# Assessing the Biodegradation of Low-Density Polyethylene Films by Candida tropicalis SLNEA04 and Rhodotorula mucilaginosa SLNEA05

Randa Harrat, Ghania Bourzama, Gaëtan Burgaud, Emmanuel Coton, Aymen Bourezgui, Boudjema Soumati

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Randa Harrat, Ghania Bourzama, Gaëtan Burgaud, Emmanuel Coton, Aymen Bourezgui, et al. "Assessing the Biodegradation of Low-Density Polyethylene Films by Candida tropicalis SLNEA04 and Rhodotorula mucilaginosa SLNEA05." Diversity, 2024, 16 (12), pp.759. 10.3390/d16120759 hal-05140927

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Article

# Assessing the Biodegradation of Low-Density Polyethylene Films by *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05

Randa Harrat <fnref n="1" />,* , Ghania Bourzama <fnref n="2" />, Gaëtan Burgaud <fnref n="3" />,<fnref n="4" />,* , Emmanuel Coton <fnref n="3" /> , Aymen Bourezgui <fnref n="5" /> and Boudjema Soumati <fnref n="1" />

**Abstract:** Environmental pollution resulting from the accumulation of plastic waste poses a major ecological challenge. Biodegradation of these polymers relies on microorganisms capable of decomposing them, generally through the biodeterioration, biofragmentation, assimilation, and mineralization stages. This study evaluates the contribution and efficacy of indigenous soil yeasts isolated from a northeastern Algerian landfill in degrading low-density polyethylene (LDPE) plastic bag films. *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05 were identified through internal transcribed spacer (ITS) and large subunit ribosomal RNA gene sequencing. These isolates were then tested for their ability to biodegrade LDPE films and utilized as the sole carbon source *in vitro* in a mineral salt medium (MSM). The biodegradation effect was examined using scanning electron microscopy (SEM), attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy, and X-ray diffraction (XRD). After 30 days of incubation at 25 °C, a significant weight loss was observed compared to the control for both cultures: 7.60% and 5.53% for *C. tropicalis* and *R. mucilaginosa*, respectively. SEM analysis revealed morphological alterations, including cracks and holes, ATR-FTIR detected new functional groups (alcohols, alkynes, aldehydes, alkenes and ketones), while XRD identified changes in the polymer crystallinity and phase composition. These findings underscore the potential of the two yeast isolates in LDPE biodegradation, offering promising insights for future environmental applications.

**Keywords:** plastic pollution; LDPE; bioremediation; yeast; SEM; ATR-FTIR; XRD

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**Citation:** Harrat, R.; Bourzama, G.; Burgaud, G.; Coton, E.; Bourezgui, A.; Soumati, B. Assessing the Biodegradation of Low-Density Polyethylene Films by *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05. *Diversity* **2024**, *16*, 759. https://doi.org/10.3390/d16120759

Academic Editor: Ipek Kurtboke

Received: 3 September 2024

Revised: 24 October 2024

Accepted: 9 December 2024

Published: 12 December 2024

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**Copyright:** © 2024 by the authors.
Licensee MDPI, Basel, Switzerland.
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4.0/).

## 1. Introduction

Plastic pollution poses a significant environmental challenge [1,2], with an estimated 9.2 billion tons of plastics produced since the 1950s, of which 6.9 billion tons have become waste [3]. Global plastic production reaches nearly 400 million tons annually [4], primarily consisting of single-use plastics with lifespans from minutes to hours yet persisting in the environment for decades to centuries [3]. Polyethylene (PE) alone constitutes about 64% of the accumulated plastic waste and is regarded as one of the most problematic polymers [5].

In Algeria, around 2.1 million tons of plastic waste are generated annually, accounting for 15.31% of household waste. Low-density polyethylene (LDPE), primarily from plastic bags, constitutes a significant part of this waste, representing nearly 8.14% of the total.

<footnote n="1">¹ Laboratory of Biochemistry and Environmental Toxicology, Badji Mokhtar-Annaba University, P.O. Box 12, Annaba 23000, Algeria; soumati_boujema@yahoo.fr</footnote>

<footnote n="2">² Laboratory of Microbiology and Molecular Biology, Badji Mokhtar-Annaba University, P.O. Box 12, Annaba 23000, Algeria; gbourzama@yahoo.fr</footnote>

<footnote n="3">³ Univ Brest, INRAE, Laboratoire Universitaire de Biodiversité et Écologie Microbienne, 29280 Plouzané, France; emmanuel.coton@univ-brest.fr</footnote>

<footnote n="4">⁴ Institut Universitaire de France, 75005 Paris, France</footnote>

<footnote n="5">⁵ National School of Advanced Sciences and Technologies of Borj Cédria (ENSTAB), University Campus of the Borj Cédria Technopole, P.O. Box 122, Hammam Chott 1164, Tunisia; bourezgui.aymen@gmail.com</footnote>

<footnote>* Correspondence: randa.harrat@univ-annaba.dz or harratranda@gmail.com (R.H.); gaetan.burgaud@univ-brest.fr (G.B.)</footnote>
Algerian households produce around 23 billion plastic bags annually, averaging 557 bags per person [6].

Traditional disposal methods, including landfilling, incineration and recycling, appear insufficient to address the growing plastic waste problem, prompting research into alternative degradation methods [5]. Biodegradation, particularly through microbial activity, may offer a promising solution [3]. Fungi have emerged as effective agents in polymer degradation due to their role in organic matter decomposition, including refractory organic matter, and metabolic versatility [7]. Fungal enzymes such as laccases, peroxidases, cutinases, and lipases catalyze the breakdown of polymers into smaller monomers, facilitating microbial uptake and subsequent mineralization [1].

Several filamentous fungal genera have demonstrated the ability to degrade polyethylene, such as *Alternaria* [8], *Aspergillus*, *Penicillium*, *Fusarium* [9–11], *Cladosporium* [12], *Gliocladium*, *Mortierella* [13], *Mucor* [14], and *Zalerion* [15]. These fungi are particularly effective at degrading low-density polyethylene (LDPE) due to their ability to release hydrophobic proteins that enhance colonization and interaction with the polymer. This is followed by the secretion of enzymes that enable them to utilize the polymer as a carbon source [16].

Yeasts, a significant group within the fungal kingdom, have been extensively utilized in various biotechnological and industrial applications due to their diverse metabolic capacities [17]. Although their potential in biodegradation studies has attracted attention, these investigations remain relatively limited compared to those involving other filamentous fungal species. Notably, *Saccharomyces* has demonstrated impressive degradation of LDPE sheets, achieving a 43% weight loss after 30 days of incubation at room temperature [18]. This value can be compared to filamentous fungal species, such as *Aspergillus clavatus* JASK1, which degraded LDPE strips (0.5 g, 2 × 2 cm) by 35% over 90 days of incubation in an enrichment medium, under shaking conditions at 120 rpm [19]. Other species, including *A. niger*, *A. flavus*, *Rhizopus oryzae*, and *Thermomyces lanuginosus*, have also exhibited varying levels of PE degradation. After 30 days of incubation, these fungi, respectively, achieved weight losses of 7.65 ± 0.92% for PE microplastics, 11% for LDPE sheets, 8.4 ± 3% for LDPE film, and 9.21 ± 0.84% for pretreated LDPE film, with the pretreatment involving the synergistic effects of UV irradiation, heat, and acid [18,20–22].

Fungi capable of degrading plastics continue to be isolated from diverse environments, including Algeria, where research on fungal plastic degradation remains limited. In this context, this study aims to address this gap by evaluating the ability of two indigenous soil yeasts, isolated from a northeastern Algerian landfill, to degrade low-density polyethylene (LDPE) plastic films. By exploring the potential of these local microbial resources, this research contributes to the development of sustainable, eco-friendly strategies for mitigating plastic pollution.

