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**Conflict of Interest:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2021 Haque, Mosharaf, Haque, Tanvir and Alam. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.
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# Description of Microbial Communities of Phosphate Mine Wastes in Morocco, a Semi-Arid Climate, Using High-Throughput Sequencing and Functional Prediction

Najoua Mghazli¹,², Laila Sbabou¹, Rachid Hakkou³,⁴, Ahmed Ouhammou⁵, Mariam El Adnani⁶ and Odile Bruneel²*

¹Center of Research Plants and Microbial Biotechnologies, Biodiversity and Environment, Team of Microbiology and Molecular Biology, Faculty of Sciences, Mohammed V University in Rabat, Rabat, Morocco, ²HydroSciences Montpellier, University of Montpellier, CNRS, IRD, Montpellier, France, ³IMED_Laboratory, Faculty of Science and Technology, Cadi Ayyad University (UCA), Marrakech, Morocco, ⁴Mining Environment and Circular Economy Program, Mohammed VI Polytechnic University (UM6P), Benguerir, Morocco, ⁵Laboratory of Microbial Biotechnologies, Agrosciences and Environment (BioMAgE), Team of Agrosciences, PhytoBiodiversity and Environment, Regional Herbarium 'MARK', Faculty of Sciences Semlalia, Cadi Ayyad University, Marrakech, Morocco, ⁶Resources Valorisation, Environment and Sustainable Development Laboratory, National School of Mines of Rabat, Mohammed V University in Rabat, Rabat, Morocco

OPEN ACCESS

**Edited by:**
Aurélie Cebron,
UMR 7360 Laboratoire
Interdisciplinaire des Environnements
Continentaux (LIEC), France

**Reviewed by:**
Pinaki Sar,
Indian Institute of Technology
Kharagpur, India
Rajeev Kaushik,
Indian Council of Agricultural
Research, India

*Correspondence:
Odile Bruneel
odile.bruneel@ird.fr

**Specialty section:**
This article was submitted to
Microbiotechnology,
a section of the journal
Frontiers in Microbiology

**Received:** 11 February 2021
**Accepted:** 07 June 2021
**Published:** 08 July 2021

**Citation:**
Mghazli N, Sbabou L, Hakkou R, Ouhammou A, El Adnani M and Bruneel O (2021) Description of Microbial Communities of Phosphate Mine Wastes in Morocco, a Semi-Arid Climate, Using High-Throughput Sequencing and Functional Prediction. *Front. Microbiol.* 12:666936.
doi: 10.3389/fmicb.2021.666936

Soil microbiota are vital for successful revegetation, as they play a critical role in nutrient cycles, soil functions, and plant growth and health. A rehabilitation scenario of the abandoned Kettara mine (Morocco) includes covering acidic tailings with alkaline phosphate mine wastes to limit water infiltration and hence acid mine drainage. Revegetation of phosphate wastes is the final step to this rehabilitation plan. However, revegetation is hard on this type of waste in semi-arid areas and only a few plants managed to grow naturally after 5 years on the store-and-release cover. As we know that belowground biodiversity is a key component for aboveground functioning, we sought to know if any structural problem in phosphate waste communities could explain the almost absence of plants. To test this hypothesis, bacterial and archaeal communities present in these wastes were assessed by 16S rRNA metabarcoding. Exploration of taxonomic composition revealed a quite diversified community assigned to 19 Bacterial and two Archaeal phyla, similar to other studies, that do not appear to raise any particular issues of structural problems. The dominant sequences belonged to *Proteobacteria*, *Bacteroidetes*, *Actinobacteria*, and *Gemmatimonadetes* and to the genera *Massilia*, *Sphingomonas*, and *Adhaeribacter*. LEfSe analysis identified 19 key genera, and metagenomic functional prediction revealed a broader phylogenetic range of taxa than expected, with all identified genera possessing at least one plant growth-promoting trait. Around 47% of the sequences were also related to genera possessing strains that facilitate plant development under biotic and environmental stress conditions, such as drought and heat.

**Keywords:** phosphate mine wastes, microbial communities, PICRUSt prediction, metabarcoding, biodiversity
# INTRODUCTION

Mining is vital for the global economy, but the extraction of valuable elements also produces large quantities of solid wastes corresponding to uneconomic materials such as waste rock, refuse material, tailings, sediment, roasted ore, and processing chemicals (Hudson-Edwards, 2016).

The mining sector, and particularly phosphate exploitation, is one of the pillars of the Moroccan economy. Among many closed mines in the country, Kettara mine located in a semi-arid region that was exploited for pyrrhotite, is particularly problematic (Hakkou et al., 2008a,b). Mining produced more than 5.2 million metric tons of pyrrhotite concentrate, producing acid mine drainage following rainfall events leading to groundwater contamination (Lghoul et al., 2014). Dominant winds also transport toxic dust and sulfur emissions, from the mine wastes to the village of Kettara (2,000 inhabitants, located in the immediate vicinity of the mine), polluting homes, cattle, and farmland, putting the population under significant environmental and health risks (Khalil et al., 2013; Babi et al., 2016). Morocco holds around three-quarters of the world's phosphate reserves and is the leading exporter of phosphate and its derivatives (Edixhoven et al., 2013; Hakkou et al., 2016). According to a United States Geological Survey<fnref n="1" /> in 2019, Morocco produced around 36 million tons of phosphate rocks representing an economical currency inflow. However, open-pit mining and beneficiation processes including crushing, screening, washing, flotation, or chemical attack generate million tons of tailings called phosphate sludge every year, which is stockpiled over a huge area (Hakkou et al., 2016; Jalali et al., 2019; Elfadil et al., 2020; Trifi et al., 2020). In semi-arid areas, the poor quality of these phosphate mine wastes, containing small quantities of nutrients and contaminated by metals and fluoride, prevents good plant growth (Cross and Lambers, 2017; Jalali et al., 2019; Elfadil et al., 2020). In Morocco, huge piles of phosphate waste rock and tailing ponds that cover thousands of hectares, with very limited vegetation and subjected to wind and water erosion, not only spoil the landscape but are also a serious source of pollution (Hakkou et al., 2016). Revegetation of phosphate mine wastes has therefore become a national priority for Morocco.

The rehabilitation scheme for Kettara mine consists of depositing a layer of alkaline phosphate waste on top of the acidic tailings to limit infiltration of water and to control acid mine drainage generation (Ouakibi et al., 2013; Bossé et al., 2015). In semi-arid climates characterized by low annual precipitation and high evaporation rates, store-and-release covers can prevent percolation of water during wet periods and allow evaporation (or evapotranspiration) during dry periods thus preventing water from penetrating acidic wastes (Bossé et al., 2013). Lab tests (using columns) and field tests (with experimental cells constructed on-site) showed very good results, proving that water infiltration can be controlled, even during extreme rainfall events (Ouakibi et al., 2014; Bossé et al., 2015). The final step in this scheme is the revegetation of the phosphate cover to limit the

dispersion of fine dust, but revegetation is highly challenging for naturally growing biota.

Microbial communities are essential for soil functioning and for the development and maintenance of plant diversity (Hassani et al., 2018; van Bruggen et al., 2019). Soil microorganisms perform essential roles in the development and maintenance of healthy soil structure, including the decomposition of biomass (cellulose degradation, etc.), the transformation or biomineralization of biogenic element, increasing the quantity and availability of nutrients, degrading or biotransforming pollutants, mobilizing/immobilizing metals, etc. (Schulz et al., 2013; Thavamani et al., 2017). Microorganisms can also influence the development of plants *via* nitrogen fixation, phosphorus solubilization, sequestration of iron by siderophores or phytohormones production, etc. (Goswami et al., 2016; Kong and Glick, 2017; Olanrewaju et al., 2017). They can also limit biotic (pathogenic) stresses and abiotic environmental stresses for plants, including drought, salinity, heat, or toxic metals, which are particularly common in alkaline mine tailings in semi-arid areas (Armada et al., 2014; Fahad et al., 2015; Kumar and Verma, 2018; Gupta and Pandey, 2019). Studies dedicated to plant-microorganism associations are currently a very active area of research, and the interest of using microbial communities to stimulate plant growth in damaged and polluted soils, in semi-arid areas has been demonstrated in many countries (Mendez and Maier, 2008; Nirola et al., 2016; Bruneel et al., 2019). Different methods are currently used to inoculate plants with microorganisms. Ju et al. (2020) demonstrated that plant root inoculation with plant growth-promoting (PGP) rhizobia increased the phytostabilization by improving alfalfa growth by decreasing Cu accumulation in shoots in a Cu-contaminated site. Seeds can also be soaked in a bacterial inoculum before germination, as shown by Ke et al. (2021) where PGP bacteria enhanced ryegrass growth and phytostabilization in Cu-Cd-polluted soil. Indeed, pot experiment revealed that PGP bacteria addition increased shoot and root biomasses and limited stress of ryegrass by decreasing metal uptake and translocation. Direct application of bacterial inocula in contaminated soils has also shown positive results for the phytoextraction of excess phosphorus in soils (Ye et al., 2020; Guo et al., 2021). Guo et al. (2021) showed, for example, in pot experiments that two native phosphate-solubilizing microorganisms increased phosphate availability by reducing soil pH. Inoculation of these strains in soil also significantly increased the total biomass of two native plants, like their root and shoot length, and chlorophyll content.

As only a few plants managed to grow naturally after 5 years on the store-and-release cover, we sought to know if any structural problem in phosphate waste communities could explain the almost absence of plants. The aims of the present work were thus to (i) investigate the structure of the microbial communities (using 16S rRNA metabarcoding approach) and (ii) identify their potential metabolic functions in phosphate mine wastes using PICRUSt2 prediction, to test this hypothesis. To our knowledge, this is the first study to describe the microbial communities present in alkaline phosphate mine wastes in Morocco using high-throughput sequencing and the first study

<footnote n="1">¹http://minerals.usgs.gov/minerals/pubs/commodity/phosphate_rock/</footnote>
on overburden phosphate waste rocks and phosphate sludge issuing from processing. This phylogenetic classification of microbial communities of phosphate mine wastes and *in silico* identification of their potential plant growth-promoting diversity will provide useful information to prepare media dedicated to the isolation of the most interesting plant growth-promoting microorganisms identified in this study, helping revegetation in a semi-arid climate.

# MATERIALS AND METHODS

## Description of Study Site and Experimental Cells

To evaluate the feasibility and effectiveness of using alkaline phosphate mine wastes as a store-and-release layer to limit infiltration of water in acidogenic tailing mine, field investigations were conducted at Kettara mine (31°52'15"N-8°10'31"W). Instrumented experimental cells (10 m × 10 m), with different configurations of store-and-release layer, and comprising two types of mine wastes, phosphate waste rock (overburden waste, removed to reach the phosphate ore) and phosphate sludge (produced during the concentration processes) were constructed in 2011 (Bossé et al., 2015; Table 1 and

**Supplementary Figure 1**). The final step in the scheme of rehabilitation is the revegetation of the phosphate cover to limit the dispersion of fine dust, but after 5 years, only a few plants naturally grew on the experimental cells. Like belowground biodiversity is a key component for aboveground functioning, we conducted an *in situ* investigation, to identify bacterial and archaeal communities present in these experimental cells. To see if the presence of some annual plants, naturally growing, modified the structure of microbial communities, the microbial communities in soils sampled close to plants and in bare soils were also compared for each cell. Due to lack of plants naturally growing on these cells, three different species were chosen, *Spergularia rubra* (L.) for cell 1, *Atriplex semibaccata* for cell 3, and *Echium vulgare* (L.) for cell 4. *Echium vulgare* (L.) is a metallicolous plant that grows naturally in mine wastes and highly tolerant to Cd soil contamination (Dresler et al., 2015; Kompala-Baba et al., 2019). *Atriplex semibaccata*, found in abandoned mine, hyperaccumulate Cu and Cd and accumulate Fe (Baycu et al., 2015). *Atriplex* is also highly salt tolerant and has already been used as a pioneer species for revegetation in semi-arid areas (Nirola et al., 2016). *Spergularia rubra* (L.) J.Presl and C.Presl has been found as a pioneer plant in mine tailings (Garcia-Carmona et al., 2019) and is known to accumulate Cu in shoots (Bech et al., 2017).

