodegradable polyesters, poly (butylene succinate) (PBS) and poly (butylene adipate-co-terephthalate) (PBAT), as well as their bio-composites containing 10 % agro-industrial fillers (orange peel and brewer's spent grain). Two bacterial consortia were isolated through selective enrichment in minimal medium using compost extracts incubated with the target materials for 200 days at 28 °C. Biodegradation assays were conducted in closed respirometric chambers following an adapted NF T51-800 / ISO 14855-1 protocol. Vermiculite activated with compost extract served as the substrate, with or without the enriched consortia, and the tested materials were the only carbon sources. Mineralization kinetics were monitored for 133 days at 28 °C. Structural changes were evaluated using DSC, FTIR, and macroscopic and SEM imaging, while microbial dynamics were characterized by CFU enumeration and Community-Level Physiological Profiling (Biolog EcoPlates™). Bioaugmentation substantially enhanced PBS biodegradation, increasing final mineralization by + 68.3 % relative to the control (43.6 %) and reaching 72.4 %, a value approaching the 90 % threshold defined for home-compostability. No improvement was observed for PBS/OP. PBAT exhibited very limited biodegradation under mesophilic conditions, with no detectable effect of bioaugmentation. Hydrolysis was confirmed as the dominant degradation mechanism. Overall, bioaugmentation appears to be a promising strategy to accelerate the mesophilic biodegradation of PBS, although its effectiveness depends strongly on polymer-filler interactions and material formulation.

## 1. Introduction

The global escalation of plastic pollution has emerged as one of the most critical environmental issues of the twenty-first century [1]. Plastics accumulate today in all major environmental compartments, including terrestrial, aquatic, and even atmospheric systems. In these environments, they exert multiple harmful effects on ecosystem functioning and human health [2].

Manufactured thermoplastic products are composed almost entirely (~99 %) of high-molecular-weight conventional polymers such as PE, PET, PP, PS, and PVC. These materials exhibit exceptional thermodynamic and kinetic stability because their macromolecular structures consist exclusively of carbon-carbon backbone linkages (e.g. PE, PS, PP). They may also contain inherently fewer stable bonds, such as the ester groups in PET. However, these bonds remain largely inaccessible and only weakly reactive due to the stabilizing influence of adjacent

aromatic benzene rings [3].

In contrast, several high-molecular-weight polymers are now classified as biodegradable (e.g. PLA, PBS, PBAT, PHAs, PCL), offering a promising avenue to mitigate the long-term accumulation of plastic waste in the environment [4]. In particular, poly (butylene succinate) (PBS) and poly (butylene adipate-co-terephthalate) (PBAT) are biodegradable polyesters that can be fully bio-based and are considered strong candidates to replace certain polyolefins. PBS provides an attractive balance of mechanical strength, ductility, flexibility, impact resistance, and thermal stability [5]. PBAT, by contrast, is highly flexible and exhibits mechanical behaviour like low-density polyethylene (LDPE), with a reported elongation at break of approximately 670 % and a tensile strength of around 21 MPa [6]. Both polymers can be used individually or in blends, together with appropriate additives, to produce flexible or semi-rigid items such as plastic bags or agricultural mulch films [5].

However, the biodegradability of biodegradable polymers strongly

<footnote>* Corresponding author.
E-mail address: jules.bellon@unilasalle.fr (J. Bellon).</footnote>
depends on the environmental compartment in which degradation occurs (e.g. compost, soil, or aquatic systems) [7]. Biodegradability is therefore not an intrinsic material property, but an extrinsic one. It results from the interaction between polymer characteristics such as chemical structure, molar mass, crystallinity, glass transition temperature ($T_g$), and hydrophilicity, and the surrounding environmental conditions. These conditions include both biotic factors (e.g. microbial abundance, community composition, and enzymatic activity) and abiotic parameters (e.g. temperature and moisture) [8].

In this respect, compost represents an environment with a strong biodegradative potential toward biodegradable polyesters, primarily due to its high microbial abundance and taxonomic diversity [9]. Two main composting systems can be distinguished: (i) centralized (i.e. industrial) composting, and (ii) decentralized composting, which encompasses community-scale and home-composting practices [10].

Decentralized composting offers several advantages over centralized systems. These include simplified logistics of waste handling by reducing the need for extensive sorting and long-distance transport. Additional benefits include lower collection and transportation costs, as well as greater citizen engagement in the local management of biodegradable waste [11,12]. Beyond operational benefits, decentralized composting has also been shown to generate positive social externalities. For example, in the municipality of Surabaya, Indonesia, Maeda [13] demonstrated that households adopting decentralized composting exhibited enhanced pro-environmental behaviours, including a 23 % reduction in annual household waste generation after three years of practice.

Decentralized composting systems predominantly operate under mesophilic conditions (20–45 °C), whereas the thermophilic phase (> 45 °C) is generally absent or, when present, only transient [11]. As a result, temperatures remain substantially lower than those reached in industrial composting which typically leads to slower biodegradation kinetics. Consequently, for several widely used polyesters, biodegradation under mesophilic conditions is often partial, markedly slower, or even negligible [14]. In particular, PBS and PBAT are not considered decentralized-compostable materials, in contrast to polymers such as poly (hydroxybutyrate-co-valerate) (PHBV), poly (ε-caprolactone) (PCL), and thermoplastic starch (TPS), which are recognised as decentralized-compostable [15].

Thus, enhancing the biodegradability of these two major biodegradable polymers under mesophilic composting conditions remains a critical research challenge [16]. Addressing this limitation is essential for establishing an effective end-of-life management pathway specifically tailored to these materials.

Several strategies have been proposed to enhance the biodegradation of biodegradable polymers under composting conditions. One strategy consists in formulating biocomposites by incorporating organic fillers into polymer matrices, which can enhance material disintegration and promote microbial colonization [17,18]. Another strategy involves the addition of biostimulants to the degradation medium, such as readily available carbon sources or targeted nutrients. This approach has been shown to enhance PLA biodegradation under mesophilic conditions [19].

Vermicomposting has also been explored, relying on the introduction of earthworm species such as *Eisenia fetida* and *Eisenia andrei*, which can increase substrate aeration and stimulate microbial activity [20,21]. In parallel, bioaugmentation strategies based on the enrichment of the degradation medium with selected microbial strains have been investigated, mainly under thermophilic composting conditions for PLA at 58 °C [22,23]. Finally, the incorporation of specific enzymes into nano-structured materials has been proposed to facilitate early hydrolysis of polymer chains [24].

However, few of these studies have focused on PBS and PBAT, and the biodegradability of these two polyesters under such conditions remains a significant challenge.

Moreover, the studies conducted to date are generally not performed

within the framework of an appropriate normative specification for mesophilic composting, such as the French standard for home-compostable plastics (NF T51-800) [25] or an equivalent (i.e. AS 5810, TÜV Austria - OK compost HOME) [26,27]. These standards provide a clear experimental framework, primarily based on the ISO 14855-1 procedure [28], and define specific requirements to be met. For instance, the mineralization extent of the tested plastic material must exceed 90 % after $t \le 365$ days of incubation at $T \le 30$ °C. It is therefore essential to operate within such a normative framework to ensure that tests are conducted under standardized and mutually comparable conditions.

In a previous study [29], we developed thermoplastic biocomposites based on the two polyester matrices poly (butylene succinate) (PBS) and poly (butylene adipate-co-terephthalate) (PBAT), with a view to their use in packaging applications. These matrices were combined with organic fillers derived from agro-industrial residues, specifically orange peel (OP) and brewer's spent grain (BSG). In doing so, the resulting biocomposites incorporated 10 wt% of these under-utilised by-products, providing an economically and environmentally sustainable filler source.

In the present work, we investigate a complementary strategy to enhance the biodegradability of these thermoplastic materials under mesophilic composting conditions, namely bioaugmentation. This approach, together with biostimulation, falls within the broader concept of bioremediation, originally developed for the treatment of contaminated soils and polluted environments [30]. In this context, the substrate is treated as a pollutant [31]. Beyond its environmental remediation roots, bioremediation has progressively inspired applied microbial technologies aimed at steering biological activity in managed degradation systems.

In recent years, this conceptual framework has translated into the emergence of commercial “compost activators”, consisting of microbial blends sometimes supplemented with enzymes or nutrient additives and intended to accelerate the early stages of organic-matter degradation. These products mainly act by promoting enzymatic hydrolysis of lignocellulosic substrates during the mesophilic phase [32]. From a technological standpoint, microbial inoculants used in such formulations can be stabilized through established routes, such as freeze-drying, ensuring adequate shelf life and transportability [33].

However, current compost activators are not designed to enhance the biodegradative capacity of composting environments toward specific recalcitrant substrates such as biodegradable polyesters. This gap suggests that, in the future, targeted biotechnological solutions could be developed. These approaches would be conceptually analogous to existing compost activators, while relying on specialized microbial consortia. Their objective would be to selectively stimulate microbial communities capable of degrading biodegradable polyesters.

The feasibility of deploying such dedicated activators is, however, strongly dependent on the operational context of composting systems. Community composting systems are increasingly documented as effective decentralized waste-management solutions and, in contrast to household composting, benefit from structured operation based on standardized procedures and dedicated management, including clear financial responsibility for system operation and maintenance. These characteristics make the integration of specialized compost activators incorporating defined microbial formulations considerably more realistic in such settings than in home-composting systems [34–36].

The implementation of such activators within community composting systems could be facilitated by public policy instruments, including incentive-based measures or performance-based requirements applied to operators managing decentralized composting infrastructures. These instruments would be consistent with extended producer responsibility frameworks and with the progressive implementation of separate bio-waste collection across Europe [37,38]. However, beyond regulatory support and operational feasibility, the effectiveness of such approaches ultimately depends on biological constraints governing microbial
establishment and function.

The success of a bioaugmentation strategy lies within the domain of microbial ecology. It critically depends on the ability of the inoculated strains to establish themselves in the new environment, persist over time, and efficiently degrade and metabolize the target carbon substrate. Such success requires the functional integration of the introduced strains within the resident microbial community, which may occur through the establishment of symbiotic or mutualistic interactions, or alternatively, through sufficient competitive ability to dominate specific ecological niches [39].

Previous investigations have consistently reported that bioaugmentation approaches enhance the mineralization performance of PLA under thermophilic composting conditions, thereby accelerating its biodegradation relative to non-augmented systems [22,40]. However, the performance of bioaugmentation under mesophilic composting conditions remains insufficiently documented. To date, no study has investigated this approach for PBS or PBAT, whether as pristine polymers or incorporated into biocomposites. This remains the case under conditions compatible with decentralized biodegradability standards (e. g. NF T51–800; < 30 °C).

Furthermore, decentralized composting tests are typically conducted using mature compost as the degradation medium [41]. This matrix plays a central role in the evaluation procedure. It simultaneously provides a physical support for the biodegradation process and acts as a source of microbial inoculum. In addition, it serves as a nutrient reservoir beyond the target substrate, as it contains a wide range of readily metabolizable compounds, including organic acids, phenolics, and polysaccharides [28,42].

However, carbon substrates present within the organic matter of mature compost can modulate microbial activity via positive or negative priming effects [43,44]. These dynamics may shift microbial metabolic allocation away from the target polymer, thereby causing its biodegradability to be over- or underestimated throughout the degradation process [45].

To address this limitation, inert degradation media such as vermiculite (a phyllosilicate of the mica group), which contains no carbon source (i.e. (Mg, Fe²⁺, Fe³⁺, Al)₃(Si, Al)₄O₁₀(OH)₂·4H₂O), can be used. It offers a compromise between the ecological representativeness of composting processes and the experimental control required, while remaining fully compliant with the NF T51–800 standard. This medium is less complex than mature compost. This reduces interference from background organic matter. At the same time, it remains more environmentally realistic than liquid cultures or agar-based systems.

In this context, we conducted a mesophilic compostability test using vermiculite activated with a low-carbon mature compost extract, following the methodological framework initially established by Bellia et al. [46] and later refined and applied by Gattin et al. [47] and Copinet et al. [48]. This approach preserves the functional diversity of compost-associated microbial communities while reducing confounding nutrient effects, thereby providing a reliable and controlled framework for assessing bioaugmentation strategies under mesophilic conditions.

Building on this controlled experimental framework, the careful selection of tested materials is essential to ensure a meaningful interpretation of biodegradation outcomes. Commercial biodegradable plastics are complex and proprietary systems, whose exact formulation is generally not disclosed. This lack of transparency concerns both polymer composition and the nature and content of additives. It prevents unambiguous attribution of biodegradation behaviour and limits robust interpretation and extrapolation of observed trends. To avoid formulation-dependent bias, well-defined, non-commercial reference polymers were intentionally used in this study, allowing the effects of bioaugmentation and agro-industrial fillers to be assessed under controlled conditions. This reduced level of system complexity represents a necessary upstream step for reliable interpretation, prior to the evaluation of these strategies on commercial formulations based on PBS and PBAT.

In this context, the bioaugmentation strategy investigated in the present study should be considered a first proof-of-concept toward the development of compost activators targeting PBS- and PBAT-based biodegradable plastics under decentralized mesophilic composting conditions. It was not intended as a directly deployable implementation. Instead, it evaluates whether targeted microbial enrichment can enhance the biodegradation potential of these materials under controlled mesophilic composting conditions. This approach provides a scientific basis for the rational design of dedicated compost activators.

## 2. Materials and methods

### 2.1. Fabrication and initial characterization of plastic materials

The fabrication process and initial characterization of the thermo-plastic materials were previously described in detail by Bellon et al. [29]. Briefly, square specimens (2 × 2 cm) were prepared from poly (butylene succinate) (PBS), poly (butylene adipate-co-terephthalate) (PBAT), and their corresponding biocomposites containing 10 wt% orange peel (PBS/OP) or brewer's spent grain (PBAT/BSG) (Fig. 1).

The two organic fillers are agri-food by-products incorporated to promote biodegradation under mesophilic composting conditions and to reduce the overall material cost. Biocomposite sheets were produced via melt extrusion followed by thermo-compression, whereas pristine polymers were processed solely by thermo-compression.

Key parameters known to influence biodegradability, including sample thickness, molecular weight, and crystallinity, are summarized in Table 1. The molecular weight values fall within the ranges commonly reported for PBS and PBAT materials produced by extrusion or thermo-compression (≥ 50.000 g·mol⁻¹). This confirms that the starting materials were representative and did not exhibit unusually low molecular weights.