**2. Materials and Methods**

**2.1. Sample Collection**

A soil sample was collected in April 2021 from the Boumehadjer public landfill (36°59′35″ N, 6°33′23″ E), located at the entrance of the city of Collo in northeastern Algeria (Figure 1). The sample was taken after removing the top five centimeters of the soil surface and was transported to the laboratory in sterile conditions, at 4 °C, and processed within 24 h of collection.
<figure><img src="image_5.png" /><figcaption>**Figure 1.** (a) Geographic location of the Boumehadjer landfill. (b) Photograph of waste at the Boumehadjer landfill.</figcaption></figure>

## 2.2. Isolation of Indigenous Fungi from Soil

Ten grams of soil, collected from a site with long-term plastic waste exposure, were added to 90 mL of sterile physiological saline solution in a sterile Erlenmeyer flask. This environment served as a natural enrichment, likely favoring the selection of microorganisms adapted to plastic degradation. The mixture was stirred for 30 min, followed by one hour of setting. The supernatant served as the stock solution for subsequent analyses. Decimal dilutions were prepared, and 0.1 mL of each dilution was inoculated onto potato dextrose agar (PDA) plates supplemented with 20 µg/mL of gentamicin to inhibit bacterial growth. The plates were incubated at 27 °C for one week [23]. Fungal isolates were then purified through successive subcultures on PDA, without the need for additional enrichment steps.

## 2.3. Molecular Characterization and Identification of Yeasts

Yeast isolates were identified based on the sequences of the internal transcribed spacer (ITS) and the D1/D2 regions of the large subunit ribosomal RNA (26S) using the following primers: ITS1 5'-CTTGGTCATTTAGAGGAAGTAA-3'/ITS4 5'-TCCTCCGCTTATTGATATGC-3' [24] and NL1 5'-GCATATCAATAAGCGGAGGAAAAG-3'/NL4 5'-GGTCCGTGTTTCAAGACGG-3' [25]. Genomic DNA was extracted using a FastDNA Spin Kit (MP Biomedicals), following the manufacturer's instructions. Amplification was performed in a 25 µL reaction mixture containing 2 µL (ITS) or 1.5 µL (D1/D2) of MgCl2, 0.5 µL of each primer, 0.5 µL of each dNTP, 0.125 or 0.0625 µL of Taq DNA polymerase, respectively, 5 µL of PCR reaction buffer, and 1 µL of genomic DNA standardized at 5 ng/ µL, with the remainder filled with ultra-pure sterile water. The PCR thermal cycling conditions were as follows: initial denaturation at 95 °C for 5 min (ITS) or 2 min (D1/D2), respectively, followed by 30 or 37 cycles of 95 °C for 1 min, annealing at 55 or 52 °C for 1 or 1.5 min, and elongation at 72 °C for 1 or 2 min, respectively. A final elongation step was conducted at 72 °C for 10 min.

Amplicons were separated by electrophoresis on a 2% agarose gel and visualized using a Quantum ST4 with Quantum-Capt software under UV light. Sequencing was performed by Eurofins (Germany) using the Sanger method. The obtained sequences were corrected using the Geneious software v. 11.0.18+10 (Biomatters Ltd., Auckland, New Zealand). The resulting sequences were analyzed using BLAST (Basic Local Alignment Search Tool, https://blast.ncbi.nlm.nih.gov/Blast.cgi, accessed on 9 April 2024). Phylogenetic analysis was conducted with MEGA 11 [26], employing the maximum likelihood (ML) method with a bootstrap analysis of 1000 repetitions and by applying the Tamura-Nei model. Representative sequences of the isolates of interest were submitted to the NCBI GenBank nucleotide database (http://www.ncbi.nlm.nih.gov, accessed on 4 June 2024) and accession numbers were assigned.
## 2.4. Biodegradation Assay of LDPE Films

Yeast isolates were tested for their ability to degrade LDPE films in vitro in mineral salt medium (MSM) supplemented with LDPE film as the sole carbon source [27]. A control flask containing plastic film without fungal inoculation was used for comparison.

### 2.4.1. Preparation of LDPE Films

LDPE plastic bags were cut into 0.2 g pieces, each measuring 13 × 14 cm. The pieces were disinfected with 70% ethanol for 30 min and then rinsed with sterile distilled water for 20 min to eliminate any potential contaminants.

### 2.4.2. Assessment of Yeast Capacity for LDPE Film Degradation

Yeast isolates were screened for their ability to degrade LDPE films in sterile MSM broth containing 16 g K₂HPO₄, 2 g KH₂PO₄, 1 g (NH₄)₂SO₄, 0.475 g MgSO₄, 0.2 g NaCl, 0.01 g CuSO₄, 1 g CaCO₃ and 1 g ZnSO₄ per 1000 mL of distilled water [28]. A 0.2 g LDPE film was placed into 150 mL of MSM adjusted to pH 6 in a 250 mL Erlenmeyer flask inoculated with five agar plugs of 5 mm diameter from a 7-day-old culture, and incubated at 25 °C for 30 days with continuous agitation at 150 rpm (Edmund Bühler GmbH, TH15). A negative control was used to confirm the degradation of the LDPE films [27].

## 2.5. LDPE Degradation Analysis

The LDPE film degradation was evaluated using four methods, namely weight loss, scanning electron microscopy (SEM), attenuated total reflectance–Fourier transform infrared Spectroscopy (ATR-FTIR), and X-ray diffraction (XRD) [29].

After incubation, the films were removed from their media, disinfected with 70% ethanol for 30 min, and gently cleaned to remove any fungal growth on their surface. The films were then rinsed with sterile distilled water and air-dried at room temperature before analysis [30]

### 2.5.1. Weight Loss

The weight of the LDPE films was measured every ten days over a one-month incubation period using an analytical balance (Radwag, Radom, Poland). Weight loss (W) was defined based on the initial (W0) and final (W1) weights using the following formula [31]:

$$W (\%) = \frac{W0 - W1}{W0} \times 100$$

The obtained values were compared to those of the control.

### 2.5.2. Scanning Electron Microscopy

Surface changes in the LDPE films were analyzed using a scanning electron microscope (Quanta 250). The films were mounted on carbon adhesive tapes, coated in an ionized chamber with water spray, and imaged at 1000×, 1500×, and 2000× magnifications [31,32].

### 2.5.3. Attenuated Total Reflectance–Fourier Transform Infrared Spectroscopy

The chemical structures and functional groups of the LDPE films were determined using attenuated total reflectance–Fourier transform infrared spectroscopy (ATR-FTIR) with an INVENIO-R (329) FTIR spectrometer (Bruker, Billerica, MA, USA). Each sample was analyzed with 30 scans at a 4 cm⁻¹ resolution across a wavenumber range of 500 to 5000 cm⁻¹ to identify functional groups [31,33].

Additionally, the carbonyl index (CI) was calculated as the ratio of the integrated absorbance of the carbonyl peak (C=O) from 1850 to 1650 cm⁻¹ to the methylene peak (CH₂) from 1500 to 1420 cm⁻¹. The CI values were determined using the following equation [34]:
$$CI = \frac{\text{Area under the band 1850-1650 cm}^{-1}}{\text{Area under the band 1500-1420 cm}^{-1}}$$

### 2.5.4. X-Ray Diffraction

The LDPE films were subjected to X-Ray diffraction (XRD) analysis to assess the crystallinity of the LDPE films before and after incubation. Samples were analyzed using a D8 ADVANCE X-ray diffractometer (Bruker, Germany). Each film was disinfected, mounted on a glass holder, and placed on a sample stage within the XRD chamber, where it was exposed to X-rays. The X-ray radiation was produced by a copper anticathode with a wavelength of 2θ (Cu) = 1.5418 Å. The samples were analyzed at a 2θ angle ranging from 5° to 80° [29,35].

The percentage of crystallinity was calculated using the following formula [36]:

$$ \text{Crystallinity (\%)} = \frac{T_c}{T_c + T_a} \times 100 $$

where *T*<sub>a</sub> represents the integrated area of all the peaks (amorphous and crystalline) and *T*<sub>c</sub> represents the integrated area of the crystalline peaks.

## 3. Results

### 3.1. Isolation and Molecular Characterization of Fungal Isolates

In this study, nine fungal isolates were obtained from the Boumehadjer landfill, including seven filamentous fungi and two yeast species. The subsequent analyses were specifically focused on the two yeast isolates, SLNEA04 and SLNEA05, due to their significant relevance to the research objectives. The selected isolates were identified based on two genetic markers, i.e., the ITS region and the D1/D2 region of the 26S rRNA gene.

Sequence homology analysis of the D1/D2 regions revealed that the first isolate, named SLNEA04, has a very high homology (99.63%) with Candida tropicalis NRRL Y-12968 (U45749), with 96% query coverage and an E-value of zero. The ITS region sequence (468 bp) and the D1/D2 region (484 bp) have been deposited in GenBank under accession numbers PP854593 and PP882831, respectively. A phylogenetic analysis based on the Tamura-Nei model was performed via maximum likelihood (ML) analysis of the D1/D2 sequence data from the isolate SLNEA04. As illustrated in Figure 2a, the resulting phylogenetic tree supports the classification of SLNEA04 within the Candida tropicalis group, with an ML bootstrap value of 64%.