<table><caption>**TABLE 1** | Description of experiment cells (10 m × 10 m), constructed in 2011 to test their capacities to limit infiltration of water and comprising two types of phosphate wastes used as store and release cover (Bossé et al., 2015).</caption><thead><tr><th>Type of material</th><th>Sample name</th><th>Thickness of cover</th><th>Fraction</th><th>Name of plants sampled</th><th>Sample code</th><th>Main composition*</th><th>Electrical conductivity**</th></tr></thead><tbody><tr><td rowspan="10">Phosphate waste rock, overburden wastes, removed to reach the phosphate ore</td><td rowspan="3">Cell 1</td><td rowspan="3">100 cm</td><td rowspan="3">Raw material</td><td rowspan="3">Spergularia rubra (L.)</td><td>C1VS1</td><td>Calcite (40.7 wt%)</td><td rowspan="3">147 to 496 µS cm<sup>-1</sup></td></tr><tr><td>C1VS2</td><td>Apatite (25.9 wt%)</td></tr><tr><td>C1VS3</td><td>Dolomite (25.2 wt%)<br>Quartz (8.2 wt%)<br>Chromium (Cr,<br>137 mg/kg)<br>Strontium (Sr,<br>769 mg/kg)<br>Fluor (F, 1.3 wt%)</td></tr><tr><td rowspan="3">Cell 4</td><td rowspan="3">100 cm</td><td rowspan="3">&lt; 1 mm</td><td rowspan="3">Echium vulgare (L.)</td><td>Bare soil</td><td>C1BS1<br>C1BS2<br>C1BS3</td><td></td></tr><tr><td>Bare soil</td><td>C4VS1<br>C4VS2<br>C4VS3</td><td></td></tr><tr><td>Bare soil</td><td>C4BS1<br>C4BS2<br>C4BS3</td><td></td></tr><tr><td rowspan="6">Phosphate sludge, produced during the concentration processes</td><td rowspan="3">Cell 3</td><td rowspan="3">100 cm</td><td rowspan="3">Raw material</td><td rowspan="3">Atriplex semibaccata</td><td>C3VS1</td><td>Fluorapatite (44 wt%)</td><td rowspan="3">178.6 to 691 µS cm<sup>-1</sup></td></tr><tr><td>C3VS2</td><td>Quartz (17 wt%)</td></tr><tr><td>C3VS3</td><td>Calcite (15 wt%)<br>Dolomite (7 wt%)<br>Cadmium (Cd,<br>131 mg/kg)<br>Chromium (Cr,<br>275 mg/kg)<br>Strontium (Sr,<br>927 mg/kg)<br>Fluor (F, 2.6 wt%).</td></tr><tr><td rowspan="3">Cell 4</td><td rowspan="3">100 cm</td><td rowspan="3">&lt; 1 mm</td><td rowspan="3">Echium vulgare (L.)</td><td>Bare soil</td><td>C3BS1<br>C3BS2<br>C3BS3</td><td></td></tr><tr><td>Bare soil</td><td>C3BS1<br>C3BS2<br>C3BS3</td><td></td></tr><tr><td>Bare soil</td><td>C3BS1<br>C3BS2<br>C3BS3</td><td></td></tr></tbody></table>

<footnote>*According to Hakkou et al., 2009, 2016.</footnote>

<footnote>**Personal communication from Rachid Hakkou.</footnote>
## Sample Collection

In this study, three experimental cells already present were investigated (cells 1, 3, and 4, Bossé et al., 2015). For each cell, soil samples were collected in triplicate at the surface (0 to 5 cm) close to the roots of each plant (some centimeters from the roots) or in bulk soils located a few meters away from the plants to avoid the influence of plants. To obtain more homogenous representative samples, each of the three topsoil replicates was formed by mixing five soils, collected in 1 m<fnref n="2" /> subsamples of bare soil and in close proximity of the roots of five annual plants. These five samples were thoroughly homogenized to form three composite samples. Samples were named as follows: C1BS and C1VS for cell 1 with "BS" for bare soils and "VS" for soils with plants and with 1, 2, and 3 as replicates; C3BS and C3VS for cell 3, and C4BS and C4VS for cell 4. All 18 samples were immediately placed in Falcon tubes and stored in the field in coolers containing blocks of ice and transported to laboratory where they were stored at -20°C until DNA extraction. One tube from each site was stored at 4°C for physicochemical analyses.

## Physicochemical Characterization

The chemical composition of the samples was determined using X-ray fluorescence (XRF, Epsilon 4 Model, Malvern Panalytical). The device contains a kV tube with a silver (Ag) anode that produces X-rays. Its detector energy resolution was 4,000–50,000 kV. The samples were ground to a powder to obtain a particle size close to that of standards. The quality of the data was assessed using duplicate sample analyses, and measurement accuracy was estimated at ± 5% for all the elements analyzed. The total organic carbon was determined by potassium bichromate digester (Behrotest Heiz block K-40). The total nitrogen was measured using sulfuric acid-salicylic acid digestion following Kjeldahl method adjusted (Bremner, 1960) and determined by Kjeldahl K-375 BUCHI Kjel Master analyzer. The pH was measured by WTW/INOLAB pH7310 following the pH-meter instructions. Hundred milliliters of soil suspension was prepared in distilled water with a ratio of 1:5, then pH was measured using 1 mol/L of KCl.

## DNA Extraction and Amplification Using Illumina MiSeq Sequencing

Soil DNA was extracted in triplicates using the DNeasy® PowerMax® Soil kit Qiagen following the manufacturer's instructions. All extracted genomic DNA samples were stored at -20°C until further analysis. Amplicon libraries were constructed according to ADNid<fnref n="2" /> dual indexing strategy. The universal primer set 519F (S-D-Arch-0519-a-S-15) (5'-CAGCMGCCGCGGTAA-3') and 805R (S-D-Bact-0785-a-A-21) (5'-GACTACHVGGGTATCTAATCC-3') designed to target bacterial and archaeal taxa (Klindworth et al., 2013) was used to amplify a 286-bp region targeting the V4 region of 16S rRNA genes. Amplicons were generated using Hot start DNA polymerase (QIAGEN). First, PCR (PCR1) was performed on 10 ng of template DNA in 15 µl total mix, submitted to an initial

denaturation of 5 min at 95°C, followed by 35 cycles of 30 s at 95°C, 1 min at 60°C, and 30 s at 72°C. A final extension of 30 min at 60°C completed the reaction. Amplicons were checked on 1.5% agarose gel and purified using AMPure beads. A second PCR step (PCR2) was performed using a specific Illumina adapter, the index and the Illumina overhang adapter primers in 2 × KAPA Hifi Hotstart Ready Mix (Roche, Switzerland). A previously prepared 5-µl volume of PCR products was used to obtain a final volume of PCR mix of 50 µl. A 3-min denaturation step at 95°C was performed followed by 12 cycles of 30 s at 95°C, 30 s at 55°C, 30 s at 72°C, and a final 5-min extension step at 72°C. PCR2 products were purified (AMPure Beads), quantified (Nanodrop, Spark), normalized at 15 ng/µl, and pooled in a final library. The library was then purified using AMPure Beads, and its quality was checked using a fragment analyzer. Final quantification was performed by qPCR (KAPA library quantification Kits, Roche, Switzerland). Sequencing runs, generating 2 bp × 250 bp, reads were performed on an Illumina MiSeq using V2 chemistry.

## Sequence Analysis of Metabarcoding Datasets

The sequencing data analysis was conducted using FROGS v3.1.0 (Find Rapidly OTUs with Galaxy Solution, Escudié et al., 2018). Briefly, paired reads were merged using FLASH (v1.2.11, Magoc and Salzberg, 2011) with a minimum length of 240 bp and an overlap length of 270 bp. After denoising and removal of primer/adapters with cutadapt (v1.18, Martin, 2011), de novo clustering was implemented with SWARM (v2.2.2), which uses a local clustering threshold (Mahé et al., 2015), with an aggregation distance of *d* = 3. Chimeric sequences were removed using VSEARCH (v2.91, Rognes et al., 2016). The dataset was filtered, and sequences present in less than two samples and sequences representing less than 0.005% of the total set of sequences were discarded, as recommended by Bokulich et al. (2013). The taxonomic affiliation of operational taxonomic units (OTUs) was performed using Blastn+ against the Silva database (release 132, December 2017) for 16S rRNA gene amplicons using an innovative multi-affiliation output to identify conflicting sequences in the database and uncertainties (Escudié et al., 2018).

## Statistical Analysis and Predictive Metagenome Analysis

All statistical analyses were performed using R software<fnref n="3" /> unless otherwise stated. The alpha diversity statistics were calculated for each sample, and the rarefaction curves and the diversity indexes (Chao1, Shannon, and Simpson) were generated. The variation of beta diversity was graphically illustrated using non-metric multidimensional scaling (nMDS) analysis. Environmental variables were vector fitted to the nMDS ordination plots using the function "envfit" in the R "vegan" package to assess possible explanatory variables. Analysis of variance (ANOVA) was used at 95% confidence level and was completed by Tukey's post hoc test to analyze significant distribution of the chemical parameters. Pearson

<footnote n="2">²www.qualtech-groupe.com, Montpellier, France</footnote>

<footnote n="3">³v3.6.0, http://www.r-project.org/</footnote>
correlations were used to assess the relationships between environmental factors, and the microbial community diversity. A linear discriminant analysis effect size (LEfSe) method (Segata et al., 2011) was also used to identify statistically significant microbial indicators associated with each cell and to identify the most discriminating clades. This analysis was done through the online interface at https://huttenhower.sph.harvard.edu/galaxy. Network analysis was used to reveal the relationship between genera present in bare soils and those present in soils sampled near plants. The preparation of data for network analysis was carried out using "psych" package as described by Batushansky et al. (2016). The "r coefficient" and "p-value" tables were generated, and only significant interactions were selected and visualized using Cytoscape V3.8.0 with the organic layout. The nodes represent the genera that differed significantly between bare soil and soil near the plant roots (p < 0.05) based on the results of ANOVA, and the edges represent the correlation between these genera.

Finally, PICRUSt2 (phylogenetic investigation of communities by reconstruction of unobserved states, Douglas et al., 2020) analysis was used to explore the possible functional profiles of the microbial communities. This was done following the pipeline command «picrust2_pipevine.py» using the "-stratified" flag as described in the PICRUSt2 GitHub<fnref n="4" />. The weighted Nearest Sequences Taxon Index (NSTI) was calculated to assess the accuracy of the PICRUSt2 analysis (Langille et al., 2013). The EC table resulting from PICRUSt2 was filtered, and only enzymes that contribute to pathways of interest in the present study were retained, i.e., phosphate solubilization, nitrogen metabolism, thiosulfate oxidation, cellulase degradation, and indole 3-acetic acid (IAA) and siderophore production. Phosphate solubilization in soil can be catalyzed by bacteria that produce enzymes including phosphatases, phosphonatases, phytases, and C-P lyase (Billah et al., 2019; Prabhu et al., 2019). For all other metabolisms, our study was based on the pathways published by the MetaCyc database<fnref n="5" />. To determine the presence of a function in one of the genera detected, all the enzymes involved in the given function's pathway must be present in the same genus. The number of enzymes/functions was converted to "1" if present and to "0" if absent, and a presence/absence heatmap was then constructed using the "ggplot2" package.