The main mechanical properties obtained from standardized tensile tests (ISO 527–3) on pristine polymers and biocomposites are

<figure><img src="image_3.png" /><figcaption>Fig. 1. Photograph of materials fabricated by thermopressing. From left to right: pure polymer, biocomposite with orange peel (OP), biocomposite with brewer's spent grain. From top to bottom: PLA and PLA-based composites; PBS and PBS-based biocomposites; PBAT and PBAT-based biocomposites.</figcaption></figure>
<caption>**Table 1**</caption>

Thickness, molecular weight and crystallinity of the pristine polymers and biocomposites.

<table><thead><tr><th>Materials</th><th>Thickness (µm)</th><th>M<sub>w</sub> (g·mol<sup>-1</sup>)</th><th>M<sub>n</sub> (g·mol<sup>-1</sup>)</th><th>D</th><th>χ (%)</th></tr></thead><tbody><tr><td>PBS</td><td>219 ± 53</td><td>146 700</td><td>54 000</td><td>2.72</td><td>54.2<br/>± 5.0</td></tr><tr><td>PBS/OP</td><td>219 ± 49</td><td>143 000</td><td>53 300</td><td>2.68</td><td>53.1<br/>± 5.0</td></tr><tr><td>PBAT</td><td>385 ± 77</td><td>151 000</td><td>59 700</td><td>2.53</td><td>6.0 ± 5.0</td></tr><tr><td>PBAT/BSG</td><td>316 ± 44</td><td>120 100</td><td>50 500</td><td>2.38</td><td>9.4 ± 5.0</td></tr></tbody></table>

**Notes.** Weight-average molecular weight (M~w~); number-average molecular weight (M~n~); and polydispersity index (D = M~w~/M~n~) were determined by SEC–MALLS. Crystallinity (χ) was calculated from the first DSC heating scan.

summarized in Table 2. A comprehensive analysis of how filler incorporation influences mechanical properties, in relation to filler characteristics, can be found in our previous study [29].

## 2.2. Isolation and identification of material-degrading bacteria

A continuous selective enrichment procedure was applied to isolate bacteria capable of degrading the target materials under mesophilic conditions in liquid media. This approach was adapted from the procedure previously reported by Castro-Aguirre et al. [22] for PLA in thermophilic composting conditions.

Mature compost samples (2 g) were collected at the end of a 200-day biodegradation experiment performed at 28 °C in laboratory-scale bio-reactors containing PBS- or PBAT/BSG-based materials. Samples were transferred into Erlenmeyer flasks containing 25 mL of sterile M9 minimal medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with five disinfected plastic fragments (approximately 2 × 2 cm) as the sole carbon source. Prior to inoculation, plastic surfaces were disinfected with 70 % ethanol and air-dried under sterile conditions following Castro-Aguirre et al. [49].

The cultures were incubated under aerobic, agitated mesophilic conditions (28 °C, 80 rpm) in the dark. Each week, under sterile conditions, the cultures were transferred into fresh M9 medium containing new sterile fragments of the same polymeric material. A 150 µL aliquot from the previous culture was used as the inoculum, together with one fragment recovered from the preceding enrichment step. This procedure was repeated for four consecutive subcultures.

This serial transfer protocol imposed a strong selective pressure, enriching for microorganisms capable of utilizing material-derived carbon under mesophilic conditions. The final enrichment cultures were serially diluted and streaked onto R2A agar plates, which were then incubated at 28 °C. Distinct colonies were purified through successive re-streaking and stored at –80 °C in cryobeads (DeltaLab, Barcelona, Spain), where they were preserved for subsequent use as bioaugmentation inocula in the vermiculite-based assays conducted in this study. Strain identification was conducted by an accredited external

<caption>**Table 2**</caption>

Mechanical properties of pristine polymers and biocomposites determined by tensile testing.

<table><thead><tr><th>Materials</th><th>Young's modulus (MPa)</th><th>Tensile Strength (MPa)</th><th>Elongation at Break (%)</th></tr></thead><tbody><tr><td>PBS</td><td>615 ± 16<sup>b</sup></td><td>30.4 ± 1.5<sup>a</sup></td><td>8.7 ± 1.5<sup>a</sup></td></tr><tr><td>PBS/OP</td><td>683 ± 30<sup>a</sup></td><td>22.8 ± 1.5<sup>b</sup></td><td>4.9 ± 1.5<sup>b</sup></td></tr><tr><td>PBAT</td><td>80.5 ± 4.0<sup>b</sup></td><td>21.3 ± 3.2<sup>a</sup></td><td>601.3 ± 3.2<sup>a</sup></td></tr><tr><td>PBAT/BSG</td><td>112.0 ± 1.4<sup>a</sup></td><td>12.3 ± 1.1<sup>b</sup></td><td>408.8 ± 1.1<sup>b</sup></td></tr></tbody></table>

**Notes.** Values are reported as mean ± standard deviation. Different superscript letters (a, b) indicate statistically significant differences between materials within a polymer type (p < 0.05).

laboratory (Eurofins BPT Biologics, France) using MALDI–TOF MS and, when necessary, complemented by long-read 16S rDNA sequencing.

## 2.3. Ultimate aerobic biodegradation test on an activated inert substrate

The experimental setup described below was designed to simulate decentralized composting systems operating under mesophilic conditions, representative of both home and community-scale composting practices.

In this study, biodegradation is operationally assessed through CO₂-based mineralization measurements, in accordance with ISO 14855–1 and related compostability standards. Under aerobic conditions, polymer carbon is first assimilated into microbial biomass and subsequently converted into CO₂ through metabolic turnover. Although biodegradation strictly encompasses both assimilation and mineralization steps, CO₂ evolution represents the dominant and normatively defined endpoint and is therefore commonly used as a quantitative proxy for biodegradation in standardized composting tests.

Ultimate aerobic biodegradation was monitored for 133 days using closed respirometric systems (ECHO Instruments, Slovenia) operated under mesophilic conditions (28 ± 1 °C) in the dark. The experimental setup followed ISO 14855–1 and was adapted from the NF T51–800 specification.

Plastic specimens (17 g) were mixed with 170 g of vermiculite (2–5 mm; inert and carbon-free) (Plantit, HydroGarden Ltd., Coventry, UK), moistened to approximately 70 % with 410 mL of a pH 7.2 buffered mineral solution. The composition of this solution (per 5 L) was: 0.650 g CaCl₂·2H₂O, 34.850 g Na₂HPO₄·2H₂O, 18.750 g KH₂PO₄, 20 g (NH₄)₂SO₄, 1 g MgSO₄·7H₂O, 0.0135 g FeSO₄·7H₂O, 0.005 g MnSO₄·7H₂O, 0.005 g ZnSO₄·7H₂O, 0.005 g H₃BO₃, 0.005 g KI, and 0.005 g (NH₄)₆Mo₇O₂·4H₂O [47,48].

Substrate moisture was maintained by weekly addition of ultrapure water to the lower compartment of the bioreactors. In addition, 15 mL of mineral solution was supplied on days 30, 60, 90, and 120 after the start of the experiment. The contents of the reactors were manually mixed once per week to improve aeration and homogenization. Mineralization extents of samples were calculated using the following equation:

$$ \text{Mineralization}_{(i,t)} (\%) = \frac{\text{Sample}_{(i,t)} - \text{Blank}_{(i,t)}}{\text{ThCO}_2 (i)} \times 100 \quad (1) $$

where Sample(i,t) denotes the cumulative CO₂ produced at time t with material i; Blank(i,t) refers to the CO₂ released by the activated medium without plastic fragments; and ThCO₂(i) is the theoretical CO₂ yield from complete aerobic mineralization of material i, calculated from its organic carbon content determined by CHN analysis:

$$ \text{ThCO}_2 (i) (g) = C_{(i)} (g) \cdot \frac{44}{12} \quad (2) $$

This theoretical CO₂ yield corresponds to the amount of CO₂ that would be released under complete aerobic oxidation, according to the following stoichiometric equation:

$$ C_x H_y O_z + \left(x + \frac{y}{4} - \frac{z}{2}\right) O_2 \rightarrow xCO_2 + \frac{y}{2} H_2O \quad (3) $$

The moisture content of the samples was estimated using the weight loss method at 105 °C in a drying oven, until stabilization of their weight. These values were incorporated into the ThCO₂(i) terms to refine the accuracy of mineralization extent estimates over time.

### 2.3.1. Microbial inoculation

#### 2.3.1.1. *Baseline inoculum derived from mature compost extract.* A biotic compost extract used as the baseline inoculum was obtained by suspending 15 g of mature compost in 150 mL of Ringer solution (Ringer tablets, Merck) (Fig. 2). The resulting mixture was homogenized for 1 h,
Preparation of mature compost extract (with low carbon content)

<figure><img src="image_4.png" /><figcaption>Fig. 2. Schematic illustration of the preparation of the mature compost extract.</figcaption></figure>

filtered through a 1 mm sieve, and centrifuged at 10,000 rpm for 20 min using an Eppendorf Centrifuge 5810 R. The resulting pellet was resuspended in 150 mL of fresh Ringer solution. The mature compost originated from an industrial composting facility (SMEDAR, Saint-Jean-du-Cardonnay, 76594, France) processing lignocellulosic green waste and was compliant with the French quality standard for composites (NF U44-051). This low-carbon inoculum was added at the beginning of the experiment to all bioreactors (15 mL/bioreactor) as described by Copinet et al. [48].

### 2.3.1.2. Bacterial enrichment cultures for bioaugmentation.
For the bioaugmentation strategy, we used bacteria previously isolated in 2.2. During selective enrichment in minimal media, several bacterial species were recovered together within the same Erlenmeyer flasks, suggesting potential synergistic, mutualistic, or facilitative interactions among the co-isolated strains [50]. To preserve these putative positive interactions, the bioaugmentation strategy adopted in this study relied on the application of two mixed consortia to the bioreactors.

Two enrichment-derived inocula were then prepared and applied according to material pairing. Consortium (A), derived from PBS enrichment, was used for both PBS and PBS/OP tests, whereas consortium (B), derived from PBAT-based materials, was applied to both PBAT and PBAT/BSG tests. Table 3 summarizes the different strains that constitute these two extracts.

For each strain, a single cryopreserved bead (stored at -80 °C) was transferred into 5 mL of Nutrient Broth (NB) in a sterile 50 mL Falcon tube and incubated at 28 °C with shaking at 120 rpm for 48 h. An aliquot of 2 mL of this preculture was then used to inoculate 75 mL of fresh NB in a 500 mL Erlenmeyer flask, followed by incubation under identical conditions for a further 48 h. Viable cell counts were determined by serial dilution and plating on R2A agar (three replicates per condition). Each bioreactor assigned to the bioaugmentation treatment was inoculated with 15 mL of the final enrichment culture (in addition to 15 mL of

<table><caption>**Table 3**

Composition of the bacterial enrichment consortia used for bioaugmentation.</caption><thead><tr><td>Enrichment inocula</td><td>Isolation method</td><td>Identified strains</td></tr></thead><tbody><tr><td>PBS-based</td><td>Selective enrichment in M9 minimal medium using plastic fragments as the sole carbon source</td><td>Rhizobium radiobacter<br>Achromobacter<br>denitrificans/xylosoxidans<br>Stenotrophomonas maltophilia<br>Pseudomonas putida</td></tr><tr><td>PBAT-based</td><td>Selective enrichment in M9 minimal medium using plastic fragments as the sole carbon source</td><td>Pseudomonas aeruginosa<br>Pseudomonas sp.</td></tr></tbody></table>

compost extract described in 2.3.1.2.).

The PBS-derived enrichment culture ($7.05 \times 10^{11}$ CFU per 15 mL) was applied to the PBS and PBS/OP bioreactors, whereas the PBAT-derived enrichment culture ($1.67 \times 10^{11}$ CFU per 15 mL) was used for the PBAT and PBAT/BSG bioreactors. In all cases, these inocula were added to approximately 2 L of decomposition medium containing 17.5 g of plastic fragments.

The inoculation levels used in this study were intentionally set high to maximize the probability of functional establishment in the presence of indigenous compost-derived microbiota. They should therefore be interpreted as exploratory upper-bound experimental values rather than optimized operational targets for field deployment.

These inoculation levels were derived from mean viable cell counts of ($4.7 \pm 0.11 \times 10^{10}$ CFU·mL⁻¹ (n = 3; CV = 23.6 %)) for the PBS-derived enrichment and ($1.11 \pm 0.13 \times 10^{10}$ CFU·mL⁻¹ (n = 3; CV = 11.8 %)) for the PBAT-derived enrichment. The same enrichment cultures were also added to the blank bioreactors, in addition to the mature compost extract, although no plastic fragments were present. This procedure enabled an accurate correction for microbial respiration unrelated to polymer degradation and therefore ensured a reliable estimation of the net mineralization of the target substrates.

The organic carbon introduced by the enrichment inocula was limited to $\leq 60$ mg C per bioreactor, based on the standard formulation of the growth medium. Under typical incubation yields, 40–60 % of this carbon is expected to be mineralized during preculture [51], leaving approximately 25–35 mg C as residual biomass or dissolved organic matter. This low carbon input ensured that plastic fragments remained the primary carbon source available to the microbial communities. These details are summarized in Table 4.

### 2.3.1.3. Microbial pre-activation phase in bioreactors.
To better emulate mature compost conditions, a five-day preactivation step was performed prior to introducing the plastic specimens into the bioreactors. These conditions are characterized by a well-established and enzymatically active microbial community. This approach was adapted from methods previously described by Longieras et al. [52], who demonstrated that nutrient-assisted microbial preconditioning improves the comparability of mineralization kinetics obtained on activated vermiculite with those measured in mature compost.

The preactivation step aimed to stimulate microbial growth and to induce, at an early stage, specific enzymatic pathways involved in polymer degradation. On inert substrates such as vermiculite, which are devoid of native organic matter, microbial colonization and metabolic activation are generally delayed compared to compost-based systems [52,53]. Therefore, this strategy was designed to reduce the initial lag phase typically observed in such environments, ultimately allowing for
<caption>**Table 4**</caption>

Inoculation conditions applied to PBS, PBS/OP, PBAT, and PBAT/BSG under non-bioaugmented and bioaugmented conditions.