For the second isolate, SLNEA05, sequence homology analysis of the D1/D2 region revealed 100% homology with Rhodotorula mucilaginosa CBS 2772 (KY109129) and CBS316 (NG055716), with 100% query coverage and an E-value of zero. The ITS region sequence (519 bp) and the D1/D2 region (507 bp) have been deposited in GenBank under accession numbers PP854582 and PP882834, respectively. A maximum likelihood phylogenetic analysis based on the Tamura-Nei model was conducted using the D1/D2 sequence data from the isolate SLNEA05. As illustrated in Figure 2b, the phylogenetic tree places SLNEA05 within the Rhodotorula mucilaginosa group, supported by a significant maximum likelihood (ML) value of 94%.
<figure><img src="image_6.png" /><figcaption>**Figure 2.** (a) Phylogenetic tree generated via maximum likelihood analysis of D1/D2 sequence data from *Candida* species. The tree was rooted using *Rhodotorula mucilaginosa* CBS 316. ML bootstrap support values greater than 50% are displayed near the nodes. The isolate of interest, SLNEA04, is indicated in bold. (b) Phylogenetic tree generated via maximum likelihood analysis of D1/D2 sequence data from *Rhodotorula* species. The tree was rooted using *Candida albicans* NRRL Y-12983. ML bootstrap support values greater than 50% are displayed near the nodes. The isolate of interest, SLNEA05, is indicated in bold.</figcaption></figure>
## 3.2. Determination of LDPE Film Weight Loss

Biodegradation of the LDPE films was evaluated by exposing the polymers separately to *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05 isolates. The weight loss was measured after 10, 20, and 30 days of incubation. The percentage weight loss of the LDPE films exposed to *C. tropicalis* SLNEA04 was 2.35%, 6.11%, and 7.60% after 10, 20, and 30 days, respectively. The films exposed to *R. mucilaginosa* SLNEA05 showed weight losses of 0.37%, 2.49%, and 5.53% at the same sampling times. The scatter plot indicates that both isolates showed a progressive increase in the LDPE films' weight loss, although with different trends. SLNEA04 exhibited a faster degradation rate, with a significant increase in weight loss by the 30-day mark. In contrast, SLNEA05 exhibited a slower degradation, but its weight loss steadily increased over time (Figure 3). Statistical analyses for each isolate revealed significant differences in degradation values between 10, 20, and 30 days (one-way ANOVA followed by Tukey's HSD post hoc test, *p*-values < 0.001; normality of the residuals was confirmed using the Shapiro test, and the homogeneity of variances was verified with the Bartlett test, *p*-values < 0.001). These results suggest that the yeasts were able to utilize the polymer as a sole carbon source, demonstrating their ability to degrade LDPE films. In contrast, the control LDPE film showed no weight loss throughout the incubation period.

<figure><img src="image_7.png" /><figcaption>**Figure 3.** Kinetics of LDPE film degradation by *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05 compared to the control (n = 3).</figcaption></figure>

## 3.3. Characterization of LDPE Film Biodegradation

### 3.3.1. SEM Analysis

The LDPE films were examined at 1000×, 1500×, and 2000× magnifications using a scanning electron microscope to observe the surface morphology in both the control and treated samples.

SEM analysis of the control LDPE film revealed an intact and smooth surface, with no observable morphological changes or surface alterations during incubation, confirming the material's stability under the tested conditions (Figure 4a-c). This clearly indicates that agitation at 150 rpm did not affect the integrity of the polymers. In contrast, the LDPE films exposed to *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05 displayed significant surface alterations, including pitting, roughness, and the formation of fissures
and cracks (Figure 4d-i). These changes highlight the impact of fungal activity on the polymer degradation process.

<figure><img src="image_8.png" /><figcaption>**Figure 4.** Surface morphology of LDPE films exposed to *C. tropicalis* SLNEA04 (**d–f**) and *R. mucilaginosa* SLNEA05 (**g–i**) for 30 days of incubation under SEM at 1000×; 1500×, and 2000× compared to the control (**a–c**). White arrows highlight examples of areas where surface alterations are visible.</figcaption></figure>

### 3.3.2. ATR-FTIR Analysis

The control and degraded LDPE films were subjected to ATR-FTIR analysis after 30 days of incubation in MSM, confirming the degradation of the LDPE through structural changes in the spectra when comparing the control (Figure 5a) to the treated films (Figure 5b,c).

The FTIR spectrum of the control LDPE film exhibited characteristic peaks at 2912.478, 2845.155, 1463.002, 1432.965, and 718.746 cm⁻¹ (Figure 5a). The prominent bands at 2912.478 and 2845.155 cm⁻¹ were attributed to the asymmetric and symmetric stretching vibrations of aliphatic C–H groups in the alkane molecules. The intense peaks at 1463.002, 1432.965 and 718.746 cm⁻¹ were assigned to the asymmetric scissoring vibration of C–H bonds (methylene -CH₂ group) and the angular deformation—rocking vibration—of the methylene group. Additional medium bands in the range of 500 to 874 cm⁻¹ were attributed
to vibrations of the polymer backbone structure, including C–C and C–H bond stretching, bending, and twisting within the polymer chains.

The LDPE films exposed to fungal isolates exhibited the same peaks as the control, but with variations in the intensity: higher for *C. tropicalis* SLNEA04 and lower for *R. mucilaginosa* SLNEA05. Notably, the peak at 1432.965 cm⁻¹ in the control shifted to 1425.251 cm⁻¹ (*C. tropicalis*) and 1429.108 cm⁻¹ (*R. mucilaginosa*). Additionally, new peaks of relatively low intensity emerged in the LDPE film exposed to *Candida* at 3855.907, 3740.634, 2360.633, 2335.560, 2155.619, 1697.187, 1648.971, and 1512.039 cm⁻¹, corresponding to O–H, C≡C, C=O and C=C stretching vibrations (Figure 5b). Similarly, the LDPE film exposed to *Rhodotorula* showed new peaks at 3621.951–3847.600, 2363.112–2335.560, 1700.288, 1648.414, and 1504.322 cm⁻¹, also attributed to O–H, C≡C, C=O and C=C stretching vibrations (Figure 5c).

<figure><img src="image_9.png" /><figcaption>Figure 5. *Cont.*</figcaption></figure>
<figure><img src="image_10.png" /><figcaption>**Figure 5.** ATR-FTIR spectrum analysis of LDPE films exposed to *C. tropicalis* SLNEA04 (b) and *R. mucilaginosa* SLNEA05 (c) for 30 days of incubation in MSM compared to the control (a).</figcaption></figure>

### 3.3.3. Carbonyl Index Measurement

The carbonyl index (CI) was determined using the ATR-FTIR analysis for both the control and degraded LDPE samples. The control LDPE film displayed a CI value of ~0.50. In comparison, the LDPE films exposed to *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05 for 30 days showed elevated CI values of ~0.79 and ~0.55, respectively (Table 1). The elevated CI values of the degraded LDPE films can be attributed to the enzymatic degradation processes initiated by the fungal isolates. After 30 days of exposure, the CI values increased, indicating a higher presence of carbonyl groups (C=O) formed as the fungal enzymes cleaved the polymer chains and facilitated oxidative degradation. The more significant increase in the CI for the *C. tropicalis* SLNEA04 treatment suggests that this strain may be more effective at breaking down LDPE, resulting in a greater accumulation of carbonyl functionalities. These findings reflect the effectiveness of microbial action in altering the chemical composition of the LDPE films, confirming the extent of the degradation facilitated by these microorganisms.

<table><caption>**Table 1.** Carbonyl index of the control and degraded LDPE films.</caption><thead><tr><td>LDPE Samples</td><td>Carbonyl Index (CI)</td></tr></thead><tbody><tr><td>Control</td><td>0.504906</td></tr><tr><td>Exposed to <i>C. tropicalis</i> SLNEA04</td><td>0.790574</td></tr><tr><td>Exposed to <i>R. mucilaginosa</i> SLNEA04</td><td>0.553643</td></tr></tbody></table>

### 3.3.4. XRD Analysis and Determination of Crystallinity Percentage

X-ray diffraction (XRD) analysis was performed on the LDPE films to assess the crystallinity changes resulting from potential fungal degradation. The XRD profiles revealed a semi-crystalline structure in the LDPE films, with the crystalline phase exhibiting the typical intense and sharply defined diffraction peaks. The XRD spectrum of the control film showed characteristic peaks of the crystalline regions at angular positions (2θ) 11.20°, 21.26°, 23.71°, 27.11°, 29.11°, 31.26°, 35.96°, and 47.37° (Figure 6a).