# RESULTS

## Physicochemical Characteristics of Phosphate Mine Wastes in Morocco

The environmental characteristics of soil samples in the three cells are presented in **Figure 1**. ANOVA was used to test differences in geochemical factors among the samples and was completed with Tukey's *post hoc* test at a confidence interval of 95%. Phosphate mining wastes were highly alkaline (pH 8–9) with low nutrient contents [total organic carbon (TOC) between 0.3 and 0.4%, and total nitrogen (N) < 0.034%] but very

high level of P₂O₅ (> 75 g kg⁻¹). Significant differences were observed between five parameters, pH, SiO₂, P₂O₅, Fe₂O₃, and to a lesser extent in MgO between the two types of phosphate mine wastes, phosphate waste rocks (cells 1 and 4), and phosphate sludge (cell 3). pH, SiO₂, P₂O₅, and MgO concentrations were statistically higher in phosphate sludge than in waste rocks. Inversely, concentrations of Fe₂O₃ were higher in waste rocks. No clear differences were found in many elements, including TOC, N, or CaO, between the three cells thus between the two types of phosphate wastes. No significant differences were found inside cells, between soils sampled near plants, and bare soils, except for TOC and N in cell 3, where the highest percentage was found in C3VS. No differences in K₂O, TiO₂, and Al₂O₃ were observed between samples.

## Diversity and Taxonomic Assignment of Microbial Communities in Phosphate Wastes

A total of 8,507,878 sequences paired-end reads were obtained from the samples collected in 18 phosphate mine wastes. After cleaning and processing, 7,633,309 high-quality sequences were obtained with an average of 428,414 sequences per sample. The number of sequences per sample was then made equal by random resampling (191,792 sequences per sample, based on the smaller number of sequences) giving 3,452,256 good-quality reads. At 97% similarity, all effective normalized sequences were distributed among 869 microbial OTUs (**Supplementary Table 1**). Rarefaction curves achieved an asymptote for all samples, suggesting that almost all OTUs were detected (**Supplementary Figure 2**). The number of OTUs estimated by Chao1 was also very similar to the richness (observed number of OTUs), ranging, respectively, from 650 to 800 for Chao1 and from 640 to 800 for richness, confirming that virtually all the microbial diversity was explored (**Supplementary Figure 3**). No statistical differences were observed between different cells but statistical differences were found between bare soils (C1BS, C3BS, and C4BS) and soils located near plants (C1VS, C3VS, and C4VS), with systematically higher richness and Chao1 in soils near plants than in bare soils. No statistical differences in Shannon and Simpson indices were found in the three cells or between bare soils and soils near plants, except for C4BS, which had lower Shannon and Simpson values.

Exploration of taxonomic composition revealed that the 869 bacterial and archaeal OTUs were assigned to 19 phyla with only two Archaea (this kingdom represented only 0.11% of the total sequences, **Figure 2A**). Over 98% of the OTUs were assigned to a taxonomic phylum with 80% confidence. Across all 18 samples, *Proteobacteria* (51.3%), *Bacteroidetes* (22%), *Actinobacteria* (9.3%), and *Gemmatimonadetes* (7.9%) were the dominant phyla represented by more than 90% of all sequences. For Bacteria, at the genus level, 59.7% of sequences were attributed to known genera, 22.1% to unknown genera, and 18.2% were multi-affiliated (**Supplementary Table 2**). The majority of sequences were affiliated to the genera *Massilia* (9.4%), *Sphingomonas* (5.7%), *Adhaeribacter* (3.9%), and *Hymenobacter* (2.7%, **Figure 2B** and **Supplementary Table 2**). Only 18 genera

<footnote n="4">⁴https://github.com/picrust/picrust2/wiki/Full-pipeline-script</footnote>

<footnote n="5">⁵https://biocyc.org/</footnote>
<figure><img src="image_95.png" /><figcaption>**FIGURE 1** | Boxplot displaying the concentrations of different chemical parameters. Organic carbon (OC) and nitrogen (N) are presented by percent. Major oxides such as silica (SiO₂), phosphorus pentoxide (P₂O₅), ferric oxide (Fe₂O₃), lime (CaO), magnesia (MgO), potash (K₂O), titanium oxide (TiO₂), and alumina (Al₂O₃) are in milligrams per kilogram. The letters present the compact display letters (CDL). Error bars represent standard deviation of the mean value (analyses performed in triplicates). Values in each column followed by the same letter are not significantly different at 0.05% (Tukey's).</figcaption></figure>

had more than 1% of total sequences. For Archaea, up to 82.7% of sequences were affiliated to *Euryarchaeota* phylum and to unknown genera, while the remaining 17.3% were affiliated to *Thaumarchaeota* phylum, among which 70.8% were multi-affiliated to different genera of the *Nitrososphaeraceae* family and 29.2% were affiliated to the genus *Nitrososphaera*.

## Structure of Microbial Communities in Phosphate Mining Wastes and the Effect of Plants

nMDS ordination and PERMANOVA was performed to explore the relationships between the structure of the microbial communities and the physicochemical characteristics of the phosphate mining wastes (Figure 3 and Supplementary Table 3). Phosphate sludge (six replicates of cell 3) and waste rocks (12 replicates of cell 1 and cell 4) were clearly separated along the first ordination axis. The second axis separated samples into two distinct groups, bare soils in all three

cells and those of soils sampled near plants. This result was confirmed by statistical analysis using Jaccard dissimilarities (PERMANOVA, *p*-value = 0.002), indicating that community composition differed significantly between the two types of mine wastes and between soils near plants and bare soils (Supplementary Table 3). Vector fitting of environmental parameters on nMDS revealed that P₂O₅, pH, SiO₂, and MgO were positively correlated with cell 3 and separated the two types of phosphate mining wastes (Figure 3 and Supplementary Table 4). Fe₂O₃ and, to a lesser degree, CaO, were negatively correlated with the first four parameters and were more abundant in cell 1 and cell 4. Vector fitting also revealed that TOC and, at a lesser degree, N, were positively correlated with soils near plants.

Among the 130 identified genera, Pearson analysis indicated that 79 were significantly correlated with environmental variables (Figure 4). Many correlations were observed with SiO₂ (35), P₂O₅ (31), pH (31), Fe₂O₃ (30), MgO (21), TOC (17), CaO (18), and N (9) but only one with Al₂O₃ and TiO₂. For
<figure><img src="image_96.png" /><figcaption>**FIGURE 2** | Microbial diversity present in the alkaline phosphate mining wastes. At the phylum level (A), all sequences are presented, while at the genus level (B), only the most abundant sequences are presented.</figcaption></figure>

SiO₂, P₂O₅, pH, Fe₂O₃, MgO, and TOC, 20, 18, 14, 20, 11, and 14 genera were, respectively, positively correlated, whereas 15, 13, 17, 10, 10, and 3 were negatively correlated. All 18 correlations found for CaO were positive, and all nine correlations found for N were negative. Very similar correlation indices were also observed between P₂O₅, pH, SiO₂, and, to a lesser extent, MgO, with relatively dominant genera such as *Adhaeribacter*, *Hymenobacter*, and *Cnuella* positively affected, while *Brevundimonas*, *Arctibacter*, and *Devosia* were strongly negatively affected. The LEfSe analysis identified 9, 20, and 11 bacterial taxa, containing 3, 10, and 6 genera that statistically distinguished, respectively, cell 1, cell 3, and cell 4 (with LDA score > 2, **Figure 5**). The family *Hymenobacteraceae* was the most differentially abundant taxon with a LDA score > 4, reflecting higher abundance in cell 3 than in cells 1 and 4. At the genus level, *Adhaeribacter*, *Cnuella*, *Blastocatella*, *Tychonema*, *Bdellovibrio*, *Segetibacter*, *Rubritepedia*, *Deinococcus*, *Pajaroellobacter*, and *Nostoc* were overrepresented in cell 3. In cell 1, only *Marmoricola*, *Bacillus*, and *Myxococcus* were prevalent, while in cell 4, *Skermanella*, *Sphingoaurantiacus*, *Ohtaekwangia*, *Geodermatophilus*, *Solirubrobacter*, and *Pedomicrobium* were more abundant. These 19 genera were also highlighted by Pearson's correlation. A correlogram of these core genera separated them into two distinct groups (**Figure 6**). The first group contained all genera found in the phosphate sludge (cell 3) identified by LEfSe and presented quite strong positive

correlations among themselves and negative correlations among genera sampled in phosphate waste rocks (except for *Bdellovibrio*, which was positively correlated with *Ohtaekwangia*). The second group comprised genera retrieved in phosphate waste rocks (cells 1 and 4), and correlations between members of this group were generally positive, while correlations with phosphate sludge were negative.

Network analysis was also used to identify the relationships between genera found in bare soils and in soils sampled near annual plants (**Supplementary Figure 4**). Fifteen genera were found to be significantly associated with bare soils and 16 were shown to be more abundant in soils near plants.

## Potential Plant Growth Promotion Traits of Microbial Communities of Phosphate Wastes

PICRUST2 analysis was used to explore the possible metabolic pathways associated with microbial communities in phosphate wastes (**Figure 7**). The NSTI values < 0.06, confirmed the prediction accuracy (**Supplementary Table 5**). All 130 OTUs affiliated to known genera were predicted to possess at least one of the interesting enzymes involved in important traits related to plant development. For the phosphate solubilization, 102 genera were predicted to possess the alkaline phosphatase enzyme while only 39 were predicted to possess phytase 1, 23 acid phosphatase,
<figure><img src="image_97.png" /><figcaption>**FIGURE 3** | Non-metric multidimensional scaling (NMDS) presenting the differences of the microbial communities between the different replicates of phosphate mining waste samples. The stress values were < 0.05, indicating that these data were very well represented by the two-dimensional representation. Arrows are the projections of possible explanation variables obtained by vector fitting. Only correlations with a false discovery rate (fdr) corrected *p* < 0.05 were indicated. The angle and length of the vector indicate the direction and strength of the variable. The *r*² correlation coefficient and the *p*-values are presented in **Supplementary Table 3**.</figcaption></figure>

18 C-P lyase, 8 phosphonatase, and only 4 phytase 2. The OTUs assigned to *Novosphingobium* appeared to have all the phosphate-solubilizing enzymes except C-P lyase. For nitrogen metabolism, all genera were predicted to possess at least one of the seven enzymes involved in N assimilation and referenced in PICRUSt2 and also to produce nitrate-reducing enzymes. Only 2 genera were predicted to be able to oxidize ammonia (*Nitrosomonas* and the archaea *Nitrososphaera*), 59 were predicted to reduce nitrite, and 10 to fix nitrogen. For phytohormones production, 96 genera were predicted to possess at least one enzyme involved in different auxin biosynthesis pathways. Twenty-six genera were predicted to possess the thiosulfate oxidation enzyme and only 14 to biosynthesize aerobactin, an enzyme involved in siderophore production.