<table><thead><tr><th rowspan="2">Materials/Inocula</th><th colspan="3">Enriched inocula</th></tr><tr><th>Compost extract inoculum (mL)</th><th>Volume (mL)</th><th>CFU·mL<sup>-1</sup></th></tr></thead><tbody><tr><td><strong>Without bioaugmentation</strong></td><td>15</td><td>—</td><td>—</td></tr><tr><td>PBS</td><td>15</td><td>—</td><td>—</td></tr><tr><td>PBS/OP</td><td>15</td><td>—</td><td>—</td></tr><tr><td>PBAT</td><td>15</td><td>—</td><td>—</td></tr><tr><td>PBAT/BSG</td><td>15</td><td>—</td><td>—</td></tr><tr><td><strong>With bioaugmentation</strong></td><td>15</td><td>15</td><td>(4.7 ± 0.11) × 10<sup>10</sup></td></tr><tr><td>PBS+</td><td>15</td><td>15</td><td>(4.7 ± 0.11) × 10<sup>10</sup></td></tr><tr><td>PBS/OP+</td><td>15</td><td>15</td><td>(1.11 ± 0.13) × 10<sup>10</sup></td></tr><tr><td>PBAT+</td><td>15</td><td>15</td><td>(1.11 ± 0.13) × 10<sup>10</sup></td></tr><tr><td>PBAT/BSG+</td><td>15</td><td>15</td><td>(1.11 ± 0.13) × 10<sup>10</sup></td></tr></tbody><tfoot><tr><td colspan="4"><strong>Notes.</strong> CFU·mL<sup>-1</sup> values correspond to viable cell counts determined by serial dilution plating on R2A agar (mean ± SD).</td></tr></tfoot></table>

more comparable mineralization kinetics between activated vermiculite and mature compost conditions.

In this context, a preliminary four-week trial was undertaken to determine the activation dose most suitable for the plastic materials under investigation. This step was informed by Longieras [52], who demonstrated that both the quantity and composition of the activation dose critically influence mineralization kinetics on activated vermiculite, which closely mirror those observed in mature compost. Two activation doses (D₁ and D₂) (Table 5) were therefore tested under identical experimental conditions using a mature compost extract, the preparation of which is described in Section 2.3.2. Following these preliminary trials, the activation doses were adjusted, and two formulations were established and tailored to the polyester type (PBS or PBAT) and their respective biocomposites.

In the case of PBS and PBS/OP, the D₁ dose caused overstimulation of microbial activity and an accelerated biodegradation resembling a bio-stimulation effect, a phenomenon that was not observed for PBAT and PBAT/BSG. Each activation formulation combined Nutrient Broth (Sigma-Aldrich/Merck KGaA, Darmstadt, Germany) and cellulose microcrystalline (≤ 20 µm) (Sigma-Aldrich/Merck KGaA, Darmstadt, Germany) as general growth substrates to promote microbial proliferation. A small amount of the target material (PBS, PBAT, or their bio-composites) (0.5 g) was also added as the primary carbon source to induce the expression of specific depolymerases. Urea was also included as a nitrogen source to set the initial C/N ratio to ~25, a value considered optimal during the early stages of composting. This supplementation further supports microbial metabolism and enhances early microbial activation [54].

For the bioaugmented treatments, the microbial enrichment cultures were introduced at this stage, together with the nutrient additives and the mature compost extract, to ensure their early integration into the vermiculite medium. The detailed composition of each activation mixture is provided in Table 5.

Fig. 3 provides a schematic overview of the microbial inoculation strategy and the pre-activation phase, as detailed in 2.3.1.

<caption>**Table 5**</caption>

Composition of the two activation doses (D₁ and D₂) used in the biodegradation experiments.

<table><thead><tr><th>Activation doses</th><th>Cellulose (g)</th><th>Nutrient Broth (g)</th><th>Urea (g)</th><th>Target material (g)</th></tr></thead><tbody><tr><td>D<sub>1</sub></td><td>1.50</td><td>1.00</td><td>0.08</td><td>0.50</td></tr><tr><td>D<sub>2</sub></td><td>0.75</td><td>0.50</td><td>0.06</td><td>0.50</td></tr></tbody></table>

## 2.4. Analytical monitoring of materials during composting

Analytical monitoring of all tested materials was performed at weeks 3 and 9 after the start of the compostability assay. These time points were selected to capture distinct stages of the mineralization kinetics. Week 3 corresponds to the active growth phase, characterized by a near-linear increase in mineralization, whereas week 9 corresponds to the late stage of the process, encompassing either the plateau or the end of the deceleration phase (Figs. 5–6). At each sampling time point, two plastic fragments were randomly collected from each bioreactor, thoroughly rinsed with ultrapure deionized water, and oven-dried at 40 °C for 72 h prior to analysis.

### 2.4.1. Macroscopic images of plastic fragment samples

Photographs of the plastic specimens were taken at weeks 3 and 9 to document changes in their macroscopic appearance, with a focus on visible signs of disintegration. These observations were used to complement mineralization data by providing qualitative information on fragmentation and surface deterioration occurring during the biodegradation process.

### 2.4.2. Scanning electron microscopy (SEM)

The surfaces of samples collected after 3 and 9 weeks of composting were examined by scanning electron microscopy (SEM) using a JEOL JSM-IT100 InTouchScope™ microscope. Observations were carried out at an accelerating voltage of 11 kV in secondary electron detection (SED) mode, with the objective of identifying micrometer-scale morphological features indicative of surface degradation, such as cavities, cracks, and erosion.

### 2.4.3. Differential scanning calorimetry (DSC) measurements

Thermal properties of the materials were analyzed by differential scanning calorimetry (DSC 214 Polyma, Netzsch) to determine their degree of crystallinity. Sample masses ranged from 8 to 12 mg. For each bioreactor, two fragments were analyzed, yielding two analytical replicates per reactor and three experimental replicates per condition (one per respirometric chamber). In total, six DSC measurements were obtained for each material–inoculum combination (compost extract or compost extract with microbial enrichment).

For both PBS- and PBAT-based materials, the DSC program consisted of a first heating from room temperature to 200 °C at 10 °C·min⁻¹, followed by an isothermal step of 3 min at 200 °C and subsequent cooling to −65 °C. Argon was used as the purge gas at a flow rate of 40 mL·min⁻¹. Thermal parameters and crystallinity indices were extracted exclusively from the first heating scan, as it reflects the thermal and morphological state of the materials at the time of sampling from the bioreactors.

The crystallinity index (χ) was therefore calculated from the first heating cycle using the following expression:

$$ \chi (\%) = \frac{(\Delta H_m - \Delta H_{cc})}{\Delta H_m (1 - \emptyset)} \times 100 \quad (4) $$

where ΔH_m is the measured melting enthalpy, ΔH_{cc} is the measured cold crystallization enthalpy, ΔH_m corresponds to the theoretical standard melting enthalpy of a fully crystalline material, and Ø represents the estimated dry mass fraction of the fillers in the biocomposite (~8 %).

The reference values for ΔH_m used for each polymer were as follows: PBS (110.3 J/g), and PBAT (114 J/g) [55]. The DSC thermograms were analysed using the Proteus Analysis software, version 8.0 for DSC (NETZSCH-Gerätebau GmbH, Selb, Germany).

### 2.4.4. Fourier transform infrared spectroscopy (FTIR-ATR)

Processed plastic specimens (neat polymers and biocomposites) were characterized by Fourier Transform Infrared Spectroscopy (FTIR) using a Nicolet iS10 spectrometer (Thermo Scientific) equipped with an
<figure><img src="image_5.png" /><figcaption>Fig. 3. Schematic illustration of the inoculation and pre-activation procedure for the bioreactors.</figcaption></figure>

attenuated total reflectance (ATR) accessory (ZnSe crystal, Smart iTX™ ATR). For each measurement, 32 scans were collected at a resolution of 4 cm⁻¹ over the spectral range 4000–500 cm⁻¹, with a background spectrum recorded prior to each acquisition.

Five replicate measurements were performed on two independent plastic fragments per bioreactor, and the resulting spectra were averaged to obtain one analytical spectrum per reactor. These spectra were then averaged across the three experimental replicates (distinct respirometric chambers), yielding a total of 15 spectra per material–inoculum condition. Spectral processing was performed using OMNIC 8.1 software and included ATR correction, smoothing, and manual baseline adjustment.

For comparative analysis, spectra were normalized to the maximum intensity of the ester C=O stretching band, located at approximately 1713 cm⁻¹ for PBS-based materials and 1711.1 cm⁻¹ for PBAT-based materials. This normalization enabled qualitative comparison of spectral features (e.g. band appearance, disappearance, and shifts) as well as semi-quantitative assessment of relative intensity changes across time points (initial state, weeks 3 and 9) and between material–inoculum combinations [56].

## 2.5. Microbial monitoring during composting

Microbial community dynamics were assessed using two complementary approaches: colony-forming unit (CFU) enumeration on R2A agar plates and community-level physiological profiling (CLPP) using Biolog EcoPlates™. Both analyses were performed for all material–inoculum combinations, with and without microbial enrichment. Measurements were conducted at weeks 3 and 9 after the start of the composting experiment, using three independent bioreactors as experimental replicates for each condition.

### 2.5.1. Sampling of the DEgradation Medium and Preparation for Microbial Monitoring

At each sampling time point, 3 g of decomposition medium were collected from each of the 24 bioreactors, excluding any visible plastic fragments. Samples were stored in airtight containers at –20 °C until analysis. For microbial suspension preparation, 2 g of the sampled

vermiculite were transferred into 50 mL Falcon™ tubes containing 45 mL of sterile 0.9 % NaCl solution and vortexed for 3 min at room temperature (TopMix FB 15024, Thermo Fisher Scientific) to detach and resuspend the associated microbial cells.

### 2.5.2. CFU enumeration in vermiculite

The microbial suspensions prepared as described above were subjected to tenfold serial dilutions up to 10⁻⁸ using sterile 0.9 % NaCl solution in 1.7 mL Eppendorf® tubes. Each dilution step was homogenized by vortexing, and 100 µL aliquots were spread onto R2A agar plates under sterile conditions. Plates were incubated in the dark at 24 ± 1 °C for 48 h. Colony-forming units (CFU) were manually counted on plates containing between 30 and 300 colonies. All counts were performed in triplicate and expressed as viable microbial abundance in CFU.g⁻¹ of wet composting medium.

### 2.5.3. Biolog ecolplates

Community-level physiological profiling (CLPP) was performed using Biolog EcoPlates™ (96-well microplates) containing 31 distinct carbon substrates, each supplied in triplicate, along with triplicate water-only control wells. Substrate utilization was assessed based on color development resulting from the reduction of the tetrazolium-based redox dye present in each well [57–59].

The microbial suspension described in 2.5.1 was centrifuged at 1000 rpm for 5 min (Eppendorf® 5810 R, Hamburg, Germany). A 7 mL aliquot of the resulting supernatant was subsequently diluted with 33 mL of sterile 0.9 % NaCl solution to prepare the working inoculum for EcoPlate™ analysis.

Prior to inoculation, the diluted suspension was homogenized by vortexing for 1 min. Then, 150 µL was dispensed into each well using a multichannel pipette. The EcoPlates™ were immediately read at 590 nm using a Biolog MicroStation® (Biotek Instruments, Winooski, VT, USA) to establish baseline absorbance values (T₀) and subsequently incubated in the dark at 24 ± 1 °C. Optical density measurements were recorded again after 72 h of incubation (T + 72 h).

The 72-h incubation time was selected as an optimal compromise. This duration is sufficient to allow measurable metabolic activity across a broad range of carbon substrates, while minimizing bias associated
with differences in initial inoculum density [60,61]. In contrast, extending the incubation period beyond 72 h may introduce artefacts, as microbial communities can progressively adapt to the specific substrates present in individual wells. Such adaptive shifts may lead to community restructuring and divergence of optical density signals from the intrinsic functional physiological profiles of the original bioreactor communities targeted in this study [62]

## 2.6. Data analysis

All statistical analyses were performed using R software (version 4.5.1). Data were first evaluated to verify compliance with the assumptions of linear models, including normality of residuals and homogeneity of variances. When these assumptions were satisfied, one-way analysis of variance (ANOVA) was applied. Significant effects were further explored using Tukey's Honest Significant Difference (HSD) post hoc test for multiple comparisons.

Pairwise comparisons between incubation times were assessed using Student's t-test when normality and variance homogeneity were met. In cases where these assumptions were violated, the non-parametric Mann-Whitney U test was used instead. For all analyses, statistical significance was defined at a threshold of $\alpha = 0.05$.

### 2.6.1. Nonlinear empirical modelling of mineralization kinetics

Mineralization kinetics were modelled using a nonlinear empirical regression based on the Hill equation. This model was selected because it adequately captures the sigmoidal behaviour typically observed in composting systems and provides a robust framework for describing and comparing mineralization kinetics across conditions [63].

The Hill model expresses the temporal evolution of the mineralization extent, $Y(t)$, as follows:

$$Y(t) = Y_{\max} \cdot \frac{t^n}{t^n + K^n} \qquad (5)$$

In this equation, $Y(t)$ represents the mineralization extent at time $t$. $Y_{\max}$ corresponds to the predicted asymptotic maximum mineralization, expressed as a percentage. The parameter $K$ is a scale parameter that defines the time required to reach half of $Y_{\max}$, while $n$ is a shape parameter controlling the steepness of the sigmoidal curve.

Model performance was evaluated using two statistical indicators: the coefficient of determination ($R^2$) and the root mean square error (RMSE). The coefficient of determination was calculated as follows:

$$R^2 = 1 - \frac{\sum (Y_{\text{obs}} - Y_{\text{pred}})^2}{\sum (Y_{\text{obs}} - \bar{Y}_{\text{obs}})^2} \qquad (6)$$

The RMSE was used to quantify the average prediction error, expressed as a percentage:

$$RMSE (\%) = \sqrt{\frac{1}{n} \sum (Y_{\text{obs}} - Y_{\text{pred}})^2} \qquad (7)$$

### 2.6.2. Biolog EcoPlates® data analysis

#### 2.6.2.1. Average well colour development (AWCD)
The average well colour development (AWCD) was calculated as an index of the overall metabolic activity of the microbial communities [64], according to the following equation:

$$AWCD = \left(\frac{1}{31}\right) \times \sum_{i=1}^{31} OD_i \qquad (8)$$

where $OD_i$ corresponds to the corrected optical density of each substrate well, calculated as the absorbance measured at $T + 72$ h minus both the absorbance at $T_0$ and that of the water control. Slightly negative values resulting from background correction were set to zero to avoid bias in subsequent analyses.