Figure 6b illustrates the XRD profile of the LDPE film exposed to *C. tropicalis* SLNEA04 for 30 days compared to the control. This analysis revealed the same peaks as the control
film, with higher intensities at 11.20°, 21.26° and 31.26°, but shifts in the peaks at 23.71°, 27.11°, 29.11°, and 47.37° to 23.61°, 27.06°, 29.06°, and 47.12°, respectively.

Figure 6c shows the XRD profile of the LDPE film exposed to *R. mucilaginosa* SLNEA05 for one month compared to the control. This analysis also revealed the presence of the same peaks as the control, with higher intensity at 11.20° and lower intensity at 31.26°. However, the peaks at 21.26°, 23.71°, 27.11°, 29.11°, 35.96°, and 47.37° shifted to 21.36°, 23.66°, 27.26°, 29.21°, 36.16°, and 47.42°, respectively. Additionally, two new peaks appeared at 43.82° and 56.38°.

<figure><img src="image_11.png" /><figcaption>Figure 6. *Cont.*</figcaption></figure>
<figure><img src="image_12.png" /><figcaption>**Figure 6.** XRD spectrum of LDPE films exposed to *C. tropicalis* SLNEA04 (b) and *R. mucilaginosa* SLNEA05 (c) for 30 days of incubation compared to the control (a). Asterisks (*) indicate new peaks that appeared following degradation.</figcaption></figure>

The crystallinity of the LDPE samples exposed to fungal isolates was compared to the control, which exhibited a crystallinity of 37.4%. The LDPE films incubated with *Candida* and *Rhodotorula* for one month showed higher crystallinity rates, estimated at 53.8% and 41.7%, respectively (Table 2).

<table><caption>**Table 2.** Crystallinity of the control and degraded LDPE films.</caption><thead><tr><td>LDPE Samples</td><td>Crystallinity Percentage (%)</td></tr></thead><tbody><tr><td>Control</td><td>37.4</td></tr><tr><td>Exposed to <i>C. tropicalis</i> SLNEA04</td><td>53.8</td></tr><tr><td>Exposed to <i>R. mucilaginosa</i> SLNEA04</td><td>41.7</td></tr></tbody></table>

## 4. Discussion

The persistent accumulation of LDPE in the environment highlights the urgent need for innovative strategies, including natural ones like bioremediation. This study evaluated the biodegradation potential of two fungal species isolated from the Boumehadjer landfill in northeastern Algeria. Fungi were targeted for their known enzymatic capabilities and their adaptability to various environmental conditions. Through controlled *in vitro* experiments, we evaluated their efficacy in degrading LDPE plastic bag films using various advanced analytical techniques, including SEM, ATR-FTIR, and XRD, to characterize the degradation process and elucidate the underlying mechanisms.

In this context, the yeast isolates *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05 were obtained and identified based on ITS and D1/D2 rRNA gene sequencing. Both fungal isolates were able to utilize LDPE as the sole carbon and energy source by exhibiting a significant weight loss of 7.60% and 5.53% after 30 days of incubation in MSM (pH 6, 25 °C, 150 rpm) for *C. tropicalis* and *R. mucilaginosa*, respectively (one-way ANOVA followed by Tukey's HSD post hoc test, *p*-values < 0.001). This weight loss corresponds to the surface deterioration due to direct contact between the fungal cells and the LDPE polymer, leading to its consumption [29].
The observed weight loss in the LDPE samples is primarily attributed to the degrada-
dation of the polymer's carbon skeleton by fungal enzymes. Fungi produce a range of
enzymes capable of breaking the chemical bonds of plastic polymers, allowing them to
be used as a carbon source. Upon adhesion and colonization on the LDPE film surface,
facilitated by the production of biosurfactants, fungal microorganisms secrete extracellular
enzymes such as hydrolases (cutinases, carboxylesterases, proteases, amidases) and oxi-
doreductases (laccases and peroxidases). These enzymes depolymerize the long carbon
chains of the polymer into simpler oligomers, dimers and monomers, which are more easily
metabolized. These monomers can then be transported across the cell membrane, entering
the fungal cell and integrating into metabolic pathways for assimilation [37–39].

The non-specificity of fungal exoenzymes enables them to decompose various plastic polymers [1], thereby enhancing cellular assimilation [40]. The primary fungal enzymes involved in PE biodegradation are lignolytic enzymes such as laccases (Lac, EC 1.10.3.2) and peroxidases (EC 1.11.1.7) [41–44]. Santacruz-Juárez et al. [41] proposed hypothetical pathways for the enzymatic degradation of PE under aerobic conditions by fungal microorganisms. One such pathway focuses on laccases, in combination with the action of hydrophobins acting as biosurfactants, thereby enabling enzymatic catalysis. In this hypothetical pathway, laccases, possessing a high redox potential, lead to the oxidation of neighboring molecules, generating PE radicals. These radicals are subsequently cleaved to form alkane radicals, which are oxidized to alcohols, then to aldehydes, and finally to carboxylic acids that enter the Krebs cycle to be mineralized into CO2 and H2O. Furthermore, the degradation of PE by the laccase enzyme produced by *Pleurotus ostreatus* and *Aspergillus flavus* has been reported by Gomez-Mendez et al. [45] and Zhang et al. [40], respectively.

Other fungal enzymes have also been identified in the degradation of plastic polymers. Bholay et al. [46] reported the capacity of fungal species such as *Aspergillus flavus*, *Aspergillus niger*, and *Fusarium graminearum* to produce manganese peroxidase and lignin peroxidase, inducing PE bag biodegradation, with *Aspergillus niger* and *Aspergillus flavus* showing the highest lignin peroxidase production. These fungal enzymes generate new functional groups, such as hydroxyl, carbonyl and carboxyl groups, that increase the hydrophilicity of the polymer, enhancing its biodegradability [47]. Additionally, Abraham et al. [48] and Awasthi et al. [21] observed an increase in the amount of carbonyl groups in LDPE when exposed to *Aspergillus nomius* and *Rhizopus oryzae*, respectively, which is directly linked to peroxidase activity [49].

Despite the lack of specific identification of the enzymes secreted by yeasts involved
in PE degradation, enzymes involved in the degradation of other polymers have been
reported. Species of the *Candida* genus are known to produce various extracellular enzymes
capable of degrading large macromolecules into smaller ones [50].

Furthermore, Hadibarata et al. [51] isolated a novel strain, Candida sp. S1, from a tropi-
cal rainforest and reported its ability to degrade pyrene by 35% after 15 days of incubation.
Farag and Soliman [52] demonstrated the capacity of Candida tropicalis to degrade petroleum
oil as well as certain hydrocarbons. Additionally, some species of the Saccharomycetes
class have been extensively used in industrial and biotechnological processes, with genera
such as Arxula, Candida, and Debaryomyces implicated in plastic degradation [53]. Zahari
et al. [54] confirmed the ability of C. tropicalis, obtained from the Animal Physiology Lab
(Malaysia Sabah University), to degrade LDPE strips (5 cm × 3 cm), with a 3.2% weight
loss after seven weeks of incubation in Ramsay broth at 30 °C, under continuous agitation
in a rotary shaker (200 rpm). This research also provided evidence of microbial growth
on the plastic surface, CO2 evolution, and notable changes in the polymer's topography.
In contrast, our study, utilizing C. tropicalis isolated from a local landfill, demonstrated a
higher weight loss of 7.60% after just one month of incubation. This suggests that isolates
from contaminated environments may have developed more effective plastic degradation
mechanisms. Moreover, species such as Saccharomyces cerevisiae are model organisms in
many studies [55]. In addition, research by Vaksmaa et al. [56] demonstrated the potential
of Rhodotorula mucilaginosa isolated from the marine environment to mineralize and assim-
ilate carbon from PE. This was evidenced by stable isotope tracing with ¹³C-labeled PE, which was incorporated into the fungal biomass.