DISCUSSION

Characteristics of Phosphate Mining
Wastes and Development of Plants

Many mine tailings disposed in artificial dumps have not
been subject to the long-term natural soil formation processes,
representing challenging conditions for biota (Cross and
Lambers, 2017). Mine wastes containing generally ultrafine
particles, present also often dysfunctional and unstable physical
and geochemical structures, altered or inadequate hydrological
functioning, associated generally with extreme pH, limited

concentrations of organic material and nutrients, and often
presenting an accumulation of toxic or radioactive substances
(Cross and Lambers, 2017; Cross et al., 2017). The phosphate
mine wastes in the present study share many of these
characteristics with highly alkaline soils (pH 8–9), very low
levels of nutrients (TOC and N), very high concentrations of
P₂O₅ (mass percent between 10% and 20%) and associated
with high levels of hazardous contaminants (e.g., Sr, Cr, Cd,
or F, Hakkou et al., 2016). Phosphate mine wastes mainly
comprise ultrafine material with significant quantities of calcite
and dolomite (Hakkou et al., 2009). In arid and semi-arid areas,
soils present usually low moisture content and limited organic
matter associated with extreme temperatures and irradiance
(Padmavathiamma et al., 2014). They are also quite often saline,
because of the high evaporation rate and the low water infiltration
(Mendez and Maier, 2008). However, phosphate mine wastes
in this study were not saline according to their electrical
conductivity (Table 1). All these harsh properties could explain
the almost absence of plants.

Structure of Microbial Communities and
Potential Physicochemical Drivers

Characterization of the microbial communities in phosphate
wastes enabled most OTUs to be classified at the phylum
level; whereas, only 55.3% of sequences were classified at
the genus level. The high proportion of unclassified genera
suggests the presence of possible novel microorganisms and is in
<figure><img src="image_98.png" /><figcaption>**FIGURE 4** | Heatmap of the statistically significant coefficients of Pearson's correlations (false discovery rate (fdr) corrected; *p* < 0.05) between the relative abundance of taxa and concentrations of environmental parameters across all sites. Only threshold higher than 0.7 was plotted.</figcaption></figure>

agreement with the results of many studies conducted in mining environments (Bruneel et al., 2017; Liu et al., 2019). Around 90% of sequences belonged to only four phyla *Proteobacteria*, *Bacteroidetes*, *Actinobacteria*, and *Gemmatimonadetes*, and the dominant genera were identified as *Massilia*, *Sphingomonas*, *Adhaeribacter*, *Hymenobacter*, and *Brevundimonas*. Archaea formed a very small minority, and only one genus was identified,

*Nitrososphaera*. Zouch et al. (2017) obtained similar results for phosphogypsum in Tunisia, where Archaea accounted for only 1.5% of the microbial community and no more than 3% in pyrrhotite mine tailings of Kettara (Bruneel et al., 2017). Among the 130 identified genera, only 18 had more than 1% of the total number of sequences. This is in agreement with the results obtained by Trifi et al. (2020), indicating that in
<figure><img src="image_99.png" /></figure>

<figure><img src="image_100.png" /><figcaption>**FIGURE 5** | Linear discriminant analysis effect size (LEfSe), (A) chart and (B) cladogram, on microbial diversity in different store-and-release phosphate mine waste layer, overburden waste rock (cells 1 and 4), and phosphate mine wastes (cell 3) produced during the concentration processes.</figcaption></figure>

Tunisia, phosphogypsum contained only five low abundance genera (between 1 and 5% of sequences) out of the 193 detected.

The composition of microbial communities differed significantly between the two types of mining wastes and was affected by soil chemical properties, particularly soil pH, P₂O₅, Fe₂O₃, MgO, CaO, and SiO₂. Phosphorus and pH have been recognized as drivers of microbial communities, and pH is generally recognized to be the best predictor of microbial composition and diversity (Rousk et al., 2010; Wu et al., 2018; Ye et al., 2020).

Our investigation, combining Pearson correlations and LEfSe, also identified 19 genera divided into two groups, those found in phosphate sludge and those found in phosphate waste rocks. These core genera probably have significant functions in phosphate mining wastes, but, apart from *Bacillus*, *Nostoc*, and, to a lesser extent, *Myxococcus*, many of them are not known as PGP microorganisms such as *Adhaeribacter*, *Cnuella*, *Blastocatella*, *Tychonema*, *Sphingoaurantiacus*, *Segetibacter*, *Rubritepida*, *Deinococcus*, and *Pajaroellobacter* (**Supplementary Table 6**). Other genera possessed only few PGP traits: *Skermanella*, *Geodermatophilus*, and *Marmoricola* are involved in nitrogen cycling, *Ohtaekwangia* sp. displays antimicrobial activity, *Solirubrobacter* sp. is able to degrade cellulose, or *Bdellovibrio* and *Myxococcus*, which are known to be predators of other bacteria.

The composition of the microbial communities differed significantly between bare soils and soils sampled near plants. Richness and estimated OTUs were also higher in wastes associated with plants than in bare soils in all the three cells. Although our study did not analyze the rhizosphere, the proximity of plants appeared to have an impact on the structure of microbial communities, and organic carbon and N were identified as the main drivers. This is consistent with many studies reporting differences in the diversity and taxonomic composition of microbial communities in bulk soils and in the rhizosphere (Edwards et al., 2015; Luo et al., 2018; Orozco-Mosqueda et al., 2020; Ye et al., 2020). Plant species or even genotypes are also known to select divergent microbial communities (Bulgarelli et al., 2013; Edwards et al., 2015; Ye et al., 2020). This is usually explained by the presence of many elements exuded by roots including sugars, amino acids, and organic acids, which can account for one-third of the carbon fixed by plants (Bulgarelli et al., 2013; Edwards et al., 2015; Olanrewaju et al., 2017). Many significantly different genera were observed in soils near plants and in bare soils, but no clear differences were observed concerning their potential PGP traits as symbiotic N₂ fixers, contrary with that reported by Luo et al. (2018). Although in agreement with Ye et al. (2020) in phosphogypsum, the relative abundance of three genera (*Blastocatella*, *Thermithiobacillus*, and *Sphingomonas*) was higher in soils associated with plants.
<figure><img src="image_101.png" /><figcaption>**FIGURE 6** | Correlogram representing the Pearson's correlation coefficient *r*² between the 19 key bacterial genera ordered by the "hclust" method.</figcaption></figure>

Other factors affect the abundance, structure, and activity of microbial communities, including the type, properties, and texture of the soil, as well as climatic conditions (Edwards et al., 2015; Luo et al., 2018; Ye et al., 2020). The structure of the microbial communities analyzed in this study differed from that identified in other studies conducted in phosphate mine wastes, such as in Ragot et al. (2013) using T-RFLP and clone libraries to study apatite, the principal phosphate mineral. Next-generation sequencing are generally used to investigate microbial communities in phosphate mining area in China (Ye et al., 2020) and in phosphogypsum in Portugal (Martins et al., 2016), in China (Wang et al., 2018), or in Tunisia (Zouch et al., 2017; ben Mefteh et al., 2019; Trifi et al., 2020). However, phosphogypsum, the main byproduct of phosphoric acid, obtained after sulfuric acid digestion, present clearly different physicochemical characteristics, compared with phosphate wastes in this study. Nevertheless, our microbial communities were quite similar to those associated with turfgrass seeds of three turf species (Festuca rubra, Lolium arundinaceae, and Lolium perenne) in low moisture climates (annual precipitation < 750 mm), containing a majority of Proteobacteria (89%) followed by Actinobacteria (6%, Chen et al., 2020). At the genus level, Chen et al. (2020) found a high proportion of Massilia (25%), and Pseudomonas (29%), followed by Sphingomonas (6%), Paenibacillus (3%), Pedobacter (1%), Flavobacterium, and Chryseobacterium (< 1%). This structure is also quite similar to the core of phyllosphere

communities composed of *Pseudomonas*, *Sphingomonas*, *Methylobacterium*, *Bacillus*, *Massilia*, *Arthrobacter*, and *Pantoea* (Bulgarelli et al., 2013).

## Potential Ecological Role in Plant Growth-Promoting Traits

Surprisingly, a huge number of microorganisms recognized as potential PGP genera, were found in alkaline phosphate mining wastes according to the PICRUSt2 predictions. Indeed, all 130 OTUs assigned to known genera were predicted to possess at least one enzyme involved in plant growth-promoting traits. These results are quite consistent with the literature and show that a very large proportion of our sequences (66.15%) resemble microorganisms that contain isolated strains known to present one plant growth-promoting trait (Supplementary Table 6). Many sequences were similar to very well-known PGP genera such as *Pseudomonas*, *Bacillus*, *Arthrobacter*, *Azospirillum*, *Herbaspirillum*, *Rhizobium*, *Flavobacterium*, *Variovorax*, *Streptomyces*, *Sphingomonas*, and *Paenibacillus* (Goswami et al., 2016, Supplementary Table 6). Some of these microorganisms are also currently sold in international markets as biofertilizers or biocontrol agents (Bio-Save®, Mycostop®, Galltrol-A, etc.) and are used by farmers in the field as inocula (Goswami et al., 2016; Pathak and Kumar, 2016; Kumari et al., 2019). Many of the most abundant sequences, representing
<figure><img src="image_102.png" /><figcaption>**FIGURE 7** | Binary heatmap highlighting presence (■) absence (■) of the interested enzymes in identified genera. Enzymes implicated in: cellulose degradation by producing the cellulase enzyme (EC:3.2.1.4); sulfur metabolism by oxidizing thiosulfate (EC:1.8.2.2); siderophores production by catalyzing aerobactin biosynthesis pathway (EC:1.14.13.59, EC:2.3.1.102, EC:6.3.2.38 and EC:6.3.2.39); phosphate solubilization by producing either one or multiple of the enzymes C-P lyases (EC:4.7.1.1), phytase 1 (EC:3.1.3.8), phytase 2 (EC:3.1.3.26), alkaline phosphatase (EC:3.1.3.1), acid phosphatase (EC:3.1.3.2), and phosphonatase (EC:3.11.1.1); nitrogen metabolism by catalyzing at least one of the following pathways: nitrogen fixation I (ferredoxin) (EC:1.18.6.1), ammonia assimilation cycle I (EC:6.3.1.2 and EC:1.4.1.14); ammonia assimilation cycle II (EC:6.3.1.2 and EC:1.4.7.1), ammonia assimilation cycle III (EC:6.3.1.2 and EC:1.4.1.13), ammonia oxidation I and IV nitrite producing (EC:1.14.99.39 and EC:1.7.2.6), nitrate reduction VI L-glutamine forming (EC:1.7.7.2, EC:1.7.7.1, and EC:6.3.1.2), and nitrate reduction VI L-glutamate forming (EC:1.7.7.2, EC:1.7.7.1, and EC:1.4.1.4), and finally in auxin biosynthesis by producing enzymes implicated in indole-3-acetate biosynthesis III (EC:1.13.12.3 and EC:3.5.1.4, or in indole-3-acetate biosynthesis IV (EC:4.2.1.84 and EC:3.5.1.4) or in indole-3-acetate biosynthesis V (EC:3.5.5.1).</figcaption></figure>
more than 1% of the total sequences, also possessed many plant growth-promoting traits including *Massilia*, *Brevundimonas*, *Lysobacter*, or *Noviherbaspirillum* or genera belonging to *Hymenobacteraceae* family (identified by LEfSe as very abundant in phosphate sludge) comprising *Adhaeribacter*, *Hymenobacter*, *Pontibacter*, or *Rufibacter*.