#### 2.6.2.2. Functional diversity indices
Microbial functional diversity was assessed using three complementary indices: substrate richness ($S$), the Shannon diversity index ($H$), and the Simpson diversity index ($D$).

Substrate richness [57] was defined as:

$$S = \sum_{i=1}^{31} I (OD_i > 0.25) \qquad (9)$$

where $OD_i$ is the blank-corrected absorbance of the ith substrate, and $I$ is an indicator function equal to 1 when the condition is met and 0 otherwise. Accordingly, $S$ represents the number of substrates significantly metabolized by the microbial community.

The Shannon index [65] was calculated as:

$$H = - \sum_{i=1}^{31} p_i \ln(p_i) \qquad (10)$$

where $p_i$ corresponds to the relative contribution of substrate $i$, defined as the absorbance of a given well normalized by the total absorbance across the 31 carbon substrates of the EcoPlate™:

$$p_i = \frac{OD_i}{\sum_{j=1}^{31} OD_j} \qquad (11)$$

Higher values of $H$ indicate greater diversity and a more even distribution of metabolic activities.

The Simpson index [65] was calculated as:

$$D = 1 - \sum_{i=1}^{31} p_i^2 \qquad (12)$$

where $p_i$ is defined as above. Compared to Shannon index, the Simpson index gives more weight to the most abundant categories and ranges from 0 (no diversity) to 1 (maximum diversity).

All indices were computed using absorbance data after blank correction, following established procedures for CLPP analyses [57,58].

#### 2.6.2.3. Principal component analysis (PCA)
Principal component analysis (PCA) was applied to explore potential clustering of samples based on their metabolic fingerprints. The analysis was performed using the mean utilization values of the 31 carbon substrates, which were grouped into six biochemical families: carbohydrates (CH), complex carbon sources (CCS), carboxylic acids (CA), amino acids (AA), amines (A), and phosphorylated carbon sources (PC). The detailed composition of each biochemical family is provided in Table 6.

Separate PCAs were performed for Weeks 3 and 9 to capture distinct stages of the composting process. For each analysis, both the correlation circle and the individual factor map were examined.

**Table 6**

Grouping of the 31 EcoPlate™ carbon substrates into six biochemical families used for PCA analysis.

<table><thead><tr><th>Biochemical families</th><th>Carbon substrates</th></tr></thead><tbody><tr><td>Carbohydrates (CH)</td><td>β-Methyl-D-Glucoside; D-Xylose; i-Erythritol; D-mannitol; N-Acetyl-D-Glucosamine; D-Cellobiose; α-D-Lactose</td></tr><tr><td>Amines (A)</td><td>Phenylethyl-Amine; Putrescine</td></tr><tr><td>Complex carbon sources (CCS)</td><td>Tween 40; Tween 80; α-Cyclodextrin; Glycogen</td></tr><tr><td>Carboxylic acids (CA)</td><td>Pyruvic Acid Methyl Ester; D-Glucosaminic Acid; D-Galactonic Acid γ-Lactone; D-Galacturonic Acid; 2-Hydroxy Benzoic Acid; 4-hydroxybenzoic acid; γ-Hydroxybutyric Acid; Itaconic Acid; α-Ketobutyric Acid; D-Malic Acid</td></tr><tr><td>Amino acids (AA)</td><td>L-Arginine; L-Asparagine; L-Phenylalanine; L-Serine; L-Threonine; Glycyl-L-Glutamic Acid</td></tr><tr><td>Phosphorylated carbon sources (PC)</td><td>Glucose-1-Phosphate; D,L-α-Glycerol Phosphate</td></tr></tbody></table>
Together, these analyses enabled the identification of distinct func-
tional metabolic profiles associated with different materials and inocu-
lation conditions at each composting stage.

**3. Results and discussion**

3.1. Mineralisation kinetics

The mineralization kinetics of the thermoplastic materials and the positive reference (microcrystalline cellulose), together with the corresponding first-derivative curves representing daily mineralization rates for PBS- and PBAT-based materials, are shown in Figs. 4–6.

These curves illustrate the mineralization behaviour of the tested
materials under two inoculation conditions. The first corresponds to
standard inoculation using mature compost extract only, while the
second represents a bioaugmented condition in which the compost
extract was supplemented with a selected bacterial consortium
composed of previously isolated and identified strains (Table 3).

For the positive reference, mineralization was assessed exclusively under standard inoculation conditions. Microcrystalline cellulose reached the 70 % mineralization threshold required by the French standard for home-compostable plastic products (NF T51–800) after 29 days of incubation. This rapid mineralization confirms the high biological activity of the compost inoculum and demonstrates that the experimental setup was appropriate for evaluating the biodegradation performance of the tested materials under mesophilic composting conditions [25,66].

3.1.1. PBS- based materials

The mineralization kinetics of PBS-based materials (pristine PBS and PBS/OP) are shown in Fig. 5. Under standard inoculation conditions, pristine PBS reached a final mineralization extent of 43.6 % after 133 days. In contrast, bioaugmentation (PBS+) resulted in a final mineralization of 73.4 %, corresponding to an average increase of + 68.3 %. This marked enhancement demonstrates that inoculation with selected microbial strains significantly improved PBS biodegradation under mesophilic composting conditions. These strains were previously isolated through a selective enrichment procedure using minimal medium.

Analysis of the first-derivative curves (Fig. 5), which represent daily
mineralization rates, indicates that both inoculation conditions followed
comparable kinetics during the initial phase (up to approximately day
5). Beyond this point, the daily mineralization rate of non-bioaugmented
PBS progressively declined, whereas that of the bioaugmented system
continued to increase, reaching a maximum around day 30 before
gradually decreasing. The two profiles converged after approximately
50 days, suggesting that bioaugmentation primarily contributed to
sustaining microbial activity during the early and intermediate stages of

biodegradation.

While the temporal shape of the mineralization kinetics was gener-
ally reproducible across replicates (latency, growth, deceleration, and
plateau phases), substantial variability was observed in the final
mineralization extents. A quantitative summary of final mineralization
levels and associated dispersion is provided in Table 7.

The reproducibility of the bioaugmentation effect on pristine PBS is illustrated in Fig. 7, which presents the mineralization kinetics obtained from the three biological replicates. Two of the three bioaugmented replicates reached high mineralization extents (85.5 % and 93.4 %), whereas the third replicate reached only 41.4 %, a value comparable to that of the non-enriched control. All controllable experimental parameters were strictly standardized across replicates, including the enrichment culture, inoculation volume, polymer mass, vermiculite substrate, moisture content, aeration, and incubation temperature. This high level of experimental control strongly limits the likelihood that the observed variability arises from experimental artefacts or uneven inoculation.

In this respect, the partial reproducibility observed is consistent with
the well-documented stochastic nature of microbial community assem-
bly in composting-like systems. In such environments, functional out-
comes can be shaped by dispersal limitation, ecological drift, and
priority effects, even under nominally identical operating conditions
[67,68]. Ecological drift arises from stochastic birth–death events at low
effective population sizes. Dispersal limitation restricts microbial access
to specific micro-habitats despite identical inoculum inputs. Together,
these processes can create early stochastic differences in microbial
attachment and surface colonization, promote priority effects, and ul-
timately lead to divergent degradation trajectories among biological
replicates, even when inocula are applied simultaneously.

In line with this interpretation, plastisphere studies under compost-
ing conditions have shown that stochastic processes may remain domi-
nant in shaping plastic-associated microbiota, with substantial
contributions of drift- and dispersal-related mechanisms [69]. More
generally, composting microbiomes have been described as highly sto-
chastic and difficult to control, reflecting strong microscale heteroge-
neity and complex interacting environmental drivers [70]. Similar
patterns have also been reported in engineered bioreactor biofilms,
where stochastic assembly dominates community construction [71].

Accordingly, the PBS+ variability observed here is plausibly
explained by probabilistic early establishment of the introduced con-
sortium within each bioreactor, combined with variable competitive
interactions with the resident compost-derived microbiota [71,72].
Several mitigation strategies could therefore be envisaged to reduce
inter-replicate variability. These include optimization of inoculation
density, as well as the implementation of repeated or staged inoculation
during the early incubation phase to increase the probability of effective
surface colonization. In addition, coupling bioaugmentation with mild

<figure><img src="image_6.png" /><figcaption>**Fig. 4.** Left: Evolution of the mineralization extent of microcrystalline cellulose using compost extract as the sole inoculum. Right: Daily mineralization rate (first derivative of the mineralization curve).</figcaption></figure>
Mineralization kinetics of PBS-based materials under mesophilic composting conditions (28 °C)

<figure><img src="image_7.png" /><figcaption>**Fig. 5.** Left: Evolution of the mineralization extent of PBS-based materials (pristine PBS and PBS/OP biocomposite) under compost extract and compost extract supplemented with bacterial enrichment (bioaugmentation). Right: Daily mineralization rate (first derivative of the mineralization curves). The horizontal red dashed line indicates the mineralization threshold required to comply with the NF T51-800 compostability standard.</figcaption></figure>

Mineralization kinetics of PBAT-based materials under mesophilic composting conditions (28 °C)

<figure><img src="image_8.png" /><figcaption>**Fig. 6.** Left: Evolution of the mineralization extent of PBAT-based materials (pristine PBAT and PBAT/BSG biocomposite) under compost extract and compost extract supplemented with bacterial enrichment (bioaugmentation). Right: Daily mineralization rate (first derivative of the mineralization curves). The horizontal red dashed line indicates the mineralization threshold required to comply with the NF T51-800 compostability standard.</figcaption></figure>

biostimulation may further promote the rapid establishment and functional expression of the introduced microbial guilds [73,74].

Moreover, it is worth noting that when only the two replicates exhibiting a clear bioaugmentation effect are considered, the mean mineralization extent reached 89.5 % after 133 days. This value is very close to the 90 % threshold required by decentralized-compostability

standards.

Nonlinear regression using the Hill model confirmed the positive effect of bioaugmentation on PBS biodegradation (Table 8). The predicted maximum mineralization ($Y_{max}$) increased from 47.9 % for PBS to 76.3 % for PBS+. In addition, the K parameter, corresponding to the time required to reach 50 % of $Y_{max}$, decreased from 42 to 32 days,
<table><caption>**Table 7**
Final mineralization of plastic materials after 133 days.</caption><thead><tr><th>Materials/Inocula</th><th>Final mineralization (%)</th><th>SD (%)</th><th>CV (%)</th></tr></thead><tbody><tr><td><strong>PBAT-based</strong></td><td>5.9</td><td>1.7</td><td>27.9</td></tr><tr><td>PBAT</td><td></td><td></td><td></td></tr><tr><td>PBAT+</td><td>7.7</td><td>3.7</td><td>47.9</td></tr><tr><td>PBAT/BSG</td><td>23.7</td><td>2.1</td><td>9.0</td></tr><tr><td>PBAT/BSG+</td><td>25.2</td><td>1.0</td><td>4.0</td></tr><tr><td><strong>PBS-based</strong></td><td>43.6</td><td>5.2</td><td>12.0</td></tr><tr><td>PBS</td><td></td><td></td><td></td></tr><tr><td>PBS+</td><td>73.4</td><td>28.0</td><td>38.1</td></tr><tr><td>PBS/OP</td><td>79.9</td><td>10.9</td><td>13.7</td></tr><tr><td>PBS/OP+</td><td>75.6</td><td>20.8</td><td>27.5</td></tr></tbody></table>

**Notes.** SD: standard deviation; CV: coefficient of variation, calculated as $(SD / mean) \times 100$

indicating accelerated early-stage biodegradation under bioaugmented conditions. The higher Hill coefficient (n) for PBS+ (2.71 vs. 2.42 for PBS) further indicates a steeper transition from the initial to the active degradation phase, suggesting a more synchronized microbial response once degradation was initiated [75,76].

The enhanced biodegradation of PBS under bioaugmented conditions can be related to known functional traits of the strains present in the enrichment consortium (Table 3). *Pseudomonas putida*, for example, can metabolize 1,4-butanediol, a key PBS hydrolysis product, through multiple metabolic pathways [77]. More broadly, *Pseudomonas* spp. are well known for expressing extracellular esterases and lipases involved in polyester chain scission, as well as for their high metabolic versatility [78,79]. Other consortium members, such as *Stenotrophomonas maltophilia* and *Achromobacter xylosoxidans*, have also been reported to contribute to polymer surface colonization and degradation processes under mesophilic conditions [80,81].

Beyond single-strain traits, the four-strain PBS consortium can be interpreted within a “metabolic division of labour” framework, in which complementary functions collectively enhance polymer-to-CO₂ conversion. In such consortia, some members primarily support surface colonization and extracellular chain scission, whereas others preferentially assimilate soluble oligomers and monomers, thereby limiting product accumulation and maintaining favourable thermodynamic and kinetic conditions for sustained hydrolysis. This functional partitioning and cross-feeding is a well-established mechanism underlying the stability and efficiency of multi-member degradation communities [82,83].

Within this framework, it can be hypothesized that at the polymer-biofilm interface, *Pseudomonas* spp. primarily contributes through robust biofilm formation and extracellular esterase/lipase activities, promoting initial ester-bond cleavage and sustained surface-associated hydrolysis [84,85]. In the present consortium, *Pseudomonas putida* may additionally act as a sink for soluble PBS hydrolysis products, while

*Stenotrophomonas maltophilia*, *Achromobacter* spp., and *Rhizobium/Agrobacterium*-related taxa likely play supportive roles in surface colonization, biofilm maturation, and complementary esterolytic activity [86, 87].

We emphasize that this interpretation remains a working hypothesis, and that resolving strain-level functional contributions would require dedicated assays, such as controlled co-culture experiments, enzyme activity profiling, and expression-based approaches.