Biodegradation of LDPE films exposed to *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05 was further confirmed by SEM, ATR-FTIR, and XRD analysis. The SEM images provided detailed insights into the surface morphology changes induced by the fungal isolates, revealing their substantial biodegradation potential. SEM analysis showed that both yeasts notably altered the surface of the polymer, with *C. tropicalis* SLNEA04 causing more pronounced surface erosion and fragmentation compared to *R. mucilaginosa* SLNEA05, particularly at higher magnifications (2000×). Initial signs of surface roughness and fragmentation were observed for both fungi at 1000× magnification. As the magnification increased to 1500× and 2000×, the extent of the surface degradation became more evident, with SLNEA04 exhibiting deeper and more extensive cracks and voids. These observations suggest that both fungi produce extracellular enzymes capable of breaking down the polymer matrix. This enzymatic activity likely involves oxidative and hydrolytic processes, which weaken the polymer chains and facilitate fragmentation. The extent of the surface degradation observed in the SEM images correlates with the biodegradation efficiency of the respective fungi. Our findings align with those of Giyahchi and Moghimi [57], who have observed fungal biofilm formation along with distinct pits and holes on the surface of polyurethane films (2 cm × 2 cm) after 30 days of exposure to *Exophiala* sp. NS-7 and *Rhodotorula* sp. NS-12 at 30 °C in a shaker incubator set at 120 rpm. These surface alterations demonstrated the ability of these strains to fragment polyurethane. Similarly, Khan et al. [58] investigated the degradation of LDPE by *Penicillium citrinum* and observed the formation of cavities and cracks on the surface of the polymer under SEM after 90 days of incubation. Other studies have also reported surface changes in LDPE films following fungal degradation [27,33,59–61].

To further substantiate the SEM findings, ATR-FTIR analysis was conducted to investigate the chemical changes in the LDPE films exposed to the two yeasts, providing additional insights by identifying specific functional groups and alterations resulting from fungal activity [29,62,63]. ATR-FTIR analysis revealed an increase in the intensity of the absorption bands at 2912, 2845, 1463 and 718 cm⁻¹ in the LDPE film exposed to *C. tropicalis* SLNEA04, attributed to the cleavage of C–H bonds and the formation of new intermediate products [64]. In contrast, the decrease in the intensity of these bands in the LDPE film exposed to *R. mucilaginosa* SLNEA05 reflects lower degradation and fewer new chemical intermediates compared to those observed with *C. tropicalis* SLNEA04. The appearance of peaks in the 3621–3855 cm⁻¹ and 2155–2363 cm⁻¹ regions in the degraded samples indicates that O–H (alcohol) and C≡C (alkyne) bonds have been altered, leading to the formation of carboxyl compounds. Bands observed at 1697 and 1700 cm⁻¹, in films exposed to SLNEA04 and SLNEA05, respectively, are related to the elongation of C=O groups in aldehydes and ketones. The formation of these carbonyl groups suggests biological hydrolysis activity by depolymerases and results from oxidation by oxidoreductases. Das and Kumar [65] noted that the formed carbonyl groups can be attacked by microorganisms. Hasan et al. [66] attributed the changes observed at the 1812.8 cm⁻¹ band to the utilization of carbonyl groups by fungi during LDPE biodegradation.

Furthermore, the bands appearing at 1648, 1512, 1504, 1429, and 1425 cm⁻¹ correspond to the elongation of C=C bonds in alkenes and the deformation vibrations of C–H bonds in methylene groups, respectively. These modifications confirm the biodegradation and the formation of new intermediate products, indicating the oxidation of LDPE by enzyme-mediated fungal activity [67]. ATR-FTIR analysis thus showed that LDPE films were degraded by *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05 through enzymatic hydrolysis, favoring microbial action [27]. Additionally, the carbonyl index (CI) value of films exposed to both yeasts was higher than that of the control film, confirming that oxidative changes occurred during the degradation of LDPE films by fungal isolates [68]. The oxidation of LDPE introduces carbonyl groups into the polymer chain, leading to the formation of carboxylic acids and other low-molecular-weight products. These mod-
ifications make the material more hydrophilic, thereby increasing and accelerating the degradation rate. Thus, the conformational changes observed in the absorption spectra of the degraded LDPE films confirm the depolymerization of these polymers and their use as a carbon source [13].

The structure of the LDPE films was further examined using XRD to examine the changes in crystallinity after exposure to the two fungal isolates. The obtained spectra showed high intensity diffraction peaks in the 2θ range of 20 to 30°, confirming the semi-crystalline structure of the tested polymer. The peaks observed at 21.26° (control and SLNEA04) and 21.36° (SLNEA05), representing (110) reflections, and the peaks at 2θ = 23.61° (SLNEA04), 23.66° (SLNEA05), and 23.71° (control), corresponding to (200) reflections [69], are attributed to the orthorhombic crystalline structure of LDPE [70]. Moreover, changes in the XRD profile of the LDPE films were observed after biodegradation, including increases or decreases in the intensity of certain peaks, increases in the crystallinity rate, shifts in some peaks, and the appearance of new bands. These alterations are consistent with the results reported by El Sayed et al. [29], who have observed changes in the crystalline structure of thermally treated LDPE films exposed to a mixed culture of two fungal isolates, *Aspergillus carbonarius* and *A. fumigatus*, over 16 weeks. Notable observations included a shift in the peak positions to 2θ = 21.6, 24.06 et 36.39°, the disappearance of the peak at 2θ = 69.558°, and a decrease in the intensity of the peaks at 2θ = 21.5 and 36.12°, reflecting a reduction in the crystal size and crystallinity during biodegradation [29]. In contrast, another study by Manzur et al. [71] revealed an increase in the relative crystallinity of LDPE after 3 months of incubation with a consortium of four filamentous fungi: *Aspergillus niger*, *Gliocladium virens*, *Penicillium pinophilum*, and *Phanerochaete chrysosporium*. Microorganisms typically act on semi-crystalline polymers in the amorphous phases, as crystalline regions are more resistant to enzymatic attacks [72]. The amorphous fraction surrounds the crystalline particles and/or delineates the crystalline blocks in the crystalline mosaic [73]. Our results suggest that both yeasts, *C. tropicalis* SLNEA04 and *R. mucilaginosa* SLNEA05, primarily target the amorphous phase, leading to increased crystallinity and the fragmentation of crystalline blocks, resulting in smaller crystals. However, these crystals located at the amorphous-crystalline interface of the polymer or formed during the initial stage may be assimilated by the fungal species, leading to an increase in the amorphous fraction and a decrease in the crystalline fraction [71,74]. Furthermore, the shifts in the diffraction peak positions result from minor changes in the dimensions of the crystal lattice. Finally, the type of biological treatment may account for the observed differences in crystallinity [10]. Notably, the LDPE film exposed to SLNEA04 exhibited the highest percentage of crystallinity, estimated at 53.8%, compared to the film exposed to SLNEA05 (41.7%) and the control (37.4%).

Finally, recent research has increasingly highlighted the diversity, abundance, and activity of microorganisms capable of plastic degradation. These findings underscore the critical need for a deeper understanding of the factors influencing microbial biodegradation, which is essential to accelerate the process [75].

**5. Conclusions**

Our study highlights the biodegradation potential of two yeast isolates, *Candida tropicalis* SLNEA04 and *Rhodotorula mucilaginosa* SLNEA05, obtained from a northeastern Algerian landfill, in degrading LDPE films *in vitro*. The degradation process was substantiated through a combination of analytical methods, including significant weight loss in the LDPE films—7.60% for SLNEA04 and 5.53% for SLNEA05—along with surface erosion and morphological changes observed under scanning electron microscopy (SEM). Additionally, new functional groups were detected by Fourier transform infrared spectroscopy (FTIR), and an increase in crystallinity was assessed using X-ray diffractometry (XRD). These findings offer valuable insights into the role of yeasts in polyethylene biodegradation, a field that has seen limited exploration compared to bacterial and fungal studies. Despite the promotion of biodegradable plastics, their high costs impede widespread adoption,
leading to continued reliance on conventional plastics. Our results suggest that yeasts could play a significant role in PE degradation and serve as a basis for developing biological solutions to address plastic waste. To further enhance our understanding of the degradation mechanisms, future research should incorporate additional analytical techniques, such as pyrolysis temperature analysis, water contact angle measurements, and atomic force microscopy (AFM). These methods could provide complementary information on the thermal, surface, and morphological changes occurring during the degradation process. Moreover, extending the incubation periods would allow for the investigation of the long-term effects of yeast activity on PE degradation and the assessment of the synergistic effects when using both yeast isolates in combination, potentially advancing the development of efficient and cost-effective biodegradation systems for plastic waste management.