According to PICRUSt2 prediction, all the microorganisms possessed at least one of the five enzymes involved in nitrogen cycling and more than 43 genera possessed isolated strains implicated in nitrogen cycling (Supplementary Table 6). Among them, *Rhizobium* is the main contributor to symbiotic nitrogen fixation in legume crops (Pathak and Kumar, 2016), but many free-living nitrogen fixers were also present, including *Azospirillum*, *Herbaspirillum*, *Bacillus*, or *Paenibacillus* (Goswami et al., 2016). *Azospirillum* is thought to fix from 5 to 20 kg/ha/year of N₂, corresponding between 20 and 25% of total nitrogen required by maize or rice (Pathak and Kumar, 2016). For Archaea, *Nitrososphaera* is known to be an ammonia-oxidizing archaeon (Stieglmeier et al., 2014) and could thus contribute to the global nitrogen cycle in soils via nitrification (Hatzenpichler, 2012). According to metagenome functional prediction, 95% of the genera identified possess enzymes involved in phosphate solubilization and 44 of our sequences resembled genera containing isolated strains with phosphate solubilization capacity and comprising very well-known phosphate solubilizers (Supplementary Table 6). These results are in agreement with those of Khan et al. (2007) and Goswami et al. (2016), who showed that phosphate-solubilizing microorganisms could account for, respectively, from 1 to 50% and 20 to 40% of culturable populations in the soil. Among dominant genera, *Massilia* could also play a key role and a novel species, *M. phosphatilytica*, has been described with a phosphate-solubilizing potential of 40.32 µg in 48 h (Zheng et al., 2017). Potassium, calcium, or zinc solubilizers such as *Pseudomonas*, *Paenibacillus*, *Pontibacter*, *Variovorax*, or *Bacillus* improve plant nutrition and were also retrieved in our sequences (Supplementary Table 6). For iron, 14 genera could possess enzymes involved in this pathway, as predicted by PICRUSt2 and 36 sequences resembled genera containing isolated strains that produce siderophores (Supplementary Table 6). Sulfate is another important nutrient for plant development (Smith et al., 2000). PICRUSt2 prediction also identified genera increasing the availability of sulfate for plants, 26 genera potentially able to oxidize thiosulfate whereas two genera were found to contain isolated strains involved in sulfur metabolism such as *Thiobacillus* or *Stenotrophomonas*. These bacteria are also able to solubilize phosphate and K/Ca- or Na-bearing minerals due to the large quantity of sulfuric acid they produce (Ullah et al., 2013; Thabet et al., 2017). Microorganisms that produce cellulase which breaks cellulose down, also facilitate plant growth (El-Sayed et al., 2014). PICRUSt2 predicted that 83 genera possess enzymes involved in this pathway, but only eight genera possessed strains able to degrade cellulose. PGP microorganisms also improve soil formation, texture, and structure, by helping to aggregate soil particles through the production of exopolysaccharides that promote the formation of strong biofilms and hence the formation of soil crust (Kaushal and Wani, 2016; Thavamani et al., 2017). Microorganisms

such as *Leptolyngbya*, *Pseudomonas*, or *Rhizobium* that are able to increase soil aggregation were retrieved in our phosphate mining wastes (Supplementary Table 6). For phytohormone, 96 genera should produce IAA according to PICRUSt2, while our literature survey indicated that 43 genera contain at least one isolated strain able to produce this hormone (Supplementary Table 6). PICRUSt2 prediction was in agreement with other studies showing that around 80% of microorganisms have been documented to produce IAA (Patten and Glick, 1996; Kochar et al., 2013). Strains producing other phytohormones were also retrieved including gibberellin (20 strains) and cytokinin (10 strains, Supplementary Table 6).

In alkaline phosphate mining wastes, more than 27 sequences resembled genera containing isolated strains that produce ACCD, able to inhibit the formation of ethylene, the stress hormone with *Sphingomonas*, *Brevundimonas*, *Arthrobacter*, *Devosia*, or *Pseudomonas* being the most abundant (Supplementary Table 6). Many microorganisms known to be capable of combat phytopathogens were also retrieved in the present study: *Pseudomonas*, *Bacillus*, *Streptomyces*, *Stenotrophomonas*, *Rhizobium*, or *Streptomyces*, as well as less well-known microorganisms, such as *Massilia* or *Sphingomonas* (Supplementary Table 6). *Pseudomonas* sp. is able to inhibit diseases caused by many pathogens and possesses antimicrobial, antifungal, and antiviral properties, in addition to providing protection against insects (Kumari et al., 2019; Pattnaik et al., 2019, Supplementary Table 6). These fast-growing genera induce systemic resistance and produce different compounds including antibiotics, polysaccharides, or siderophores (limiting iron for phytopathogens, Santoyo et al., 2012). *Bacillus* sp. is known to produce a wide range of antibacterial and antifungal antibiotics (Goswami et al., 2016). *Streptomyces* are also known to produce many antibiotics (Olanrewaju and Babalola, 2019). Different genera, able to decrease or even suppress development of nematodes, were also retrieved, including *Arthrobacter*, *Pseudomonas*, *Streptomyces*, *Bacillus*, *Rhizobium*, or *Variovorax* (Topalovic et al., 2020). *Sphingomonas* sp. can suppress disease symptoms and limit the growth of the foliar pathogen of tomato, *Pseudomonas syringae* (Bulgarelli et al., 2013). Strains such as *Paenibacillus*, *Streptomyces*, *Bacillus*, or *Pseudomonas* produce lytic enzymes.

Many microorganisms present in our phosphate mining wastes are also related to strains able to improve growth of plants under drought stress; *Pseudomonas*, *Azospirillum*, *Paenibacillus*, *Rhizobium*, *Bacillus*, or *Variovorax* are among these well-known microorganisms (Supplementary Table 6). Indeed, *Azospirillum* sp. is able to mediate drought tolerance in many plants, even in deserts, by modifying root development and increasing water contents, grain yield, and mineral quality (Mg, K, and Ca) by producing IAA, gibberellin, or abscisic acid (a stress phytohormone that controls stomatal closure and stress signal transduction, Vurukonda et al., 2016; Fukami et al., 2018). *Bacillus* sp. is also known to increase the growth of *Lavandula* under drought conditions by producing IAA, improving physiology, increasing metabolic activity, and enhancing plant nutrition (Armada et al., 2014). Saline stress is also currently considered to be one of the main constraints
affecting both plants and microorganisms (Kaushal and Wani, 2016; Etesami and Beattie, 2018; Numan et al., 2018; Gupta and Pandey, 2019). Even if electrical conductivity was not high in our study, the high evaporation rate in the area could lead to high salinity in soil surface. Many microorganisms can improve the growth of plants in saline conditions, including strains of *Stenotrophomonas*, *Azospirillum*, *Bacillus*, and *Pseudomonas* (Supplementary Table 6). *Pseudomonas* sp. has been shown to improve the tolerance of wheat seedlings and sorghum to high temperatures by increasing levels of cellular metabolites (Ali et al., 2009; Vurukonda et al., 2016).

Diverse sequences related to resistant genera to different types of metals were identified, reflecting the stressors present in phosphate mining wastes in Morocco (Supplementary Table 6). This was particularly true for three metals that are present in large quantities, Cd, Cr, and Sr. Twenty-six genera contained strains resistant to Cd, but only six were resistant to Cr and four to Sr. High concentrations of Cd can reduce photosynthesis, nutrient, and water uptake, leading to growth inhibition, chlorosis, etc., and finally to death (Yadav et al., 2010). Chromium excess inhibits plant growth, causes nutrient imbalance, chlorosis, and root injury, and affects germination (Yadav et al., 2010), while excess Sr usually inhibits plant growth and weight (Burger and Lichtscheidl, 2019). *Sphingomonas* sp. was shown to actively increase the growth and stress tolerance of many plants in the presence of metals such as cadmium (Asaf et al., 2020, Supplementary Table 6). Long et al. (2017) also highlighted the capacity of *Bacillus* to immobilize Sr from aqueous solution by sorption. Fluorine, also present in quite large quantities in this study, can have also detrimental effects on the environment and human health (Fordyce, 2011; Wang et al., 2018) and can affect the growth of microorganisms (Li et al., 2014). In our phosphate wastes, various microorganisms were related to strains known to be tolerant to fluorine including *Bacillus*, *Actinobacter*, or *Pseudomonas* (Wang et al., 2018, Supplementary Table 6).

In light of these results, the microbial community present in the phosphate store-and-release cover seems not to be responsible for the almost absence of plants, explaining that their supplement in sufficient quantities may help in the phytostabilization of the phosphate cover. Further studies need to be done in order to address this conundrum.

## CONCLUSION

This study examined the composition and structuration of bacterial and archaeal communities in alkaline phosphate mining wastes in a Moroccan mine. The dominant sequences belonged to *Proteobacteria*, *Bacteroidetes*, *Actinobacteria*, and *Gemmatimonadetes* and to the genera *Massilia*, *Sphingomonas*, *Adhaeribacter*, *Hymenobacter*, and *Brevundimonas*. Significant differences in the composition of microbial communities were found between the two types of mining wastes studied (phosphate waste rocks and phosphate sludge), and differences were also found due to the presence of some indigenous annual plants. Soil pH, P₂O₅, Fe₂O₃, MgO, CaO, SiO₂, TOC, and N were, respectively, identified as the main drivers influencing microbial

community structure. This study improves our knowledge of the microbial communities present in alkaline phosphate mining wastes. This work also highlights the very high abundance of predicted plant growth-promoting microorganisms that can improve soil formation, nutrient uptake, and alleviating the biotic and abiotic stresses of this harsh environment in semi-arid areas. Furthermore, a great majority of our sequences resemble genera possessing more than one potential plant growth trait, which could be very useful in revegetation projects. In light of these results, the exploration of taxonomic composition of the phosphate store-and-release cover seems not to be able to explain the almost absence of plants, with a quite diversified communities, similar to other studies, without any particular structural issues and with a majority of microorganisms presenting plant growth-promoting factors. This is probably due to the harsh conditions of this damaged soil in semi-arid area that could prevent the plants from growing *in situ*. Future efforts should be oriented toward isolation of these interesting genera, using them as an inoculum to help the growth of local plants in such harsh environment. An in-depth understanding of the phosphate mine waste microbial communities and of their potential metabolic capabilities can provide useful information to prepare media dedicated to the isolation of the most interesting plant growth-promoting microorganisms identified in this study, a work currently in progress.

## DATA AVAILABILITY STATEMENT

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found below: https://www.ebi.ac.uk/ena, PRJEB39890.

## AUTHOR CONTRIBUTIONS

OB, LS, ME, and RH designed and contributed to the conception of the work. AO identified the plants and contributed to the sampling along with NM, OB, and RH. NM performed the experiments. NM and OB analyzed the data. RH, ME, AO, and LS contributed to the data interpretation. NM and OB wrote the first draft of the manuscript. All authors reviewed and approved the final version of this manuscript.

## FUNDING

This work was supported by the Moroccan Ministry of Higher Education (MESRSFC), the CNRST under Grant Number PPR1/2015/64, and the Ph.D. fellowship of NM by IRD (French National Research Institute, 2018–2020).

## SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2021.666936/full#supplementary-material
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**Conflict of Interest:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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# Microbial Diversity and Activity During the Biodegradation in Seawater of Various Substitutes to Conventional Plastic Cotton Swab Sticks

Justine Jacquin<sup>1,2</sup>, Nolwenn Callac<sup>1,3</sup>, Jingguang Cheng<sup>1</sup>, Carolane Giraud<sup>1,3</sup>, Yonko Gorand<sup>4</sup>, Clement Denoual<sup>5</sup>, Mireille Pujo-Pay<sup>1</sup>, Pascal Conan<sup>1</sup>, Anne-Leila Meistertzheim<sup>6</sup>, Valerie Barbe<sup>7</sup>, Stéphane Bruzaud<sup>5</sup> and Jean-François Ghiglione<sup>1*</sup>

OPEN ACCESS

**Edited by:**
Mechthild Schmitt-Jansen,
Helmholtz Centre for Environmental
Research (UFZ), Germany

**Reviewed by:**
Gabriela Vázquez-Rodríguez,
Autonomous University of the State
of Hidalgo, Mexico
Christian Lott,
HYDRA Marine Sciences GmbH,
Germany

*Correspondence:
Jean-François Ghiglione
ghiglione@obs-banyuls.fr

**Specialty section:**
This article was submitted to
Microbiotechnology,
a section of the journal
Frontiers in Microbiology

Received: 09 September 2020

Accepted: 12 May 2021

Published: 15 July 2021

**Citation:**
Jacquin J, Callac N, Cheng J, Giraud C, Gorand Y, Denoual C, Pujo-Pay M, Conan P, Meistertzheim A-L, Barbe V, Bruzaud S and Ghiglione J-F (2021) *Microbial Diversity and Activity During the Biodegradation in Seawater of Various Substitutes to Conventional Plastic Cotton Swab Sticks*. *Front. Microbiol.* 12:604395.
doi: 10.3389/fmicb.2021.604395

<sup>1</sup> CNRS, UMR 7621, Laboratoire d'Océanographie Microbienne, Observatoire Océanologique de Banyuls, Sorbonne Université, Paris, France, <sup>2</sup> Innovation Plasturgie et Composites, Biopole Clermont Limagne, Saint-Beauzire, France, <sup>3</sup> CNRS, UMR 9220 ENTROPIE, Ifremer (LEAD-NC), IRD, Univ Nouvelle-Calédonie, Univ La Réunion, Nouméa, New Caledonia, <sup>4</sup> Plateforme EnRMAT, Laboratoire PROMES, Rembla de la Thermodynamique, Perpignan, France, <sup>5</sup> UMR CNRS 6027, Institut de Recherche Dupuy de Lôme (IRDL), Université de Bretagne-Sud, Lorient, France, <sup>6</sup> SAS Plastic@Sea, Observatoire Océanologique de Banyuls, Banyuls-sur-Mer, France, <sup>7</sup> Génomique Métabolique, Genoscope, Institut François Jacob, CEA, CNRS, Univ Evry, Université Paris-Saclay, Evry, France

The European Parliament recently approved a new law banning single-use plastic items for 2021 such as plastic plates, cutlery, straws, cotton swabs, and balloon sticks. Transition to a bioeconomy involves the substitution of these banned products with biodegradable materials. Several materials such as polylactic acid (PLA), polybutylene adipate terephthalate (PBAT), poly(butylene succinate) (PBS), polyhydroxybutyrate-valerate (PHBV), Bioplast, and Mater-Bi could be good candidates to substitute cotton swabs, but their biodegradability needs to be tested under marine conditions. In this study, we described the microbial life growing on these materials, and we evaluated their biodegradability in seawater, compared with controls made of non-biodegradable polypropylene (PP) or biodegradable cellulose. During the first 40 days in seawater, we detected clear changes in bacterial diversity (Illumina sequencing of 16S rRNA gene) and heterotrophic activity (incorporation of ³H-leucine) that coincided with the classic succession of initial colonization, growth, and maturation phases of a biofilm. Biodegradability of the cotton swab sticks was then tested during another 94 days under strict diet conditions with the different plastics as sole carbon source. The drastic decrease of the bacterial activity on PP, PLA, and PBS suggested no bacterial attack of these materials, whereas the bacterial activity in PBAT, Bioplast, Mater-Bi, and PHBV presented similar responses to the cellulose positive control. Interestingly, the different bacterial diversity trends observed for biodegradable vs. non-biodegradable plastics allowed to describe potential new candidates involved in the degradation of these materials under marine conditions. This better understanding of the bacterial
diversity and activity dynamics during the colonization and biodegradation processes contributes to an expanding baseline to understand plastic biodegradation in marine conditions and provide a foundation for further decisions on the replacement of the banned single-used plastics.

Keywords: plastisphere, biofouling, plastic biodegradation, single-used plastics, microbial colonization

INTRODUCTION

The growing worldwide use of plastics together with waste mismanagement resulted in an estimated release of more than five trillion plastic particles weighing over 250,000 tons afloat in the oceans, resulting in dramatic toxicological effects along the marine trophic chain (Eriksen et al., 2014; Wang et al., 2019). A large majority of plastic items found at sea is made of polyethylene (PE), polypropylene (PP), and polystyrene (PS), which are more likely to float, and polyvinyl chloride (PVC), nylons, and polyethylene terephthalate (PET), which are more likely to sink (Auta et al., 2017).

Government regulations have been recently established to restrict the use of single-use plastics and promote the biodegradable plastic industry. For example, since 2018, at least 127 countries have established regulations against plastic bags (UNEP, 2018). The European Parliament voted to ban many other single-use plastics for 2021, including plates, cutlery, straws, food containers, and expanded polystyrene cups as well as sticks used for cotton swabs and plastic balloons (Europarl, 2019). Biodegradable plastics have become the focus of recent researches in order to replace conventional materials made of PE, PP, PS, PVC, PET, and other high-molecular compounds. Today, commercially available biodegradable materials are of petrochemical origin, such as poly(butylene succinate) (PBS) and polybutylene adipate terephthalate (PBAT; brand name Ecoflex®), or are bio-based such as polylactic acid (PLA) and polyhydroxybutyrate-valerate (PHBV). Natural starch is also frequently offered as a solution, blended with petrochemical-based (mainly potato starch for Bioplast®) or bio-based materials (corn starch with PBAT for Mater-Bi®).

Plastic biodegradation is the microbial conversion of all its organic constituents to carbon dioxide, new microbial biomass and mineral salts under oxic conditions, or to carbon dioxide, methane, new microbial biomass, and mineral salts, under anoxic conditions (Science Advice for Policy by European Academies [SAPEA], 2020). The biodegradability of a plastic is defined according to several standards reflecting disposal condition in soil, compost, or aquatic environments. Tests are generally based on respirometry (biological oxygen demand and CO2 production) attributed to the biodegradation of the plastic by microbial communities, without any other source of carbon in artificial medium, in comparison with cellulose, which serves as a reference control (standards ISO 18830:2016, ISO 19679:2016, ASTM D6691-09). Several drawbacks were pointed out by a recent paper on the ability of current standard protocols to realistically predict biodegradability in the marine environment (Harrison et al., 2018). For example, the use of preselected strains does not take into consideration the large diversity of

microorganisms colonizing plastics in natural conditions, also known as the “plastisphere” (Zettler et al., 2013) which is still poorly studied. In addition, the conventional techniques used to measure the biodegradation activity by evolution of O₂ or CO₂ generally require substantial equipment and can limit the number of experimental replicates. Other activity tests can be considered as alternative methods, such as the use of radiolabeled leucine incorporation that has been used for decades in marine microbiology (Kirchman et al., 1985) or tests based on ATP measurements (Fontanella et al., 2013). When introduced in seawater, plastics are rapidly colonized by bacteria. This process has been extensively studied, showing successive phases of biofilm formation (Dang and Lovell, 2000; Lobelle and Cunliffe, 2011; Eich et al., 2015; Erni-Cassola et al., 2019). During the first days at sea, a “conditioning film” made of inorganic and organic matter supports the initial colonization phase generally initiated by Gammaproteobacteria and Alphaproteobacteria, regardless of the material type (Oberbeckmann et al., 2015). A second phase of rapidly growing bacteria is then usually characterized by a succession of Bacteroidetes, Acidobacteria, Actinobacteria, Cyanobacteria, Firmicutes, and Planctomycetes (Salta et al., 2013). The last phase corresponds to a mature biofilm that generally appears after 15–30 days at sea and remains stable for several months, and where clear differences in bacterial abundance, diversity and activity, were found between non-biodegradable and biodegradable plastics (Eich et al., 2015; Dussud et al., 2018). Hydrocarbonoclastic bacteria (HCB) such as *Alcanivorax* sp., *Aestuariicella hydrocarbonica*, and *Marinobacter* sp. became abundant in the mature biofilm and were suggested to be potentially involved in the degradation of fossil-based plastics. However, no study has extended the experience by incubating the mature biofilm in minimum medium with the plastic as the only carbon source in order to prove its biodegradation capabilities.

Here, we used hollow cylinder sticks, such as those used for cotton swabs, made of different conventional (PP) and “biodegradable” materials as potential candidates for substitutes to conventional single-use plastics (PLA, PBS, PBAT, Mater-Bi, Bioplast, PHBV, and cellulose). We hypothesized that the properties of the distinct materials would select different marine bacterial communities with specific biodegradation capabilities. We elaborated an original two-step experimental design based on a first colonization step in natural seawater (40 days) followed by a second step of incubation in minimum medium with each plastic type as sole source of carbon and energy (94 days). Temporal dynamics of the biofilms associated with each plastic type were followed by measuring bacterial activity (³H-leucine incorporation), morphology [scanning electron microscopy (SEM)], and diversity (Illumina sequencing of 16S rRNA genes) during the successive colonization and biodegradation phases.
# MATERIALS AND METHODS

## Plastic Stick Manufacturing Process

The following list respectively, mentions the material type, its trade name, and the supplier: PP (ISPLEN PP 030 G1E, Repsol), PBAT (Ecoflex C1200, BASF), PLA (Ingeo 7001D, Natureworks), Bioplast (Bioplast 400, Biotec), PHBV (PHBHV ENMAT Y1000P, TianAn Bio), Mater-Bi (Mater-Bi EF04P, Novamont), PBS (bioPBS FZ91PB, PTT MCC Biochem), and cellulose (U Bio). PHBV was preferred in this study because it is a short-chain-length polyhydroxyalkanoate (scl-PHA) that is commercially available. Extended physical and chemical characteristics of the material types are given in **Supplementary Table 1**.

In order to manufacture plastic sticks that correspond to the hollow cylinder classically used for cotton swabs, we used a machine (except for cellulose sticks) with a spinning line composed of a SCAMEX single screw extruder with a 20-mm diameter and a L/D = 20 ratio. The machine was equipped with a vertical shaping die tube, a cooling tray, a drawing bench, an online dimensional controller (tolerances ± 0.01 mm), and a winding element. In order to conform the plastic materials to the desired stick form, the extruder was equipped with a suitable tube die tailor-made by the extruder manufacturer.

Extrusion conditions were adapted according to the used plastics and to their physical and chemical characteristics (**Table 1**). Finally, sticks were cut to equivalent dimensions as commercial sticks, i.e., 72-mm length, 2.5-mm outer diameter, and 1.6-mm inner diameter. This study required 300 fragments of 5 mm for each tested material. Except for cellulose, all materials had a hole in the stick.

## Incubation Under Natural Seawater Conditions (40 Days)

Each material type was incubated in aquariums with direct circulation to the sea, as previously described (Dussud et al., 2018). Briefly, we used nine identical aquariums consisting of trays with a 1.8-L capacity (Sodispan, Madrid, Spain), in which 1.5 L of 200-µm filtered seawater was continually renewed by direct pumping at 14-m depth in the Banyuls Bay, close to the SOLA marine observatory station (42°29'300 N-03°08'700 E, France). A flow rate of 50 ml min⁻<fnref n="1" /> was chosen to ensure a sufficient renewal of natural bacteria (every 30 min) and a homogeneous distribution of the plastic pieces in the aquariums during the entire experiment.