For the PBS/OP biocomposite, the final mineralization extent reached 79.9% after 133 days under standard inoculation conditions (compost extract only). Under a purely additive scenario, the expected mineralization extent would therefore be 49.2%. The experimentally observed value exceeds this prediction by + 30.7%, corresponding to a + 62.3% synergistic enhancement beyond the intrinsic biodegradability of the filler alone. This marked deviation from additivity indicates that the presence of the orange peel filler does not merely contribute as an additional biodegradable fraction but actively modifies the degradation behaviour of the composite material.

To elucidate the physical mechanisms underlying this synergistic effect, Fig. 8 presents water vapor sorption isotherms measured at 28 °C by dynamic vapor sorption (DVS) for orange peel, pristine PBS, and the PBS/OP biocomposite over a relative humidity range of 10–90%. As discussed in detail in our previous work [29], orange peel exhibits a markedly higher water sorption capacity than pristine PBS, reflecting its hydrophilic and porous nature. Incorporation of this filler into the PBS matrix therefore leads to a substantial increase in the water sorption capacity of the biocomposite. This enhanced water uptake is expected to promote swelling of the filler phase through water absorption under

<table><caption>**Table 8**
Parameters estimated from the nonlinear Hill model fitted to mineralization kinetics for each material-inoculum combination.</caption><thead><tr><th>Materials/Inocula</th><th>Y<sub>max</sub> (%)</th><th>K (days)</th><th>n</th><th>R<sup>2</sup></th><th>RMSE (%)</th></tr></thead><tbody><tr><td>Cellulose</td><td>76.4</td><td>7.88</td><td>2.05</td><td>0.994</td><td>1.23</td></tr><tr><td><strong>PBAT-based</strong></td><td>6.75</td><td>14.60</td><td>0.89</td><td>0.996</td><td>0.07</td></tr><tr><td>PBAT</td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>PBAT+</td><td>9.00</td><td>17.16</td><td>0.87</td><td>0.994</td><td>0.12</td></tr><tr><td>PBAT/BSG</td><td>25.1</td><td>19.71</td><td>1.44</td><td>0.998</td><td>0.25</td></tr><tr><td>PBAT/BSG+</td><td>27.3</td><td>21.70</td><td>1.41</td><td>0.999</td><td>0.24</td></tr><tr><td><strong>PBS-based</strong></td><td>47.9</td><td>42.42</td><td>2.42</td><td>0.994</td><td>1.20</td></tr><tr><td>PBS</td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>PBS+</td><td>76.3</td><td>32.19</td><td>2.71</td><td>0.998</td><td>1.22</td></tr><tr><td>PBS/OP</td><td>85.2</td><td>38.87</td><td>2.57</td><td>0.997</td><td>1.56</td></tr><tr><td>PBS/OP+</td><td>79.7</td><td>35.58</td><td>2.48</td><td>0.999</td><td>0.98</td></tr></tbody></table>

**Notes.** Ymax (%) is the predicted maximum extent of mineralization; K (days) corresponds to the time required to reach 50% of Ymax; n is the Hill coefficient describing curve steepness. RMSE (%) denotes the root mean square error of the model fit.

<figure><img src="image_9.png" /><figcaption>**Pristine PBS mineralization - individual replicates under mesophilic composting conditions (28 °C)**</figcaption></figure>

**Fig. 7.** Evolution of the mineralization extent across the three replicates of pristine PBS, illustrating the variability of treatment effects. Non-bioaugmented PBS (compost extract only) is shown on the left, and bioaugmented PBS on the right.
<figure><img src="image_10.png" /><figcaption>Fig. 8. Water sorption isotherms measured at 28 °C for agro-industrial by-products (OP, BSG), neat PBS-, PBAT-, and their corresponding biocomposites. C<sub>eq</sub> denotes the equilibrium concentration of permeant molecules (g g<sup>-1</sup>) at a given water activity (a<sub>w</sub>).</figcaption></figure>

decomposition conditions. The resulting volumetric expansion is likely
to generate mechanical stress within the polymer matrix. Such stress
facilitates the formation of microcracks and progressive fragmentation
of the material, as observed by SEM at the surface of the biocomposites
but not in the pristine polymers at weeks 3 and 9 after the onset of
composting (Figs. 9–12).

Taken together, these physical and morphological changes provide a coherent mechanistic basis for the pronounced synergistic effect observed in PBS/OP. This effect is therefore primarily attributed to enhanced physical and morphological destabilization of the bio-composite during the early stages of composting. This destabilization increases the effective accessibility of PBS chains to microbial enzymes, microorganisms, and water diffusion. As a result, chain scission and the subsequent microbial bio-assimilation of lower-molecular-weight compounds are promoted [88,89]. This interpretation is further supported by macroscopic observations (Fig. 13), which reveal early surface erosion and fragmentation of PBS/OP, whereas pristine PBS remained visually intact under the same incubation conditions.

Importantly, this synergistic enhancement resulting from the incor-
poration of the organic filler cannot be ascribed to intrinsic differences
in the initial molecular characteristics or structural state of the polymer
matrix. Indeed, the initial molecular weight of the polyester matrix in
the biocomposite (M<sub>w</sub> = 143 kDa; M<sub>n</sub> = 53.3 kDa; D̅ = 2.68) is very
similar to that of pristine PBS (M<sub>w</sub> = 146.7 kDa; M<sub>n</sub> = 54 kDa; D̅ = 2.72).
This similarity indicates that neither the absolute molecular weight nor
its distribution can account for the substantially higher mineralization
observed for the biocomposite [90]. In addition, the crystallinity degrees
of PBS and PBS/OP are very close (Table 1), indicating that differences
in crystalline-amorphous phase balance are unlikely to drive the
observed enhancement in biodegradation.

By contrast, bioaugmentation did not further enhance the biode-
gradability of PBS/OP, as the final mineralization level under bio-
augmented conditions (75.6 %) remained comparable to that obtained
under standard inoculation. This absence of an additive effect can be
rationalized by considering the biochemical nature of the orange-peel
filler. Indeed, in composting environments, biodegradation outcomes
are not governed solely by the functional potential of the microbial
community but are strongly modulated by the availability and degrad-
ability of carbon substrates, which drive microbial succession and
functional dominance over time [91].

In the case of PBS/OP, this general ecological framework is directly

relevant given the intrinsic biochemical composition of the orange-peel
filler. As reported in detail in our previous study [29], the orange peel
filler used here consists predominantly of water-soluble compounds
(80.1 ± 0.2 wt%), together with cellulose (15.3 ± 0.3 wt%) and hemi-
cellulose (3.9 ± 0.4 wt%). Moreover, the water-soluble fraction is re-
ported in the literature to be largely composed of readily hydrolysable
and microbially accessible substrates, such as simple sugars, organic
acids, and proteins. These compounds are efficiently degraded and
metabolized by microbial communities under mesophilic aerobic con-
ditions [92].

In contrast, PBS is a high molecular weight polyester that requires
specific enzymatic adaptation before efficient chain scission can occur.
Several composting studies have shown that, in biocomposite systems,
organic fillers composed of low complexity substrates are preferentially
utilized during the early stages of incubation. In contrast, polymer
degradation typically exhibits a longer lag phase associated with mi-
crobial adaptation to more complex substrates [93].

Nevertheless, the earlier degradation of the OP filler is not directly
demonstrated in the present study. It is instead proposed as a working
hypothesis to interpret the observed biodegradation trends. Confirming
this hypothesis would require targeted analyses of filler and polymer
degradation over time, which are beyond the scope of the present work.

The likely rapid availability of these labile substrates could prefer-
entially stimulate indigenous heterotrophic populations, thereby shift-
ing biodegradation control toward bottom-up, resource-driven
processes rather than the expression of specialized polymer-degrading
functions [94]. Under such conditions, this early stimulation may
further limit the establishment and functional expression of the intro-
duced strains through competitive exclusion and priority effects during
the initial stages of surface colonization [95].

This context-dependent “window of efficacy” of bioaugmentation
remains hypothetical. However, it is in line with broader literature
showing that biodegradation outcomes emerge from the interplay be-
tween polymer properties, substrate availability, and microbial ecolog-
ical dynamics [91,96,97].

3.1.2. PBAT- based materials

In the case of PBAT-based materials, microbial enrichment did not
substantially improve the mineralization of the co-polyester. Pristine
PBAT reached a final mineralization extent of 5.9 %, compared to 7.7 %
for the enriched condition (PBAT+). Similarly, the PBAT/BSG
PBS-based materials - SEM images (x60)

<figure><img src="image_11.png" /><figcaption>Fig. 9. SEM images of the surface of PBS-based materials collected at weeks 3 and 9 of the biodegradation experiment, under both inoculation conditions (bio-augmented and non-bioaugmented), at a magnification of × 60.</figcaption></figure>

biocomposite mineralized up to 23.7 % under standard inoculation (i.e. mature compost extract only), versus 25.2 % for its enriched counterpart (PBAT/BSG+). Overall, these marginal differences indicate that the enrichment strategy did not translate into a measurable enhancement of PBAT mineralization under the tested mesophilic conditions, despite the apparent suitability of the microbial selection approach.

To clarify this apparent discrepancy, it is important to recall the
rationale underlying the enrichment procedure. The microbial con-
sortium was obtained through functional isolation under strictly mineral
conditions (M9 medium), in the dark, using PBAT-based materials as the
sole carbon and energy source. This selection strategy ensures that the
retained microorganisms can exploit at least part of the material in the
absence of alternative organic substrates, thereby providing functional
evidence of microbial interaction with PBAT-related components.

However, although the PBAT-enrichment consortium comprised
metabolically versatile *Pseudomonas* strains, including *Pseudomonas
aeruginosa* [98], which have been reported to enhance PBAT degrada-
tion under controlled liquid-culture conditions [99]. This potential did
not translate into a measurable increase in mineralization in the present

non-sterile system.

Given the intrinsically heterogeneous structure of PBAT as an ali-
phatic-aromatic copolymer, preferential utilization of the more acces-
sible and hydrolytically labile aliphatic butylene adipate segments is
expected. In contrast, the aromatic terephthalate units are likely to
remain largely recalcitrant under mesophilic conditions at 28 °C,
potentially imposing a kinetic limitation on overall PBAT biodegrada-
tion through the persistence of its aromatic fraction [100]. In line with
this interpretation, several studies have reported that PBAT chain scis-
sion is predominantly driven by abiotic processes and is strongly
accelerated at temperatures approaching its melting point (T<sub>m</sub> ≈ 120
°C). In contrast, chemical hydrolysis under mesophilic conditions re-
mains very slow, thereby limiting subsequent microbial assimilation
[101].

This interpretation is further supported by FTIR trends showing a
progressive relative increase in the intensity of aromatic signatures in
the 900–700 cm⁻¹ region during composting (Figs. 16–17). Such spec-
tral evolution is consistent with a relative surface enrichment in tere-
phthalate units resulting from the preferential degradation of aliphatic
PBS-based materials - SEM images (x200)

<figure><img src="image_12.png" /><figcaption>**Fig. 10.** SEM images of the surface of PBS-based materials collected at weeks 3 and 9 of the biodegradation experiment, under both inoculation conditions (bio-augmented and non-bioaugmented), at a magnification of × 200.</figcaption></figure>

domains. It supports the hypothesis that PBAT degradation is kinetically constrained by the persistence of its aromatic segments. Importantly, the temporal evolution of these aromatic markers was nearly identical under bioaugmented and non-bioaugmented conditions. This similarity suggests that bioaugmentation did not overcome the intrinsic resistance of the aromatic PBAT segments and that the overall process was primarily limited by the lack of enzymatic potential capable of degrading and metabolizing the aromatic fraction under the experimental conditions investigated.

Taken together, these observations suggest that the microbial con-
sortium selected under mineral conditions can exploit the more readily
hydrolysable aliphatic domains of PBAT. In contrast, it does not signif-
icantly enhance the transformation of the aromatic terephthalate rich
fraction under mesophilic conditions. Nevertheless, this interpretation
remains a working hypothesis.

Addressing this question would require targeted follow up studies,
such as degradation assays in simplified liquid systems, enzymatic tests
on model compounds, and detailed structural characterization of PBAT
by NMR spectroscopy.

For the PBAT/BSG biocomposite containing 10 wt% milled brewer's
spent grain, the final mineralization extent reached 23.7 % after 133
days under standard inoculation conditions. Under a purely additive
scenario, the expected mineralization extent would therefore be 15.3%.
The experimentally observed value exceeds this prediction by + 8.4 %,
corresponding to a + 54.8 % synergistic enhancement beyond the
intrinsic biodegradability of the filler alone.

The physicochemical characteristics of the BSG filler have been re-
ported in detail by Bellon et al. [29]. Its biochemical composition con-
sists mainly of water-soluble compounds (14.4 ± 0.1 wt%), cellulose
(49.0 ± 0.5 wt%), and hemicellulose (24.6 ± 0.4 wt%).

As previously discussed for PBS/OP, incorporation of a hydrophilic
organic filler increases water uptake, as evidenced by the water vapor
sorption isotherms (Fig. 8). This enhanced sorption likely induces local
swelling of the filler phase. The resulting volumetric expansion is ex-
pected to generate mechanical stress at the polymer-filler interface.

Such stress promotes early surface cracking, fragmentation, and
progressive physical disintegration of the material, as observed by SEM
during composting (Figs. 9–12). These morphological changes increase
PBAT-based materials - SEM images (x60)

<figure><img src="image_13.png" /><figcaption>**Fig. 11.** SEM images of the surface of PBAT-based materials collected at weeks 3 and 9 of the biodegradation experiment, under both inoculation conditions (bioaugmented and non-bioaugmented), at a magnification of × 60.</figcaption></figure>

the effective surface area accessible to water diffusion, extracellular
hydrolytic enzymes, and microorganisms.

Importantly, this synergistic enhancement cannot be explained by
differences in the initial molecular or structural characteristics of the
polymer matrix. The molecular weight of PBAT in the biocomposite (M<sub>w</sub>
= 120.1 kDa; M<sub>n</sub> = 50.5 kDa; D̄ = 2.38) is comparable to that of pristine
PBAT (M<sub>w</sub> = 151.0 kDa; M<sub>n</sub> = 59.7 kDa; D̄ = 2.53). These differences are
insufficient to account for the markedly higher mineralization observed
for PBAT/BSG.