**Author Contributions:** Conceptualization, R.H. and G.B. (Ghania Bourzama); methodology, R.H., G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud) and A.B.; validation, G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud) and E.C.; formal analysis, R.H., G.B. (Gaëtan Burgaud) and A.B.; investigation, R.H., G.B. (Ghania Bourzama) and G.B. (Gaëtan Burgaud); resources, R.H., G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud) and E.C.; data curation, R.H. and G.B. (Ghania Bourzama); writing—original draft preparation, R.H.; writing—review and editing, R.H., G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud) and E.C.; visualization, R.H., G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud) and E.C.; supervision, G.B. (Ghania Bourzama), G.B. (Gaëtan Burgaud), E.C. and B.S.; project administration, G.B. (Ghania Bourzama). All authors have read and agreed to the published version of the manuscript.

**Funding:** The authors thank the Ministry of Higher Education and Scientific Research (Algeria) and the French National Research Agency (ANR-19-CE04-0001-01 MycoPLAST project) for the funding.

**Institutional Review Board Statement:** Not applicable.

**Data Availability Statement:** Data is contained within the article.

**Acknowledgments:** The authors thank the National Higher School of Mines and Metallurgy, Annaba (Algeria), the General Directorate for Scientific Research and Technological Development (Algeria), the National Polytechnic School of Constantine -Malek Bennabi (Algeria), the University Laboratory of Biodiversity and Microbial Ecology (France) and the National School of Advanced Sciences and Technologies of Borj Cedria (Tunisia).

**Conflicts of Interest:** The authors declare no conflicts of interest.

## References

1. Temporiti, M.E.E.; Nicola, L.; Nielsen, E.; Tosi, S. Fungal Enzymes Involved in Plastics Biodegradation. *Microorganisms* **2022**, *10*, 1180. [CrossRef] [PubMed]

2. Philippe, A.; Noël, C.; Eyheraguibel, B.; Briand, J.F.; Paul-Pont, I.; Ghiglione, J.F.; Coton, E.; Burgaud, G. Fungal Diversity and Dynamics during Long-Term Immersion of Conventional and Biodegradable Plastics in the Marine Environment. *Diversity* **2023**, *15*, 579. [CrossRef]

3. Bautista-Zamudio, P.A.; Flórez-Restrepo, M.A.; López-Legarda, X.; Monroy-Giraldo, L.C.; Segura-Sánchez, F. Biodegradation of Plastics by White-Rot Fungi: A Review. *Sci. Total Environ.* **2023**, *901*, 165950. [CrossRef] [PubMed]

4. Li, S.; Yang, Y.; Yang, S.; Zheng, H.; Zheng, Y.; Jun, M.; Nagarajan, D.; Varjani, S.; Chang, J.S. Recent Advances in Biodegradation of Emerging Contaminants—Microplastics (MPs): Feasibility, Mechanism, and Future Prospects. *Chemosphere* **2023**, *331*, 138776. [CrossRef]

5. Sangale, M.K.; Shahnawaz, M.; Ade, A.B. Potential of Fungi Isolated from the Dumping Sites Mangrove Rhizosphere Soil to Degrade Polythene. *Sci. Rep.* **2019**, *9*, 5390. [CrossRef]

6. Agence Nationale des Déchets (AND). *Les Déchets Plastiques En Algérie: Regard Croisé Sur Les Plastiques à Usage Unique. Le Bulletin de Veille Technologique N°02-2021*; AND: Mohamed Belouizdad, Algeria, 2020.

7. Zeghal, E.; Vaksmaa, A.; Vielfaure, H.; Boekhout, T.; Niemann, H. The Potential Role of Marine Fungi in Plastic Degradation—A Review. *Front. Mar. Sci.* **2021**, *8*, 738877. [CrossRef]

8. Gao, R.; Liu, R.; Sun, C. A Marine Fungus *Alternaria alternata* FB1 Efficiently Degrades Polyethylene. *J. Hazard. Mater.* **2022**, *431*, 128617. [CrossRef] [PubMed]

9. Sheik, S.; Chandrashekar, K.R.; Swaroop, K.; Somashekarappa, H.M. Biodegradation of Gamma Irradiated Low Density Polyethylene and Polypropylene by Endophytic Fungi. *Int. Biodeterior. Biodegrad.* **2015**, *105*, 21–29. [CrossRef]

10. Volke-Seplveda, T.; Saucedo-Castaeda, G.; Gutirrez-Rojas, M.; Manzur, A.; Favela-Torres, E. Thermally Treated Low Density Polyethylene Biodegradation by *Penicillium pinophilum* and *Aspergillus niger*. *J. Appl. Polym. Sci.* **2002**, *83*, 305–314. [CrossRef]
11. Zahra, S.; Abbas, S.S.; Mahsa, M.T.; Mohsen, N. Biodegradation of Low-Density Polyethylene (LDPE) by Isolated Fungi in Solid Waste Medium. *Waste Manag.* **2010**, *30*, 396–401. [CrossRef]

12. Gong, Z.; Jin, L.; Yu, X.; Wang, B.; Hu, S.; Ruan, H.; Sung, Y.J.; Lee, H.G.; Jin, F. Biodegradation of Low Density Polyethylene by the Fungus *Cladosporium* sp. Recovered from a Landfill Site. *J. Fungi* **2023**, *9*, 605. [CrossRef] [PubMed]

13. Nowak, B.; Pajak, J.; Drozd-Bratkowicz, M.; Rymarz, G. Microorganisms Participating in the Biodegradation of Modified Polyethylene Films in Different Soils under Laboratory Conditions. *Int. Biodeterior. Biodegrad.* **2011**, *65*, 757–767. [CrossRef]

14. Pramila, R.; Ramesh, K.V. Biodegradation of Low Density Polyethylene (LDPE) by Fungi Isolated from Marine Water– a SEM Analysis. *Afr. J. Microbiol. Res.* **2011**, *5*, 5013–5018. [CrossRef]

15. Paço, A.; Duarte, K.; da Costa, J.P.; Santos, P.S.M.; Pereira, R.; Pereira, M.E.; Freitas, A.C.; Duarte, A.C.; Rocha-Santos, T.A.P. Biodegradation of Polyethylene Microplastics by the Marine Fungus *Zalerion maritimum*. *Sci. Total Environ.* **2017**, *586*, 10–15. [CrossRef]

16. Mohan, S.K.; Suresh, B. Studies on Biodegradation of Plastics by *Aspergillus* sp. Isolated from Dye Effluent Enriched Soil. *Indo Am. J. Pharm. Sci.* **2015**, *2*, 1636–1639.

17. Shruthi, B.; Deepa, N.; Somashekaraiah, R.; Adithi, G.; Divyashree, S.; Sreenivasa, M.Y. Exploring Biotechnological and Functional Characteristics of Probiotic Yeasts: A Review. *Biotechnol. Rep.* **2022**, *34*, e00716. [CrossRef]

18. Muthumani, S.; Anbuselvi, S. Waste Management and Enviornment Protection-A Study with Isolation and Characterisation of Low Density Polyethylene Degradating Microbes. *Int. J. ChemTech Res. CODEN* **2014**, *7*, 2841–2845.