Approximately 250 pieces of each material type (PP, PLA, PBS, PBAT, Bioplast, Mater-Bi, PHBV, or cellulose) in the form of sticks of 0.5-cm length were washed three times with molecular ethanol and rinsed several times with sterile milliQ water before being placed in the aquariums, with the exception of the “control aquarium,” which only contained the circulating seawater. Pieces were sampled after 7 (D7), 15 (D15), 30 (D30), and 40 days (D40). Aquariums were kept in the dark to avoid UV-driven degradation of the materials. Throughout the incubation, seawater temperature (between 12.5 and 13.5°C) and salinity (38.5) were monitored in the aquariums and were similar to the data

found at the SOLA observatory in the Banyuls Bay during this period<fnref n="1" />.

## Incubation in Minimum Medium With Plastic Sticks as Sole Carbon Source (94 Days)

After 40 days under natural seawater conditions, the remaining pieces of each of the eight material types were carefully collected with sterilized tweezers and placed in 4-ml flasks (ref. 294330, Intechim, Montluçon, France) containing 2 ml of minimum medium with no other carbon source than the materials. The composition of the minimum medium was modified from Fegatella et al. (1998): NaCl (420 mM), Na₂SO₄ (28.8 mM), KCl (9.39 mM), NaBr (0.84 mM), H₃BO₃ (0.485 mM), MgCl₂·6H₂O (0.546 mM), CaCl₂ (0.105 mM), NH₄Cl (9.35 mM), SrCl₂·6H₂O (0.0638 mM), NaF (0.0714 mM), NaNO₃ (0.88 mM), NaH₂PO₄·H₂O (0.036 mM), KH₂PO₄ (0.106 mM), CuSO₄·5H₂O (0.04 µM), ZnSO₄·7H₂O (0.08 µM), CoCl₂·6H₂O (0.04 µM), MnCl₂·4H₂O (0.9 µM), Na₂MoO₄·2H₂O (0.03 µM), FeCl₃ (1.85 µM), thiamine (0.3 µM), biotin (2.1 nM), and vitamin B12 (0.317 nM). Pieces of each material type were sampled after 3 (D40 + 3), 7 (D40 + 7), 15 (D40 + 15), and 94 days (D40 + 94).

## Scanning Electron Microscopy

Three pieces of each of the eight material types were fixed at the beginning (D0) and at the end of the experiment (D40 + 94) with the addition of 3% (v/v) glutaraldehyde (final concentration) and stored for at least one night at 4°C. SEM was performed as previously described (Goldstein et al., 1992). Briefly, samples were washed in sodium cacodylate buffer and fixed with 1% osmium tetroxide before using successive EtOH baths (70, 95, and 100%) and liquid CO₂ for dehydration. About 10-nm conductive layer of Au-Pd was then applied on the surface of the samples to allow electron microscopy observation (Hitachi SEM FEG S-4500, Tokyo, Japan).

## Heterotrophic Bacterial Production

Incorporation of ³H-leucine is a classical proxy of the bacterial activity in seawater (Kirchman et al., 1985), which has been recently optimized for the plastisphere by Dussud et al. (2018). Bacterial production (BP) was measured on each material type at each sampling time during the colonization and biodegradation phases. A cell detachment pretreatment was applied using three cycles of 3-min sonication bath (Deltasonic, Meaux, France) followed by 3-min vortex at maximum speed (SkyLine, Elmi Ltd., Moscow, Russia). Immediately after cell detachment, ³H-leucine (specific activity 123.8 Ci mmol⁻<fnref n="1" />; PerkinElmer, Waltham, MA, United States) was added to all samples, which consisted of 1.5 ml of sterilized seawater containing the plastic pieces together with the detached cells. Biotic samples were treated in triplicates, and a fourth one was killed with trichloroacetic acid (TCA 50%) and served as a control of the reaction. Abiotic controls (consisting of plastic sticks of the eight material types incubated in sterile

<footnote n="1">¹https://www.somlit.fr/</footnote>
<table><caption>TABLE 1 | Extrusion conditions for the different plastic types.</caption><thead><tr><th rowspan="2">Plastic</th><th>Temperature zone 1</th><th>Temperature zone 2</th><th>Temperature zone 3</th><th>Temperature die</th><th rowspan="2">Screw speed<br>tr/min</th><th rowspan="2">Pulling speed<br>m/min</th></tr><tr><th>°C</th><th>°C</th><th>°C</th><th>°C</th></tr></thead><tbody><tr><td>PP</td><td>220</td><td>220</td><td>220</td><td>220</td><td>20</td><td>3.50</td></tr><tr><td>PLA 7001D</td><td>170</td><td>180</td><td>190</td><td>180</td><td>25</td><td>3.90</td></tr><tr><td>PHA Y1000P</td><td>180</td><td>180</td><td>180</td><td>180</td><td>25</td><td>4.00</td></tr><tr><td>PBAT C1200</td><td>140</td><td>160</td><td>160</td><td>160</td><td>15</td><td>2.90</td></tr><tr><td>PBS FZ91PB</td><td>140</td><td>140</td><td>140</td><td>140</td><td>20</td><td>3.50</td></tr><tr><td>Mater-Bi EF04P</td><td>140</td><td>160</td><td>160</td><td>160</td><td>15</td><td>3.15</td></tr><tr><td>Bioplast 400</td><td>160</td><td>160</td><td>160</td><td>160</td><td>15</td><td>3.00</td></tr></tbody></table>

PP, polypropylene; PLA, polylactic acid; PHA, polyhydroxyalkanoate; PBAT, polybutylene adipate terephthalate; PBS, poly(butylene succinate).

seawater) were analyzed following the same protocol. The final leucine concentrations were 36 nmol L⁻¹ (³H-leucine at 4 nmol L⁻¹) during the colonization phase and 150 nmol L⁻¹ after the transfer to minimum medium (³H-leucine at 1 nmol L⁻¹). The saturation of the leucine incorporation was checked during the biodegradation phase, and a higher final leucine concentration was required in order to avoid leucine being limiting (data not shown), explaining why the final leucine concentrations differed between the colonization and degradation phases. The final ³H-leucine concentration was 4.3 nmol L⁻¹ for control seawater samples. All samples were treated in triplicates and incubated in the dark at *in situ* temperature (between 12 and 15°C) for 2 h. Incubations were stopped by the addition of 50% TCA (5% final concentration), and the plastic sticks or cell pellets were washed with diluted TCA and then with cold ethanol. Radioactivity was counted after addition of liquid scintillator (Filter Count PerkinElmer) on a HIDEX 300 SL scintillation counter (LabLogic Systems, Inc., Tampa, FL, United States).

The empirical conversion factor of 1.55 ng C pmol⁻¹ of incorporated leucine was used to calculate BP (Simon and Azam, 1989). Because the exact surface of plastic sticks was difficult to measure (several layers for the cellulose, etc.), each piece was weighted with precision balance of sensitivity 0.1 mg (Precisa 125A, Swiss Quality; Precisa Gravimetrics AG, Dietikon, Switzerland), and the results were expressed in ng C·g⁻¹·h⁻¹.

## DNA Extraction, PCR, and Sequencing

For each material type, individual pieces were taken at all sampling times and stored at -80°C until DNA extraction. In parallel, 1 L of seawater was sampled in the control aquarium and successively filtered onto 3- and 0.2-µm pore size polycarbonate filters (47-mm diameter, Nucleopore), and filters were stored at -80°C until analysis. DNA extractions were performed on plastic sticks or filters using a classical phenol-chloroform method (Sauret et al., 2016) with slight modifications of the method for plastic samples (Debeljak et al., 2016). Briefly, the same cell detachment pretreatment was used as for BP (see above) before enzymatic and chemical cell lysis [incubation with 1 mg ml⁻¹ of lysozyme at 37°C for 45 min followed by 1 h at 50°C with 0.2 mg ml⁻¹ of proteinase K and 1% sodium dodecyl sulfate (SDS)]. DNA was quantified by spectrophotometry (Quant-iT™ PicoGreen™ dsDNA Assay Kit, Invitrogen; Thermo Fisher Scientific, Waltham, MA, United States).

PCR amplification of the V4-V5 region was performed using the primers 515-FY (5'-GTG YCA GCM GCC GCG GTA A-3') and 926-R (5'-CCG YCA ATT YMT TTR AGT TT-3') (Fuhrman et al., 1989), which are particularly well suited for marine samples (Parada et al., 2016). Sequencing was performed on Illumina MiSeq by MrDNA (Molecular Research LP, Shallowater, TX, United States) and Integrated Microbiome Resource (IMR; Dalhousie University, Halifax, NS, Canada). The sequences were analyzed using FROGS pipeline hosted in the Galaxy platform (Escudie et al., 2018), following the guidelines given in the publication. Briefly, the forward and reverse reads were assembled; and sequences were clustered using the SWARM algorithm (Mahé et al., 2014) and an aggregation distance of 3. Then, chimeras were removed using the FROGS Vsearch tool (Rognes et al., 2016). An additional filter step was used on the abundance at a 5 × 10⁻⁵ threshold to select the most relevant operational taxonomic unit (OTU) sequences. The taxonomic affiliation was assigned by standard nucleotid BLAST (Camacho et al., 2009) using the SILVA 132 database 16S (Quast et al., 2013). Chloroplast, mitochondrial, and eukaryotic sequences were removed. All the 16S rRNA data are available in the National Center for Biotechnology Information (NCBI) Sequence Read Archive (SRA) repository (accession number PRJNA649857).

## Statistical Analysis

In order to allow normalization and comparisons of the data, bacterial sequences of each sample were randomly resampled in the OTU file based on the sample with the fewest sequences (n = 9,698). These data sets were used to conduct all the analyses performed using the Rstudio software<fnref n="2" />. The α-diversity indices (Chao1, Shannon, evenness, and Simpson) and the β-diversity analysis (the Bray-Curtis similarity) were calculated with the phyloseq package (McMurdie and Holmes, 2013).

Differences between material types were tested using an ANOVA test and confirmed by a *post hoc* Tukey test. Results were considered to be significant if the *p* value was less than 0.05. Dendrogram was obtained based on the dissimilarity matrix of the Bray-Curtis and applying the Ward algorithm (Ward, 1963).

OTUs responsible for dissimilarities between pairs of clusters from the start and the end of incubation in

<footnote n="2">²https://www.r-project.org/</footnote>
minimum medium were identified using similarity percentage (SIMPER, PRIMER 6).

# RESULTS

## Scanning Electron Microscopy on the Different Plastic Types

Polypropylene, PLA, and PBS surfaces showed no visible signs of biodegradation (cracking, deformation, and holes). In addition, bacterial-cell structures on these materials were scarce and dispersed on the surface after 40 days in natural seawater followed by 94 days in minimum medium (D40 + 94). Abundant microbial colonization was observed on the surface of the other materials (PHBV, Mater-Bi, cellulose, and Bioplast), which is typical of a mature biofilm (Figure 1).

Three-dimensional biofilm structures were visible on the more colonized plastics, where cells colonized not only the surface layers but also the inner part of the visible cracks. A large diversity of morphological forms including spherical, rod-shaped, or spiral-shaped bacterial-like structures were observed on the surface of most of the materials. Extensive exopolysaccharide matrix was found except for PP, PLA, and PBS.