Similarly, crystallinity effects can be ruled out. Both PBAT and PBAT/BSG exhibit low and overlapping crystallinity degrees (6.0 ± 5.0 % and 9.4 ± 5.0 %, respectively; Table 1), indicating that differences in crystalline–amorphous phase balance are unlikely to drive the observed synergistic effect.

Therefore, the enhanced mineralization observed for the PBAT/BSG
biocomposite relative to pristine PBAT is primarily attributed to
enhanced physical disintegration of the material during the early stages
of composting, rather than to intrinsic changes in the chemical or
structural state of the polymer matrix.

Such fragmentation increases the surface-to-volume ratio and

improves the accessibility of PBAT chains to extracellular hydrolytic
enzymes, thereby facilitating the microbial assimilation of lower-
molecular-weight degradation products [88,89].

This interpretation is further supported by macroscopic observations (Fig. 13), which show earlier and more pronounced surface erosion of PBAT/BSG fragments compared with pristine PBAT.

Nonlinear Hill modelling provides further insight into the degrada-
tion kinetics of PBAT-based materials (Table 8). For pristine PBAT, the
predicted maximum mineralization remained very low (Ymax =
6.75–9.00 %), and bioaugmentation had only a marginal effect. This is
consistent with the limited mineralization observed experimentally. The
K parameter showed only minor variations between PBAT and PBAT+,
indicating no acceleration of early-stage degradation under bio-
augmented conditions.

In contrast, incorporation of brewer's spent grain markedly
increased the predicted maximum mineralization. For PBAT/BSG and
PBAT/BSG+, Ymax reached approximately 25-27%, reflecting both the
intrinsic biodegradability of the filler and enhanced accessibility of the
polymer matrix induced by composite disintegration.

Furthermore, the Hill coefficient (n) was below unity for both PBAT
PBAT-based materials - SEM images (x200)

<figure><img src="image_14.png" /><figcaption>**Fig. 12.** SEM images of the surface of PBAT-based materials collected at weeks 3 and 9 of the biodegradation experiment, under both inoculation conditions (bioaugmented and non-bioaugmented), at a magnification of × 200.</figcaption></figure>

and PBAT⁺ (n ≈ 0.9), reflecting a very gradual and weakly coordinated degradation process. Such values indicate the absence of a synchronized or cooperative biodegradation phase and are consistent with kinetically constrained degradation dominated by slow hydrolysis and sporadic microbial assimilation.

In contrast, incorporation of brewer's spent grain substantially modified the kinetic signature. For PBAT/BSG and PBAT/BSG+, n increased to values above unity (n ≈ 1.4), indicating a steeper transition toward the active degradation phase. This shift suggests partial kinetic structuring induced by filler-driven physical effects, such as enhanced water uptake and fragmentation, rather than by bioaugmentation itself.

Overall, Hill modelling highlights a clear distinction between pristine PBAT, which remains weakly reactive and poorly synchronized, as reflected by n values below unity. In contrast, PBAT/BSG composites exhibit moderate cooperativity (n > 1), indicating that filler-induced physical destabilization partially structures the degradation process without overcoming the intrinsic limitations imposed by the aromatic PBAT fraction.

## 3.2. Material monitoring during biodegradation

### 3.2.1. Macroscopic and SEM observations during biodegradation

Macroscopic photographs and SEM images of the surfaces of material fragments collected from biological replicate n°2 at weeks 3 and 9 of the biodegradation experiment are presented in Figs. 9–13. These images were used to assess changes in surface morphology, structural integrity, and the extent of visible disintegration over time.

Overall, pristine PBS and PBAT retained an almost intact macroscopic appearance during the early stages of the process. This observation applied both to bioaugmented and non-bioaugmented conditions (i. e. week 3). In contrast, both PBS- and PBAT-based biocomposites displayed clear signs of surface alteration. The PBS/OP biocomposite exhibited earlier and more pronounced visual alterations than PBAT/BSG, with marked discoloration and edge erosion already observable by week 3.

At the micrometre scale, cracks, fissures, and microcavities were observed on SEM images at the surface of all biocomposites as early as week 3. In contrast, the neat polymers exhibited no such features, or only very limited ones, throughout the sampling period.
<figure><img src="image_15.png" /><figcaption>**Fig. 13.** Photographs of sample fragments at two stages of the biodegradation process: week 3 (upper panels) and week 9 (lower panels), for the bioreactors located in chamber no. 2 (replicate no. 2). Missing fragments indicate an apparent total disintegration of the materials, as they were no longer visually discernible from the decomposition medium at the time of sampling. (A) PBS-based materials sampled at week 3: top left PBS, top right PBS+, bottom left PBS/OP, bottom right PBS/OP+. (B) PBAT-based materials sampled at week 3: top left PBAT, top right PBAT+, bottom left PBAT/BSG, bottom right PBAT/BSG+. (C) PBS-based materials sampled at week 9: top left PBS, top right PBS+, bottom left PBS/OP, bottom right PBS/OP+. (D) PBAT-based materials sampled at week 9: top left PBAT, top right PBAT+, bottom left PBAT/BSG, bottom right PBAT/BSG+.</figcaption></figure>

By week 9, both PBS/OP and PBS+ specimens were no longer visually distinguishable from the surrounding compost matrix. At this stage, the mineralization extent of PBS+ reached 87.2 % (replicate 2), indicating that the material had been nearly fully converted into CO₂. In contrast, PBS/OP reached only 58.1 % (replicate 2), implying that a substantial proportion of polymer-derived carbon remained in the degradation medium despite the absence of any visible residues.

This discrepancy suggests that, in the case of PBS/OP, a substantial proportion of the non-mineralized fraction likely persisted as micro- or nano plastic debris dispersed within the composting matrix. Conversely, the small residual fraction observed in PBS+ is more plausibly attributed to a much smaller proportion of nano-/microplastic particles, with the remaining difference corresponding to transient microbial biomass accumulation [48].

The markedly faster disintegration of PBS/OP can be explained by the hydrophilic nature of the orange-peel filler, whose strong water-sorption capacity was previously quantified by Dynamic Vapor Sorption (DVS) (Fig. 8) [29]. This high moisture uptake results from the transfer of water from the degradation medium (~70 % RH) into the filler phase and is expected to induce swelling of the filler particles during the first weeks of incubation. Such swelling likely generates internal mechanical stresses at the polymer-filler interface, thereby weakening interfacial adhesion. Consequently, the biocomposite becomes more susceptible to fracture under the physical constraints' characteristic of the composting environment (e.g. compaction and manual homogenization), in agreement with the disintegration

mechanisms described by Thia gamani et al. [102].

Importantly, fragmentation should not be conflated with biodegradation. While fragmentation refers to the mechanical or physicochemical breakdown of materials into smaller pieces, biodegradation involves the depolymerization and bioassimilation of polymer-derived carbon, followed by its conversion into CO₂, H₂O, and biomass under aerobic conditions [103]. Nevertheless, fragmentation substantially facilitates biodegradation by increasing both the external surface area available for microbial colonization and the internal reactive surface area through the formation of cracks and channels that enhance water diffusion and enzymatic penetration [104].

### 3.2.2. FTIR-ATR spectral characterization

FTIR-ATR spectra of the materials at the initial state, and after 3 and 9 weeks (W₃, W₉) of biodegradation, under the two inoculation conditions (i.e. compost extract and compost extract supplemented with bacterial enrichment), are presented in Figs. 14–17.

The infrared spectra of the materials at the initial state were characterized in detail in a previous study [29]. Briefly, PBS exhibits the typical bands of an aliphatic polyester, dominated by C=O stretching (1713 cm⁻¹), C=O stretching (1150–1260 cm⁻¹), and CH₂ absorptions (~2945, ~2860, 1455 cm⁻¹) [105]. PBAT exhibits characteristic aliphatic ester absorptions together with aromatic contributions, notably the C=C stretching band (1500–1600 cm⁻¹) and the aromatic C–H out-of-plane bending signals (~870 and ~720 cm⁻¹) [106]. For the biocomposites, no additional bands, peak shifts, or major intensity
<figure><img src="image_16.png" /><figcaption>PBS FTIR spectra over composting time</figcaption></figure>

**Fig. 14.** ATR-FTIR spectra normalized to the maximum intensity of the ester v(C=O) band, shown in the 1800–650 cm⁻¹ (right) and 4000–2750 cm⁻¹ (left) spectral regions, for pristine PBS and for both inoculation conditions (bioaugmented and non-bioaugmented) at the initial time (W₀), after three weeks of biodegradation (W₃), and after nine weeks (W₉).

<figure><img src="image_17.png" /><figcaption>PBSOP FTIR spectra over composting time</figcaption></figure>

**Fig. 15.** ATR-FTIR spectra normalized to the maximum intensity of the ester v(C=O) band, shown in the 1800–650 cm⁻¹ (right) and 4000–2750 cm⁻¹ (left) spectral regions, for the PBS/OP biocomposite under bioaugmented and non-bioaugmented conditions at the initial time (W₀), after three weeks of biodegradation (W₃), and after nine weeks (W₉).

<figure><img src="image_18.png" /><figcaption>PBAT FTIR spectra over composting time</figcaption></figure>

**Fig. 16.** ATR-FTIR spectra normalized to the maximum intensity of the ester v(C=O) band, shown in the 1800–650 cm⁻¹ (right) and 4000–2750 cm⁻¹ (left) spectral regions, for pristine PBAT under bioaugmented and non-bioaugmented conditions at the initial time (W₀), after three weeks of biodegradation (W₃), and after nine weeks (W₉).

changes were observed, indicating limited chemical interactions at the filler-matrix interface and predominantly physical adhesion.

Normalized FTIR spectra, scaled to the maximum peak intensity ($I_{norm} = I_i / I_{max}$), were used to semi-quantitatively assess the evolution of surface chemical composition throughout the biodegradation process. In all cases, the normalization reference ($I_{max}$) was the ester C=O stretching band (1713 cm⁻¹ for PBS-based materials and 1711.1 cm⁻¹ for PBAT-based materials), which consistently exhibits the highest absorbance across all spectra. This approach enables both qualitative and semi-quantitative assessment of spectral changes, including peak shifts,

the emergence or disappearance of bands, and pronounced variations in relative intensity [107].

During the biodegradation process, the most notable spectral change is the progressive development of a broad v(OH) absorption band (~3700–3000 cm⁻¹), characteristic of hydroxyl stretching vibrations. This feature is consistently observed across all materials (PBS, PBS/OP, PBAT, and PBAT/BSG) and under both inoculation conditions (i.e. compost extract alone and compost extract with bacterial enrichment). It reflects the generation of hydroxyl and carboxylic groups through ester bond hydrolysis, providing direct evidence of surface chemical
<figure><img src="image_19.png" /><figcaption>**Fig. 17.** ATR-FTIR spectra normalized to the maximum intensity of the ester v(C=O) band, shown in the 1800–650 cm⁻¹ (right) and 4000–2750 cm⁻¹ (left) spectral regions, for the PBAT/BSG biocomposite under bioaugmented and non-bioaugmented conditions at the initial time (W₀), after three weeks of biodegradation (W₃), and after nine weeks (W₉).</figcaption></figure>

transformations occurring during the biodegradation process [108,109].

For PBS-based materials, the relative OH signal showed a consistent increase across all formulations and inoculation conditions by week 3 (PBS: 0.094; PBS+: 0.105; PBS/OP: 0.112; PBS/OP+: 0.113). In contrast, by week 9 clear divergences emerged: PBS (0.142) and PBS+ (0.146) displayed only moderate increases relative to week 3, whereas PBS/OP (0.245) and PBS/OP+ (0.347) exhibited a marked enhancement of the OH band. These differences cannot be explained solely by the extent of mineralization at week 9, since the mineralization levels at that time were 31.9 % for PBS, 63.1 % for PBS+, 61.4 % for PBS/OP, and 63.4 % for PBS/OP+. This indicates a decoupling between the degree of mineralization at week 9 and the intensity of surface hydrolysis reflected by the OH band.

This marked increase indicates that the presence of orange peel filler substantially promotes hydrolysis-driven surface chemistry, particularly between the third and ninth week of biodegradation. Thus, while differences between systems are negligible at the early stage (W₃), the much stronger OH signal in PBSOP and PBSOP+ at W₉ clearly reflects a more advanced hydrolytic degradation compared to neat PBS.

For PBAT and PBAT+, the increase is similarly homogeneous and of comparable magnitude (PBAT: 0.056 → 0.083 from W₃ to W₉; PBAT+: 0.058 → 0.081), albeit less pronounced than in the PBS-based systems. In contrast, the brewer's spent grain biocomposite (PBAT/BSG) exhibit distinct kinetics: PBAT/BSG shows a strong but transient OH signal at W₃ (0.147), only moderate in PBAT/BSG+ (0.072), both decreasing by W₉ (PBAT/BSG: 0.082; PBAT/BSG+: 0.059). We hypothesize that this behaviour may reflect: (i) an initially rapid hydrolysis, facilitated by the hydrophilic filler, which enhances water uptake and generates interfacial zones where PBAT ester bonds are more readily attacked; and (ii) a subsequent leaching of soluble hydrolysis products (e.g., oligomers, organic acids) into the compost matrix, thereby attenuating the OH envelope at W₉ despite ongoing chemical transformations.

For all material-inoculum systems, a shoulder emerges in the v(C=O) band near ~1680 cm⁻¹, consistent with the formation of new carboxylic functions (–COOH/–COO⁻) generated through ester hydrolysis [110]. However, its temporal evolution differs across systems: in PBS and PBS+, the shoulder remains evident at week 9 relative to week 3 (and to the initial state), whereas in PBSOP and PBSOP⁺ it attenuates by week 9 compared with week 3 (and even with week 0 for PBSOP). For PBAT-based material-inoculum systems (PBAT and PBAT/BSG, with and without bacterial enrichment), this broadening at the base of the ester v(C=O) band is similar across conditions and remains of the same order between weeks 3 and 9. The only exception is PBATBSG at week 3, which instead shows a narrowing relative to its initial state.

Additional spectral changes further support hydrolysis-driven chemistry. First, a relative decrease is observed in the CH stretching domain (3000–2800 cm⁻¹) across all materials under both inoculation conditions, indicating reduced contributions from hydrophobic

methylene groups [111].

In addition, PBS-based systems display a progressive increase in the absorption between 1200 and 1100 cm⁻¹ in the C–O and C–O–C stretching region, consistent with the accumulation of hydroxyl and carboxylate functionalities [112].