19. Gajendiran, A.; Krishnamoorthy, S.; Abraham, J. Microbial Degradation of Low-Density Polyethylene (LDPE) by *Aspergillus clavatus* Strain JASK1 Isolated from Landfill Soil. *3 Biotech* **2016**, *6*, 52. [CrossRef]

20. Duan, Y.; Yin, Y.; Ni, Z.; Liu, J.; Gui, H.; Wu, D.; Wu, X.; Wang, L. The Effective and Green Biodegradation of Polyethylene Microplastics by the Screening of a Strain with Its Degrading Enzymes. *Biochem. Eng. J.* **2024**, *210*, 109429. [CrossRef]

21. Awasthi, S.; Srivastava, N.; Singh, T.; Tiwary, D.; Mishra, P.K. Biodegradation of Thermally Treated Low Density Polyethylene by Fungus *Rhizopus oryzae* NS 5. *3 Biotech* **2017**, *7*, 73. [CrossRef]

22. Chaudhary, A.K.; Chaitanya, K.; Dalmia, R.; Vijayakumar, R.P. Synergistic Effect of UV, Thermal, and Chemical Treatment on Biological Degradation of Low-Density Polyethylene (LDPE) by *Thermomyces lanuginosus*. *Environ. Monit. Assess.* **2021**, *193*, 513. [CrossRef] [PubMed]

23. Davet, P.; Rouxf, F. *Détection et Isolement Des Champignons Du Sol*; INRA Editions: Paris, France, 1997.

24. Gardes, M.; Bruns, T.D. ITS Primers with Enhanced Specificity for Basidiomycetes—Application to the Identification of Mycorrhizae and Rusts. *Mol. Ecol.* **1993**, *2*, 113–118. [CrossRef] [PubMed]

25. Kurtzman, C.P.; Robnett, C.J. Identification of Clinically Important Ascomycetous Yeasts Based on Nucleotide Divergence in the 5 End of the Large-Subunit (26S) Ribosomal DNA Gene. *J. Clin. Microbiol.* **1997**, *35*, 1216–1223. [CrossRef]

26. Tamura, K.; Stecher, G.; Kumar, S. MEGA11: Molecular Evolutionary Genetics Analysis Version 11. *Mol. Biol. Evol.* **2021**, *38*, 3022–3027. [CrossRef]

27. Khruengsai, S.; Sripahco, T.; Pripdeevech, P. Low-Density Polyethylene Film Biodegradation Potential by Fungal Species from Thailand. *J. Fungi* **2021**, *7*, 594. [CrossRef]

28. Nakajima-Kambe, T.; Shigeno-Akutsu, Y.; Nomura, N.; Onuma, F.; Nakahara, T. Microbial Degradation of Polyurethane, Polyester Polyurethanes and Polyether Polyurethanes. *Appl. Microbiol. Biotechnol.* **1999**, *51*, 134–140. [CrossRef]

29. El-Sayed, M.T.; Rabie, G.H.; Hamed, E.A. Biodegradation of Low-Density Polyethylene (LDPE) Using the Mixed Culture of *Aspergillus carbonarius* and *A. fumigatus*. *Environ. Dev. Sustain.* **2021**, *23*, 14556–14584. [CrossRef]

30. Khruengsai, S.; Sripahco, T.; Pripdeevech, P. Microbial Degradation of Low-Density Polyethylene by *Neopetalotiopsis phangngaensis*. *J. Gen. Appl. Microbiol.* **2022**, *68*, 287–294. [CrossRef] [PubMed]

31. DSouza, G.C.; Sheriff, R.S.; Ullanat, V.; Shrikrishna, A.; Joshi, A.V.; Hiremath, L.; Entoori, K. Fungal Biodegradation of Low-Density Polyethylene Using Consortium of *Aspergillus* Species under Controlled Conditions. *Heliyon* **2021**, *7*, e07008. [CrossRef]

32. Černoša, A.; Cortizas, A.M.; Traoré, M.; Podlogar, M.; Danevčič, T.; Gunde-Cimerman, N.; Gostinčar, C. A Screening Method for Plastic-Degrading Fungi. *Heliyon* **2024**, *10*, e31130. [CrossRef]

33. Kučić Grgić, D.; Miloloža, M.; Ocelić Bulatović, V.; Ukić, Š.; Slouf, M.; Gajdosova, V. Screening the Efficacy of a Microbial Consortium of Bacteria and Fungi Isolated from Different Environmental Samples for the Degradation of LDPE/TPS Films. *Separations* **2023**, *10*, 79. [CrossRef]

34. Almond, J.; Sugumaar, P.; Wenzel, M.N.; Hill, G.; Wallis, C. Determination of the Carbonyl Index of Polyethylene and Polypropylene Using Specified Area under Band Methodology with ATR-FTIR Spectroscopy. *E-Polym. 2020*, *20*, 369–381. [CrossRef]

35. Nasrabadi, A.E.; Ramavandi, B.; Bonyadi, Z. Recent Progress in Biodegradation of Microplastics by *Aspergillus* sp. in Aquatic Environments. *Colloids Interface Sci. Commun.* **2023**, *57*, 100754. [CrossRef]

36. Kyaw, B.M.; Champakalakshmi, R.; Sakharkar, M.K.; Lim, C.S.; Sakharkar, K.R. Biodegradation of Low Density Polythene (LDPE) by *Pseudomonas* Species. *Indian J. Microbiol.* **2012**, *52*, 411–419. [CrossRef]

37. Srikanth, M.; Sandeep, T.S.R.S.; Sucharitha, K.; Godi, S. Biodegradation of Plastic Polymers by Fungi: A Brief Review. *Bioresour. Bioprocess.* **2022**, *9*, 42. [CrossRef]

38. Puliga, F.; Zuffi, V.; Baldo, D.; Cavatorta, D.; Zambonelli, A.; Francioso, O.; Sanchez-Cortes, S. *Cladosporium cladosporioides* (Strain Clc/1): A Candidate for Low-Density Polyethylene Degradation. *Chem. Biol. Technol. Agric.* **2023**, *10*, 50. [CrossRef]
39. Varshney, S.; Gupta, V.; Yadav, A.N.; Rahi, R.K.; Devki; Neelam, D. K. An Overview on Role of Fungi in Systematic Plastic Degradation. *J. Appl. Biol. Biotechnol.* **2023**, *11*, 61–69. [CrossRef]

40. Zhang, J.; Gao, D.; Li, Q.; Zhao, Y.; Li, L.; Lin, H.; Bi, Q.; Zhao, Y. Biodegradation of Polyethylene Microplastic Particles by the Fungus *Aspergillus flavus* from the Guts of Wax Moth *Galleria mellonella*. *Sci. Total Environ.* **2020**, *704*, 135931. [CrossRef] [PubMed]

41. Santacruz-Juárez, E.; Buendia-Corona, R.E.; Ramírez, R.E.; Sánchez, C. Fungal Enzymes for the Degradation of Polyethylene: Molecular Docking Simulation and Biodegradation Pathway Proposal. *J. Hazard. Mater.* **2021**, *411*, 125118. [CrossRef]

42. Malachová, K.; Novotný, Č.; Adamus, G.; Lotti, N.; Rybková, Z.; Soccio, M.; Šlosarčíková, P.; Verney, V.; Fava, F. Ability of *Trichoderma hamatum* Isolated from Plastics-Polluted Environments to Attack Petroleum-Based, Synthetic Polymer Films. *Processes* **2020**, *8*, 467. [CrossRef]

43. Sowmya, H.V.; Ramalingappa, Krishnappa, M.; Thippeswamy, B. Degradation of Polyethylene by *Penicillium* simplicissimum Isolated from Local Dumpsite of Shivamogga District. *Environ. Dev. Sustain.* **2015**, *17*, 731–745. [CrossRef]

44. Shimao, M. Biodegradation of Plastics. *Curr. Opin. Biotechnol.* **2001**, *12*, 242–247. [CrossRef]

45. Gómez-Méndez, L.D.; Moreno-Bayona, D.A.; Poutou-Piñales, R.A.; Salcedo-Reyes, J.C.; Pedroza-Rodríguez, A.M.; Vargas, A.; Bogoya, J.M. Biodeterioration of Plasma Pretreated LDPE Sheets by *Pleurotus ostreatus*. *PLoS ONE* **2018**, *13*, e0203786. [CrossRef] [PubMed]

46. Bholay, A.D.; Bhorkhataria, B.V.; Jadhav, P.U.; Kaveri, S.P.; Mayuri, D.; Pravin, M.N.; Nalawade, P.M. Bacterial Lignin Peroxidase: A Tool for Biobleaching and Biodegradation of Industrial Effluents. *Univers. J. Environ. Res. Technol.* **2012**, *2*, 58–64.