Finally, in PBAT-based systems, terephthalate aromatic signatures become progressively more pronounced, as shown by the reinforcement of the out-of-plane aromatic C–H band (900–700 cm⁻¹). For PBAT, normalized intensity increased from 0.162 (W₀) to 0.320 (W₃) and 0.378 (W₉), with nearly identical evolution in PBAT+ (0.162 → 0.321 → 0.373). A comparable trend was observed in PBAT/BSG under compost extract (0.176 → 0.446 → 0.374), and under bacterial enrichment (PBATBSG+: 0.176 → 0.342 → 0.375). This monotonic increase reflects the preferential degradation of aliphatic domains, which are intrinsically more prone to hydrolytic attack than their aromatic counterparts. As degradation progresses, this selective susceptibility leads to a relative surface enrichment in butylene terephthalate segments [113].

Altogether, these spectral modifications indicate that hydrolysis is the dominant degradation pathway under mesophilic conditions. This pathway is likely predominantly enzymatic, with only a minor contribution from purely chemical reactions, due to the relatively low incubation temperature (28 °C) [16]. In general, the broad OH band (≈3000–3700 cm⁻¹) is more intense at week 9 than at week 3, reflecting a more advanced stage of hydrolytic degradation consistent with the temporal progression of the process.

### 3.2.3. Crystallinity

The crystallinity degree was determined for all materials sampled across the 24 bioreactors, with two analytical replicates per bioreactor, at weeks 3 and 9 under both conditions (i.e. mature compost extract alone, or mature compost extract combined with bacterial enrichment). Crystallinity was calculated from the melting and cold crystallization enthalpies measured during the first heating cycle, to reflect the structural state of the polymers now of sampling [114]. The results are presented in Fig. 18.

Only marginal, non-significant variations in crystallinity were observed between the initial state, week 3, and week 9 after the onset of the biodegradation process. This trend suggests that the degradation of amorphous domains does not substantially precede that of crystalline regions but rather occurs in parallel. In other words, both phases appear to be progressively affected at a comparable rate, leading to a relatively stable overall crystallinity throughout the process. This behaviour contrasts with the common assumption that amorphous regions are preferentially hydrolysed due to their higher accessibility to water [16]. Instead, it is consistent with a surface-mediated, enzyme-driven hydrolytic process, in which the cleavage of macromolecular chains occurs in both amorphous and crystalline domains, rather than with a purely chemical hydrolysis mechanism, as water alone cannot readily penetrate
<figure><img src="image_20.png" /><figcaption>**Fig. 18.** Evolution of the crystallinity extent (first heating scan) of the different material-inoculum combinations during the biodegradation process, from the initial time (W₀) to week 3 (W₃) and week 9 (W₉).</figcaption></figure>

crystalline structures.

## 3.3. Microbial monitoring during biodegradation

### 3.3.1. Evolution of CFU·g⁻¹ concentration during biodegradation

The mean concentrations (CFU·g⁻¹ of degradation medium) of culturable bacteria enumerated on R2A nutrient agar, for all experimental conditions (i.e. material-inoculum combinations) at weeks 3 and 9 after the start of the experiment, are reported in Table 9.

The mean CFU·g⁻¹ concentration was substantially higher at week 3 compared with week 9 across all conditions. At week 3, values ranged from 2.78 × 10⁷ CFU·g⁻¹ for PBAT to 5.64 × 10⁸ CFU·g⁻¹ for PBS+. Overall, the CFU·g⁻¹ concentration of the degradation medium was positively correlated with the mineralization extent at the time of sampling in week 3. This correlation was moderate and statistically significant at the 5 % alpha level (r = 0.47, R² = 0.22, p = 0.02, n = 24). At week 9, the positive linear relationship between CFU·g⁻¹ concentration and the mineralization extents was weaker and no longer statistically significant at the 5 % alpha level (R² = 0.12, r = 0.34, p = 0.11, n = 24) (Fig. 19).

The positive linear relationship between CFU·g⁻¹ in the degradation medium and the mineralization extent across all materials and inocula reflects the central role of microbial abundance in driving the biodegradation of biodegradable polymers. Higher CFU·g⁻¹ values indicate more active microbial communities, which in turn translate into greater CO₂ release and higher mineralization rates.

The progressive decline in CFU·g⁻¹ concentration from week 3 to week 9, along with the weakening statistical association between microbial abundance and mineralization extent, likely reflects the temporal dynamics of the biodegradation process. At week 3, the

<caption>**Table 9**</caption>

Culturable bacterial abundance at Weeks 3 and 9 for each material-inoculum combination.

<table><thead><tr><th>Materials/Inocula</th><th>Week 3 (CFU·g<sup>-1</sup>)</th><th>Week 9 (CFU·g<sup>-1</sup>)</th><th>W<sub>9</sub>/W<sub>3</sub> (%)</th></tr></thead><tbody><tr><td><strong>PBAT-based</strong></td><td>2.78·10<sup>7</sup> ± 0.21</td><td>7.53·10<sup>6</sup> ± 5.50</td><td>27.1</td></tr><tr><td>PBAT</td><td></td><td></td><td></td></tr><tr><td>PBAT+</td><td>1.61·10<sup>8</sup> ± 0.53</td><td>1.13·10<sup>7</sup> ± 0.34</td><td>7.0</td></tr><tr><td>PBAT/BSG</td><td>2.00·10<sup>8</sup> ± 1.20</td><td>1.64·10<sup>7</sup> ± 0.19</td><td>8.2</td></tr><tr><td>PBAT/BSG+</td><td>1.49·10<sup>8</sup> ± 0.11</td><td>5.47·10<sup>6</sup> ± 0.67</td><td>3.7</td></tr><tr><td><strong>PBS-based</strong></td><td>1.21·10<sup>8</sup> ± 0.85</td><td>1.08·10<sup>7</sup> ± 0.25</td><td>8.9</td></tr><tr><td>PBS</td><td></td><td></td><td></td></tr><tr><td>PBS+</td><td>5.64·10<sup>8</sup> ± 4.5</td><td>1.46·10<sup>7</sup> ± 0.80</td><td>2.6</td></tr><tr><td>PBS/OP</td><td>9.45·10<sup>7</sup> ± 3.5</td><td>1.31·10<sup>7</sup> ± 0.40</td><td>13.8</td></tr><tr><td>PBS/OP+</td><td>2.03·10<sup>8</sup> ± 1.4</td><td>1.18·10<sup>7</sup> ± 0.32</td><td>5.8</td></tr></tbody></table>

**Notes.** Counts were obtained by serial dilution plating on R2A nutrient agar after 72 h at 24 ± 1 °C in the dark. W₉/W₃ indicates the percentage of CFU remaining at Week 9 relative to Week 3.

composting environment was still in an active phase, during which the mineralization curves displayed linear or near-exponential increases. At this stage, bacterial communities proliferated rapidly without apparent constraints related to substrate accessibility or the accumulation of inhibitory metabolites, resulting in high microbial densities (CFU·g⁻¹).

In contrast, by week 9 the mineralisation kinetics had entered a decelerating or plateau phase (Figs. 5 and 6), characterized by reduced microbial turnover and lower bioavailability of polymer-derived carbon, particularly due to the depletion of easily hydrolysable fractions.

### 3.3.2. Physiological profiling of bacterial communities (CLPP)

Figs. 20–25 present the main results of the functional metabolic profiling (CLPP) of microbial communities sampled at weeks 3 and 9.

This approach was based on Biolog EcoPlates™, which contain 96 wells organized into triplicates of 31 different carbon substrates plus a water control. The metabolic response of the microbial communities was assessed by measuring colour development at 590 nm (i.e. conversion of tetrazolium into violet formazan) in the wells as an indicator of substrate utilization [115].

Such profiling is particularly relevant in the context of polymer biodegradation, as it provides insights into potential shifts in the functional diversity and metabolic versatility of microbial consortia exposed to different substrates during composting [116,117]. In our system, this approach is particularly informative, as the tested materials (i.e. PBS, PBS/OP, PBAT, and PBAT/BSG) constituted the only available and utilizable carbon sources in the vermiculite medium. Consequently, the CLPP responses observed here directly reflect the adaptation of microbial communities to these substrates, without interference from alternative carbon inputs such as those typically present in soils, mature compost, or mixed biowaste environments (e.g. humic substances in soils, lignocellulosic residues in mature compost, or easily degradable sugars, proteins, and lipids in mixed biowaste environments) [46].

#### 3.3.2.1. Functional diversity.
The comparative analysis of functional diversity indices provides a quantitative assessment of the metabolic potential and niche breadth of the microbial communities exposed to the different polymer-based materials and biocomposites across the various inoculum combinations. Shannon's index reflects both the richness of metabolised carbon sources and the contribution of rare functions, capturing the overall breadth of community activity. In contrast, Simpson's index gives greater weight to dominant substrates and is less sensitive to rare ones, thus emphasizing patterns of dominance versus evenness [118]. In parallel, functional richness (number of substrates with OD > 0.25) provides a direct measure of the breadth of significantly utilized substrates [57]. Together, these indices offer complementary perspectives on community functional diversity (Fig. 20).

At week 3, one-way ANOVA demonstrated significant overall treatment effects for each of the three diversity indices. These indices included Shannon (F = 4.98, p < 0.01), Simpson (F = 6.28, p < 0.01), and Richness (F = 5.74, p < 0.01).

Overall, pristine PBS inoculated solely with the mature compost extract exhibited the highest Shannon diversity (3.29 ± 0.10) and functional richness (29.7 ± 0.6) of all treatments. Both indices differ significantly from the bioaugmented PBS+ condition, the first at the 5 % level and the second at the 10 % level. Accordingly, PBS+ displayed lower functional diversity (Shannon = 3.14 ± 0.07; Richness = 26.7 ± 1.2) than PBS without bioaugmentation, suggesting that microbial communities originating exclusively from the mature compost extract mobilized a broader metabolic repertoire.

In contrast, PBAT+ displayed the lowest functional diversity (Shannon = 3.03 ± 0.01; Richness = 20.67 ± 0.57) among all treatments, with values for Simpson and Richness significantly lower (p < 0.05) than those of PBAT/BSG, PBAT/BSG+, PBS+, and PBS/OP+. PBAT without bioaugmentation exhibited similar diversity values (Shannon = 3.14 ± 0.02; Richness = 24.67 ± 0.57), with no statistically
Linear regression between microbial abundance and mineralization

<figure><img src="image_21.png" /><figcaption>**Fig. 19.** Relationship between the logarithm of viable cell counts [log₁₀ (CFU g⁻¹) of fresh decomposition medium] and the corresponding mineralization extent for the 24 bioreactors, determined on R2A agar at week 3 (left) and week 9 (right). Linear regression lines are shown together with the corresponding regression equations and coefficients of determination (R²).</figcaption></figure>

<figure><img src="image_22.png" /><figcaption>**Fig. 20.** Functional diversity indices (Shannon, Simpson (1-D), and functional richness, defined as the number of substrates with OD > 0.25) for the different material-inoculum combinations at week 3 and week 9.</figcaption></figure>

significant differences between the two inoculation conditions.

By week 9, all indices had declined significantly relative to week 3, consistent with a progressive functional specialization of microbial communities during biodegradation. Nevertheless, differences between the two inoculation strategies (bioaugmented vs. non-bioaugmented) persisted across treatments, as indicated by the one-way ANOVA test, which revealed significant effects for Shannon (F = 6.06, p < 0.01), Simpson (F = 5.91, p < 0.01), and Richness (F = 4.34, p < 0.01).

Across all conditions, Shannon diversity at W₉ ranged from 1.41 to 2.10, whereas at W₃ it had ranged from 3.03 to 3.30. In general, richness also declined, although to a lesser extent. For instance, in PBS it decreased from 29.7 ± 0.6 at W₃ to 25.0 ± 1.7 at W₉.

For PBS, Shannon diversity remained higher in the non-bioaugmented condition (1.69 ± 0.10) than in PBS+ (1.55 ± 0.06; p = 0.024), while richness declined in both but was better preserved without bioaugmentation (PBS: 25.0 ± 1.7; PBS+: 23.0 ± 1.0).

Regarding the biocomposites, PBAT/BSG maintained higher richness in the non-bioaugmented system (26.0 ± 0.0 vs. 23.7 ± 0.6; p < 0.05). In contrast, PBS/OP exhibited statistically indistinguishable outcomes between the two conditions (Shannon ≈ 1.6–1.7; Richness ≈ 24–25; p > 0.05). This indicates that bioaugmentation did not substantially modify the CLPP-based functional profile of the microbial communities associated with PBS/OP, in contrast to PBAT/BSG, which exhibited clear functional specialisation following enrichment. However, in both cases, these shifts did not translate into significant differences in the mineralization kinetics between the bioaugmented and non-bioaugmented

systems.

Taken together, these results indicate that bioaugmentation actively restructured the functional communities, leading to lower richness and Shannon diversity, particularly in PBS- and PBAT/BSG-based systems. This trend suggests that the enriched strains introduced in the bio-augmented treatments promoted a more specialized metabolic activity, preferentially targeting selected substrates rather than sustaining a broad functional repertoire. In contrast, microbial communities originating solely from the mature compost extract (non-bioaugmented condition) displayed greater metabolic versatility, particularly in PBS systems.

In the case of PBS+, this reduction in functional richness coincided with an increase in polymer mineralization (Fig. 5). In contrast, for PBAT/BSG, the final mineralization extent remained unchanged between the bioaugmented and non-bioaugmented conditions.

Correlation analysis (Fig. 21) revealed that at week 3, the three indices were strongly associated (Simpson vs Shannon: R² = 0.98, p < 0.001; Shannon vs. richness: R² = 0.94, p < 0.001; Simpson vs. richness: R² = 0.94, p < 0.001), indicating that functional diversity was mainly driven by a broad and homogeneous substrate utilization profile. By week 9, Simpson and Shannon remained highly correlated (R² = 0.99, p < 0.001), but their correlations with richness declined (Shannon vs. richness: R² = 0.72, p < 0.001; Simpson vs. richness: R² = 0.65, p < 0.001).

Beyond correlation strength, regression slopes also changed markedly over time. Specifically, the slope of the Shannon–richness
Linear regressions between diversity indices (Week 3 and Week 9)

<figure><img src="image_23.png" /><figcaption>Fig. 21. Linear relationships between the three functional diversity indices (Shannon, Simpson (1-D), and functional richness) at weeks 3 and 9, with the corresponding regression equations and coefficients of determination (R²).</figcaption></figure>

PCA - Variable Contribution Analysis (Week 3)

<figure><img src="image_24.png" /><figcaption>Fig. 22. Contributions of the six variables to the first two principal components (PC1 and PC2) at week 3. The horizontal red dashed line indicates the significance threshold calculated as (1/N) × 100, where N is the total number of variables; here, (1/6) × 100 = 16.66 %.</figcaption></figure>

relationship increased from 0.030 at week 3–0.069 at week 9, while that of Simpson versus richness increased from 0.0015 to 0.019. This indicates that, at later stages of biodegradation, increases in the number of metabolized substrates were associated with proportionally larger gains in both Shannon and Simpson diversity. In other words, substrate richness contributed more strongly to overall functional diversity at week 9 than at week 3.

Combined with the persistent positive correlations observed at both time points, these results suggest a partial re-coupling between substrate range and evenness during the later stages of composting. Although overall diversity levels declined over time, the substrates that remained metabolized at week 9 tended to be utilized in a more balanced manner across the community. This pattern is consistent with the emergence of a more specialized yet functionally coherent microbial consortium, in

which fewer substrates support a relatively even distribution of metabolic activity.

3.3.2.2. AWCD (Average Well Colour Development). The AWCD (Average Well Colour Development) represents the overall metabolic activity potential of microbial communities across all carbon substrates tested in EcoPlates™ [119]. It provides an integrative measure of the overall metabolic potential of microbial communities [120]. A high AWCD reflects intense catabolic enzymatic activity and expressed functional diversity, whereas a low AWCD may indicate a weakly active community constrained by unfavourable environmental conditions, or one specialized in a narrow spectrum of substrates [121].

At week 3, PBS exhibited a significantly higher value (1.58 ± 0.16) compared to the other material-inoculum combinations. At week 9, the
PCA - Variable Contribution Analysis (Week 9)

<figure><img src="image_25.png" /><figcaption>Fig. 23. Contributions of the six variables to the first two principal components (PC1 and PC2) at week 9. The horizontal red dashed line indicates the significance threshold calculated as (1/N) × 100, where N is the total number of variables; in this case, (1/6) × 100 = 16.66 %.</figcaption></figure>

PCA Summary - Week 3

<figure><img src="image_26.png" /><figcaption>Fig. 24. Principal component analysis (PCA) of the 31 carbon substrates grouped into six main biochemical families at week 3 of the biodegradation process. Left: Correlation circle of the six variables. Right: score plot of the material-inoculum combinations. Ellipses represent 90 % confidence intervals around the group centroids.</figcaption></figure>

PCA Summary - Week 9

<figure><img src="image_27.png" /><figcaption>Fig. 25. Principal component analysis (PCA) of the 31 carbon substrates grouped into six main biochemical families at week 9 of the biodegradation process. Left: Correlation circle of the six variables. Right: score plot of the material-inoculum combinations. Ellipses represent 90 % confidence intervals around the group centroids.</figcaption></figure>

highest values were again observed for PBS (1.17 ± 0.10), but also for PBAT/BSG (1.17 ± 0.13). Conversely, the lowest values were
consistently associated with PBAT+ (W₃: 0.94 ± 0.05; W₉: 0.87 ± 0.03).

In general, like the number of carbon sources utilized (richness index), AWCD tends to decrease between weeks 3 and 9. This decrease is significant at the 5 % level for PBAT, PBAT/BSG+, PBS, and PBS/OP+. Such a trend suggests an overall decline in catabolic activity and/or expressed functional diversity within microbial communities, likely related to the progressive reduction in the availability of readily assimilable substrates and/or to an increasing specialization of the communities throughout the biodegradation process.

### 3.3.2.3. Principal component analysis (PCA).
Principal component analyses (PCA) were performed based on the complete set of optical density values obtained for the 31 utilizable carbon sources of the Biolog EcoPlates®, which were grouped into six major biochemical families: (A) amines, (AA) amino acids, (CA) carboxylic acids, (CCS) complex carbon sources, (PCS) phosphorylated carbon sources, and (CH) carbohydrates. The detailed list of carbon substrates belonging to each biochemical family is provided in Table 6.

#### 3.3.2.3.1. Structuring of PCA Axes.
The structuring of the first two dimensions of the principal component analyses at weeks 3 and 9, highlighting the contribution of the six biochemical families to the construction of the axes, is shown in Figs. 22 and 23.

At week 3, the first two principal components accounted for 93.2 % of the total variance (PC1 = 83.9 %; PC2 = 9.3 %). PC1 was mainly structured by the utilization of carboxylic acids (18.9 %), amino acids (18.9 %), carbohydrates (16.6 %), and complex carbon sources (17.4 %) (Fig. 22), reflecting a gradient in the exploitation of primary and structurally diverse energy substrates.

A size effect was observed on this axis, as all families were positively correlated in the same direction, indicating that PC1 essentially captured the overall metabolic intensity. Accordingly, inoculum-material combinations positioned on the right-hand side of the individuals plot correspond to communities with a generally higher capacity to metabolize the six biochemical families of carbon substrates, whereas those located on the left-hand side represent lower global metabolic activity.

At week 9, a size effect was again apparent on PC1 (65.6 % of the variance), with all six substrate families remaining positively correlated. Compared with week 3, the relative contributions shifted moderately, with amino acids (23.8 %), carboxylic acids (22.4 %), carbohydrates (22.0 %), and amines (20.9 %) emerging as the most influential families. This pattern indicates a rebalancing of the relative contribution of substrate families rather than a fundamental alteration of the metabolic profile, since three of the four major biochemical groups continued to dominate the overall variation along PC1.

At week 3, the second axis (PC2) was largely structured by amines (65.8 %) (Fig. 22), underscoring the role of nitrogen-rich compounds in driving community differentiation. Taken together, the two axes thus represent a continuum ranging from high-energy-yielding carbon substrates (i.e. HC, AC, SCC, AA) on PC1 to nitrogen-containing compounds (A) on PC2, thereby capturing the two principal forces structuring functional diversity at this stage. Indeed, although both amino acids and amines are nitrogen-containing substrates, their metabolic roles differ. Amino acids fuel central carbon metabolism by providing both energy and anabolic precursors [122], whereas amines primarily reflect nitrogen recycling through ammonium release, with limited contribution to energy metabolism [123]. This distinction likely explains why amines structured PC2 independently of amino acids.

By week 9, the second axis (PC2 = 19.0 %) was dominated by phosphorylated carbon substrates (PCS, 74.0 %), followed by complex carbon sources (CCS, 15.9 %) (Fig. 23). This shift indicates that, at this later stage of biodegradation, secondary differentiation between inoculum-material combinations was primarily driven by phosphorylated substrates, in contrast to the amine-based structuring observed at week

3.

#### 3.3.2.4. Correlation Circle and Individuals Factor Map.
The correlation circles of the variables and the scatterplots of the individuals (i.e. material-inoculum pairs) are presented in Figs. 24 and 25 for weeks 3 and 9, respectively.

At week 3, PBS was the most positively correlated with PC1, which is consistent with the previous AWCD results indicating its generally higher metabolic activity across the 31 carbon substrate sources. It exhibited markedly higher values for two substrate families compared to PBS+ (HC: 0.42 ± 0.05 vs. 0.31 ± 0.04; AC: 0.38 ± 0.06 vs. 0.27 ± 0.05), suggesting a broader metabolic mobilization of readily degradable carbon sources in the absence of bioaugmentation.

However, PBS+ displayed higher absolute values for amino acids compared to PBS (AA: 0.35 ± 0.04 vs. 0.28 ± 0.03). Unlike carbohydrates or other carbon sources, which mainly provide readily available energy without necessarily sustaining prolonged microbial activity, amino acids supply both carbon and assimilable nitrogen [122]. Their mobilization therefore supports microbial growth and enzyme production. This specialization is consistent with the higher mineralization extent observed for PBS+, indicating that bioaugmentation fostered more metabolically active microbial communities.

This trend was further reflected in the ordination space, where PBS+ communities were positioned closer to the centre and displayed a marked orientation along axis 2. As this axis was strongly structured by the Amines family, the normalized mean optical density for this family remained slightly higher in PBS than in PBS+ (A: 1.57 ± 0.19 vs. 1.46 ± 0.26). Hence, the partitioning and divergence of PBS+ from PBS and from the other modalities (i.e. the different inoculum-material communities) were primarily driven by the predominance of the relative weight of amines in the total AWCD (PBS+: 9.16 %; PBS: 6.42 %), rather than by higher absolute values.

These results point to an early metabolic imbalance between carbon- and nitrogen-based pathways. Whereas PBS communities relied more heavily on readily available carbon sources, PBS+ communities preferentially mobilized amino acids and amines, thereby coupling carbon assimilation with a direct supply of assimilable nitrogen. This shift towards C- and N-rich substrates likely promoted microbial growth and enzyme production, which in turn sustained the higher mineralization rate observed during this initial stage of biodegradation [124].

These results were not observed for the PBS/OP+ biocomposite, despite being enriched with the same specific bacterial inoculum as the PBS+ treatment. This indicates that the incorporation of orange peel powder profoundly altered the trajectory of the microbial community's functional structuring associated with this biocomposite during the early stages of biodegradation.

This phenomenon is most likely explained by an early selective pressure exerted by the presence of low-complexity carbon substrates (e. g. such as simple sugars, or proteins) released from the filler [94]. Indeed, these readily assimilable carbon compounds promote the rapid growth of opportunistic or generalist microorganisms already present in the compost extract used as the basal inoculum. Consequently, by week 3, the microbial community observed in the PBS/OP+ treatment appeared structurally and functionally more like that of the non-enriched PBS/OP, with fewer divergences than those observed between PBS and PBS+. This suggests that the effect of bacterial enrichment was less decisive than the selective pressure arising from the organic filler, which rapidly reshaped the community trajectory.

In other words, microorganisms from the indigenous compost-extract community, likely better adapted to the low-complexity molecular substrates released by the OP filler, rapidly occupied the ecological niche, thereby limiting the establishment and/or persistence of the strains introduced through enrichment [125,126]. This illustrates a typical case of native community resilience to an enrichment attempt and highlights that the success of such a strategy primarily depends on
the initial ecological conditions (e.g. availability of carbon sources, community structure) [127]. These findings underscore the importance of the three-dimensional interactions between the polymer matrix, the organic filler, and the microbial community in shaping the dynamics of ecological niche colonization and subsequent biodegradation of the target substrate.

At week 9, the ordination plot showed no clear clustering, with individuals dispersed across both axes. This contrasted with week 3, where communities were distinctly structured along PC1 and PC2.

### 3.4. Bioaugmentation under field-relevant conditions

In this study, bioaugmentation was used as an exploratory strategy to identify biological limitations and potential improvement pathways for polyester biodegradation under mesophilic, decentralized composting conditions. It was not intended as a direct, user-driven practice at the household level. Further investigations are therefore required to assess the robustness and transferability of these findings to more complex commercial formulations and more heterogeneous composting matrices.

Although bioaugmentation appears promising for enhancing the biodegradability of certain polyesters under mesophilic conditions, its practical implementation is more realistically envisioned through the development of dedicated compost activators managed at the community or institutional scale, rather than through individual household practices.

At this scale, inoculants could be formulated using established preservation routes (e.g., freeze-drying and carrier-based formulations), enabling shelf-life and transportability, as already practiced for agricultural and environmental microbial products. From a translational perspective, the economic feasibility of such interventions is expected to be strongly scale-dependent and more realistic when composting operations are collectively managed, enabling standardized practices and economies of scale.

## 4. Conclusion

This work demonstrates that bioaugmentation with a consortium composed of *Pseudomonas putida*, *Stenotrophomonas maltophilia*, *Achromobacter denitrificans*/*xylosoxidans*, and *Rhizobium radiobacter*, previously isolated through a selective enrichment procedure in minimal medium, can substantially enhance the mesophilic (28 °C) compostability of pristine PBS. Under these conditions, bioaugmentation yielded a + 68.3 % increase in final mineralization after 133 days compare to extract with compost mature, raising the extent to 73.2 % and thereby bringing the material closer to the 90 % threshold defined in home-compostability standards. However, this effect was not observed for the PBS/OP biocomposite, which already exhibited an + 83.2 % increase in final mineralization compared to pristine PBS under non-bioaugmented conditions. In contrast, PBAT remains a recalcitrant aliphatic-aromatic copolyester under mesophilic conditions, showing very limited mineralization and no significant improvement through bioaugmentation, whether tested alone or as biocomposite (i.e. PBAT/BSG).

Overall, this study highlights that the effective biodegradation of certain biodegradable polyesters, such as PBS, under decentralized mesophilic composting conditions may require specific biological strategies beyond those achieved with conventional compost alone. These strategies may include the use of dedicated compost activators containing selected microbial strains. Such approaches appear more realistically applicable within community-scale composting systems managed by local authorities or private operators, potentially under appropriate regulatory incentives, rather than as user-driven practices at the individual household level.

### CRediT authorship contribution statement

**Feriel Bacoup:** Validation, Supervision, Project administration, Funding acquisition. **Jules Bellon:** Writing – review & editing, Writing – original draft, Visualization, Validation, Software, Resources, Project administration, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. **Richard Gattin:** Validation, Supervision, Project administration, Funding acquisition.

### Funding

This work was supported by the French region of Normandy [RIN50: 22 E01400 - 00123402]; and the Unilasalle Polytechnic Institute [Jean-Baptiste Gagne Foundation].

### Declaration of Competing Interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper

### Acknowledgements

The authors would like to thank and acknowledge the French Region of Normandy and the UniLaSalle Polytechnic Institute for their financial support of this work. The Agrobiotech research platform, labelled GIS IBISA (French Infrastructure in Biology, Health and Environment), is also gratefully acknowledged. Grace Ifembo Sikumbili, Rebecca Djamba, Jean-Baptiste Besnier, Samantha Vespier-Mori, Lisa Castel, and Anne Cauchois are warmly thanked for their invaluable assistance with the experimental work.

### Data availability

Data will be made available on request.

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