47. Shah, A.A.; Hasan, F.; Hameed, A.; Ahmed, S. Biological Degradation of Plastics: A Comprehensive Review. *Biotechnol. Adv.* **2008**, *26*, 246–265. [CrossRef]

48. Abraham, J.; Ghosh, E.; Mukherjee, P.; Gajendiran, A. Microbial Degradation of Low Density Polyethylene. *Environ. Prog. Sustain. Energy* **2017**, *36*, 147–154. [CrossRef]

49. Sudhakar, M.; Doble, M.; Murthy, P.S.; Venkatesan, R. Marine Microbe-Mediated Biodegradation of Low- and High-Density Polyethylenes. *Int. Biodeterior. Biodegrad.* **2008**, *61*, 203–213. [CrossRef]

50. Neji, S.; Hadrich, I.; Trabelsi, H.; Abbes, S.; Cheikhrouhou, F.; Sellami, H.; Makni, F.; Ayadi, A. Virulence Factors, Antifungal Susceptibility and Molecular Mechanisms of Azole Resistance among *Candida parapsilosis* Complex Isolates Recovered from Clinical Specimens. *J. Biomed. Sci.* **2017**, *24*, 37. [CrossRef]

51. Hadibarata, T.; Khudhair, A.B.; Kristanti, R.A.; Kamyab, H. Biodegradation of Pyrene by *Candida* sp. S1 under High Salinity Conditions. *Bioprocess. Biosyst. Eng.* **2017**, *40*, 1411–1418. [CrossRef]

52. Farag, S.; Soliman, N.A. Biodegradation of Crude Petroleum Oil and Environmental Pollutants by *Candida tropicalis* Strain. *Braz. Arch. Biol. Technol.* **2011**, *54*, 821–830. [CrossRef]

53. Ekanayaka, A.H.; Tibpromma, S.; Dai, D.; Xu, R.; Suwannarach, N.; Stephenson, S.L.; Dao, C.; Karunarathna, S.C. A Review of the Fungi That Degrade Plastic. *J. Fungi* **2022**, *8*, 772. [CrossRef] [PubMed]

54. Zahari, N.Z.; Abdullah, S.N.; Tuah, P.M.; Cleophas, F.N. Biodegradation of Low-Density Polyethylene (LDPE) and Starch—Based Plastic (SBP) by Thermophiles *Bacillus subtilis* and *Candida tropicalis*. *IOP Conf. Ser. Mater. Sci. Eng.* **2021**, *1173*, 012035. [CrossRef]

55. Suh, S.O.; Blackwell, M.; Kurtzman, C.P.; Lachance, M.A. Phylogenetics of Saccharomycetales, the Ascomycete Yeasts. *Mycologia* **2006**, *98*, 1006–1017. [CrossRef] [PubMed]

56. Vaksmaa, A.; Polerecky, L.; Dombrowski, N.; Kienhuis, M.V.M.; Posthuma, I.; Gerritse, J.; Boekhout, T.; Niemann, H. Polyethylene Degradation and Assimilation by the Marine Yeast *Rhodotorula mucilaginosa*. *ISME Commun.* **2023**, *3*, 68. [CrossRef]

57. Giyahchi, M.; Moghimi, H. Aerobic Biodegradation of Untreated Polyester-Polyether Urethanes by Newly Isolated Yeast Strains *Exophilia* sp. NS-7 and *Rhodotorula* sp. NS-12. *Sci. Rep.* **2023**, *13*, 5016. [CrossRef]

58. Khan, S.; Ali, S.A.; Ali, A.S. Biodegradation of Low Density Polyethylene (LDPE) by Mesophilic Fungus 'Penicillium citrinum' Isolated from Soils of Plastic Waste Dump Yard, Bhopal, India. *Environ. Technol.* **2023**, *44*, 2300–2314. [CrossRef]

59. Ameen, F.; Moslem, M.; Hadi, S.; Al-Sabri, A.E. Biodegradation of Low Density Polyethylene (LDPE) by Mangrove Fungi from the Red Sea Coast. *Progress. Rubber Plast. Recycl. Technol.* **2015**, *31*, 125–143. [CrossRef]

60. Alshehrei, F. Biodegradation of Low Density Polyethylene by Fungi Isolated from Red Sea Water. *Int. J. Curr. Microbiol. Appl. Sci.* **2017**, *6*, 1703–1709. [CrossRef]

61. Verma, N.; Gupta, S. Assessment of LDPE Degrading Potential *Aspergillus* Species Isolated from Municipal Landfill Sites of Agra. *SN Appl. Sci.* **2019**, *1*, 701. [CrossRef]

62. Montazer, Z.; Habibi-Najafi, M.B.; Mohebbi, M.; Oromiehei, A. Microbial Degradation of UV-Pretreated Low-Density Polyethylene Films by Novel Polyethylene-Degrading Bacteria Isolated from Plastic-Dump Soil. *J. Polym. Environ.* **2018**, *26*, 3613–3625. [CrossRef]

63. Ghatge, S.; Yang, Y.; Ahn, J.H.; Hur, H.G. Biodegradation of Polyethylene: A Brief Review. *Appl. Biol. Chem.* **2020**, *63*, 27. [CrossRef]

64. Esmaeili, A.; Pourbabaee, A.A.; Alikhani, H.A.; Shabani, F.; Esmaeili, E. Biodegradation of Low-Density Polyethylene (LDPE) by Mixed Culture of *Lysinibacillus xylanilyticus* and *Aspergillus niger* in Soil. *PLoS ONE* **2013**, *8*, e71720. [CrossRef] [PubMed]

65. Das, M.P.; Kumar, S. An Approach to Low-Density Polyethylene Biodegradation by *Bacillus amyloliquefaciens*. *3 Biotech* **2015**, *5*, 81–86. [CrossRef]

66. Hasan, F.; Shah, A.A.; Hameed, A.; Ahmed, S. Synergistic Effect of Photo- and Chemical Treatment on the Rate of Biodegradation of Low Density Polyethylene by *Fusarium* sp. AF4. *J. Appl. Polym. Sci.* **2007**, *105*, 1466–1470. [CrossRef]
67. Das, M.P.; Kumar, S.; Das, J. Fungal-Mediated Deterioration and Biodegradation Study of Low-Density Polyethylene (LDPE) Isolated from Municipal Dump Yard in Chennai, India. *Energy Ecol. Environ.* **2018**, *3*, 229–236. [CrossRef]

68. Ratanakamnuan, U.; Aht-Ong, D. Photobiodegradation of Low-Density Polyethylene/Banana Starch Films. *J. Appl. Polym. Sci.* **2006**, *100*, 2725–2736. [CrossRef]

69. Musuc, A.M.; Badea-Doni, M.; Jecu, L.; Rusu, A.; Popa, V.T. FTIR, XRD, and DSC Analysis of the Rosemary Extract Effect on Polyethylene Structure and Biodegradability. *J. Therm. Anal. Calorim.* **2013**, *114*, 169–177. [CrossRef]

70. Morancho, J.M.; Ramis, X.; Fernández, X.; Cadenato, A.; Salla, J.M.; Vallés, A.; Contat, L.; Ribes, A. Calorimetric and Thermo-gravimetric Studies of UV-Irradiated Polypropylene/Starch-Based Materials Aged in Soil. *Polym. Degrad. Stab.* **2006**, *91*, 44–51. [CrossRef]

71. Manzur, A.; Limón-González, M.; Favela-Torres, E. Biodegradation of Physicochemically Treated LDPE by a Consortium of Filamentous Fungi. *J. Appl. Polym. Sci.* **2004**, *92*, 265–271. [CrossRef]

72. Shogren, R.L.; Thompson, A.R.; Felker, F.C.; Harry-Ókuru, R.E.; Gordon, S.H.; Greene, R.V.; Gould, J.M. Polymer Compatibility and Biodegradation of Starch-Poly(Ethylene-co-acrylic Acid)-Polyethylene Blends. *J. Appl. Polym. Sci.* **1992**, *44*, 1971–1978. [CrossRef]

73. Manzur, A.; Cuamatzi, F.; Favela, E. Effect of the Growth of *Phanerochaete chrysosporium* in a Blend of Low Density Polyethylene and Sugar Cane Bagasse. *J. Appl. Polym. Sci.* **1997**, *66*, 105–111. [CrossRef]

74. Weiland, M.; Daro, A.; David, C. Biodegradation of Thermally Oxidized Polyethylene. *Polym. Degrad. Stab.* **1995**, *48*, 275–289. [CrossRef]

75. Cai, Z.; Li, M.; Zhu, Z.; Wang, X.; Huang, Y.; Li, T.; Gong, H.; Yan, M. Biological Degradation of Plastics and Microplastics: A Recent Perspective on Associated Mechanisms and Influencing Factors. *Microorganisms* **2023**, *11*, 1661. [CrossRef] [PubMed]

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