## Bacterial Production During the Successive Phases of Colonization and Transfer to Minimum Medium

During the colonization phase in natural seawater, BP was generally higher during the growth phase of the biofilm from 7 to 15 days. The highest BP values were found for PP, Bioplast, and PHBV (15.9, 12.4, and 10.5 ng C·g⁻¹·h⁻¹, respectively), whereas others (PLA, PBS, PBAT, and cellulose) ranged from 2.2 to 6.5 ng C·g⁻¹·h⁻¹ during this colonization phase (Figure 2A). At the end of the colonization phase in natural seawater (D40), BP was even more homogeneous between the material types, ranging from 1.6 to 4.0 ng C·g⁻¹·h⁻¹ for PP, PLA, PBS, PBAT and from 4.7 to 8.1 ng C·g⁻¹·h⁻¹ for Mater-Bi, Bioplast, PHBV, and cellulose. Bacteria living under control seawater conditions showed stable BP during the 40 days, ranging from 12.4 to 22.3 ng C·L⁻¹·h⁻¹ (Supplementary Table 1). No comparison was made between the BP values recorded between the seawater and the plastic sticks because of different measure units.

In order to anticipate the decrease of bacterial heterotrophic activity after the transfer of the plastic together with their biofilms into a minimum medium with the plastic as sole carbon source, we have changed the dilution factor of ³H-leucine added to the samples at day 40 accordingly to previous results (data not shown). It did not modify the leucine uptake, but it resulted in an increase of BP values by a factor of 5.8 ± 2.9 (n = 8) for all material types. This transfer from natural seawater to minimum medium resulted in different response scenarios according to the plastic types (Figure 2B). A first scenario was found for PP, PLA, and PBS where the BP increased during the first week of incubation in minimum medium (with maxima of 58.8, 85.9, and 94.2 ng C·g⁻¹·h⁻¹ at D40 + 3 or D40 + 7) and dramatically decreased thereafter (7.3, 8.7, and 11 ng C·g⁻¹·h⁻¹ at D40 + 94).

A second scenario was observed for PBAT, Mater-Bi, Bioplast, PHBV, and cellulose, where the BP remained relatively stable after the transfer into minimum medium (41.0, 18.6, 39.8, 59.2, and 21.9 ng C·g⁻¹·h⁻¹ at D40 + 94).

## Bacterial Community Structure, Diversity Indices, and Taxonomy During the Successive Phases of Colonization and Transfer to Minimum Medium

A total of 5,427,894 paired-end reads were acquired after sequencing all samples. After normalization based on the sample with the fewest merged sequences (n = 9,698), 3,772,255 sequence tags were selected belonging to 978 OTUs (0.03 distance threshold). The seawater control at D40 was sequenced several times, but each sequencing failed for unknown reasons.

### Changes in α-Diversity Indices

α-Diversity was studied using Chao1 richness, Pielou evenness, and Shannon diversity indices. Regardless of the plastic type, richness rapidly increased during the first 15 days of colonization and reached between 120 and 384 estimated OTUs after 40 days (Supplementary Table 2). The transfer to minimum medium did not influence richness (± 50 estimated species) for some material types including PP, PLA, PBS, and Bioplast but resulted in a decrease for PBAT, Mater-Bi, PHBV, and cellulose when comparing D40 and D40 + 94. We observed that the Chao1 index was statistically linked to the material type after the transfer to minimum medium (ANOVA, *p* value = 0.0147).

The Pielou evenness index showed relatively homogeneous diversity during the colonization phase, with evenness > 0.5 for all plastics after 40 days (Supplementary Table 2). A slight increase was found after the transfer in minimum medium for PP, Bioplast, PHBV, and cellulose, as correlated with the Chao1 index (p < 0.05), whereas it slightly decreased for PLA, PBS, PBAT, and Mater-Bi at the end of the incubation.

The resulting Shannon diversity was consistent with the Pielou and Chao1 indices. During the colonization phase, a decrease in the Shannon diversity index was found for the PP (2.4) due to its decrease in evenness (Table 1). A diversity increase was found for all the other plastics at D40 (between 2.64 and 4.40). The transfer to minimum medium resulted in an increase in diversity until the end of the experiment (between 3.67 and 3.9 at D40 + 94) for PP, PHBV, and cellulose according to the increase in evenness. Conversely, a decrease of the Shannon diversity (between 2.1 and 3.64) was caused by a decrease in richness for PBS, PBAT, and Mater-Bi or by a decrease in evenness for PLA and Bioplast.

### Microbial Communities on Plastics During the Colonization Phase in Seawater

The first colonizers growing on five of the seven plastic types immersed in seawater (PP, PLA, PBS, PBAT, and Mater-Bi) presented very similar communities (D7 and D15 of all material types grouped in Cluster 1) (ANOVA, *p* value = 0.221 and 0.121 for D7 and D15, respectively). Alphaproteobacteria represented 52% of the total OTU abundance in each sample, followed by Gammaproteobacteria (30%) and Bacteroidia
<figure><img src="image_103.png" /></figure>

**FIGURE 1** | Scanning electron microscopy of the eight polymer types (PP, PLA, PBS, PBAT, Bioplast, Mater-Bi, PHBV, and cellulose) showing the diversity of morphologies and abundance of bacteria cell-likes structures after 40 days in natural seawater followed by 94 days in minimum medium (D40 + 94). PP, polypropylene; PLA, polylactic acid; PBS, poly(butylene succinate); PBAT, polybutylene adipate terephthalate; PHBV, polyhydroxybutyrate-valerate.

<figure><img src="image_104.png" /></figure>

**FIGURE 2** | Bacterial production (in ng C·g⁻¹·h⁻¹) (A) during the colonization phase in natural seawater at days 7 (D7), 15 (D15), and 40 (D40) and (B) after the transfer to minimum medium during 3 (D40 + 3), 7 (D40 + 7), 15 (D40 + 15), 30 (D40 + 30), and 94 days (D40 + 94). The vertical bars represent the average of the bacterial production for each material (n = 3) ± standard deviation. Changes in BP values at D40 between panels (A,B) correspond to changes on dilution factor of the ³H-leucine added (see section "Materials and Methods"). BP, bacterial production.
(9%). The two most represented families were affiliated to the Alphaproteobacteria, with the Rhodobacteraceae (24%) (mainly *Sulfitobacter* sp. and *Pseudopelagicola* sp.) and the Metyloligellaceae (21%). Flavobacteriaceae (6.7%) (mainly *Croceibacter* sp. and *Aquibacter* sp.) and Nitrinocolaceae (mainly *Neptunibacter* sp.) (5.5%) were also abundant during this stage. Bacteria growing on Bioplast and cellulose were grouped in a separate cluster (Cluster 3, **Figure 3**), with a lower majority of Alphaproteobacteria (32%) and higher presence of Bacteroidia (25%), together with similar abundance of Gammaproteobacteria (30%) as compared with the other plastic types (see above). Alteromonadaceae family dominated (24.5%) and was mainly represented by the *Alteromonas* sp. and *Marinobacter* sp. Flavobacteriaceae family represented 22% of the biofilm community, mainly represented by the *Wenyingzhuangia* sp. Finally, like the other materials, the Rhodobacteriaceae family was very abundant (21%) and was mainly represented by the genus *Sulfitobacter* sp.

Clear changes in bacterial community structures were observed after the growth phase, with clear dissimilarities between the distinct plastic types after 40 days in natural seawater (D40). PP and PLA biofilms (Cluster 2) were dominated by Alphaproteobacteria (Rhodobacteriaceae) and Bacteroidia (Flavobacteriaceae). PBS and PBAT (Cluster 5) were dominated by Alphaproteobacteria (mainly Rhodobacteraceae) and Gammaproteobacteria (mainly Solimonadaceae). Bioplast and cellulose (Cluster 4) were dominated by Alphaproteobacteria (mainly Hyphomonadaceae) and Gammaproteobacteria (mainly Alteromonadaceae).

PHBV was dominated by Alphaproteobacteria (Rhodobacteraceae) followed by Gammaproteobacteria (Cellvibrionaceae) and finally Bacteroidia (Flavobacteriaceae). As for Mater-Bi, its mature biofilm was dominated by Alphaproteobacteria (Rhodobacteraceae), followed by Gammaproteobacteria (Spongiibacteraceae) and Bacteroidia (Flavobacteriaceae).

## Biofilm Specialization After Transfer to Minimum Medium

The transfer from natural seawater to minimum medium resulted in bacterial community changes following three different scenarios according to the material types. First, the structures of the bacterial communities associated with PP, PBAT, and PBS remained stable and distinct during the 94 days of incubation. The second scenario was shared between most of the plastic types, where the initial biofilm, established during the colonization phase, remained stable for 1 week after the transfer in minimum medium (D40 + 7) and then switched to another community that remained stable until the end of the experiment (D40 + 94). This scenario was found for Bioplast, Mater-Bi, PHBV, and cellulose, even though bacterial communities growing on each material were grouped in distinct sub-clusters. A third scenario was found for PLA, which presented a chaotic dispersion of the bacterial communities among all other clusters, even if the community at the end of the experiment clustered in the Cluster 7, together with three sub-clusters composed of (i) Mater-Bi, (ii) Bioplast, and (iii) PLA, PHBV, and cellulose.

A drastic increase of Bacteroidetes and a decrease of Alpha- and Gammaproteobacteria was observed at the end of the experiment (D40 + 94) in Cluster 7. The evolution of the major OTUs found on each plastic type at the beginning (D40) and at the end (D40 + 94) of the incubation in minimum medium is described in detail in **Figure 4**.

During seawater incubation, the mature biofilm associated with the non-biodegradable PP and PLA (D40) was largely dominated by *Polycyclovorans* sp. and *Marivita* sp. (more than 62 and 49% for PP and PLA, respectively). These two OTUs became scarce at the end of the incubation in minimum medium (<2.3% at D40 + 94). Bacterial lineages present on PP after 94 days under this drastic condition were hydrocarbonoclasts *Balneola* sp., *Marinobacter* sp., *Winogradskyella* sp., and *Lewinella* sp., explaining 13.15% of the difference between D40 and D40 + 94 and representing 20% (SIMPER analysis) of the relative abundance of the biofilm during the specialization phase. The major OTU at D40 + 94 on PLA was *Planctomicrobium* sp. (56.7% of the community), which contributed to 32.31% of the difference with D40 (SIMPER analysis), where it was not detected. Other genera have emerged at D40 + 94 on this plastic such as *Arenibacter* sp., *Aureisphaera* sp., and *Marixanthomonas* sp. representing almost 11% of the relative abundance at the end of the incubation.

The mature biofilm on PBS (D40) was dominated by *Maribius* sp., *Neptuniibacter* sp., and *Bacillus* sp. representing nearly 21% of the total OTUs, while these three genera represented only 0.64% at D40 + 94. At the end of the experiment (D40 + 94), the biofilm was dominated by *Filomicrobium* sp. (31%), *Thalassobius* sp. (3%), and *Bythopirellula* sp. (2%) explaining about 23% of the disparity with the D40. These three bacterial genera were scarce until transfer into minimum medium (3%).

The dominant OTUs found in the mature biofilm (D40) on PBAT were Rhodobacteraceae at 20.37% (*Marivita* sp., *Lentibacter* sp.), Solimonadaceae at 7.1% (*Lewinella* sp.), and Methylologellaceae at 5.6% and then switched to Saprospiraceae (42.5%) and *Filomicrobium* sp. (15.8%) at D40 + 94. These two OTUs alone explained 34.6% of the difference between the two sampling dates. As already found for PP and PLA, *Polycyclovorans* sp. was very abundant at D40 (7.1%) and was no longer detected at D40 + 94.

Incubation in minimum medium with Mater-Bi as sole carbon source resulted in the selection of *Aquimarina* sp., *Winogradskyella* sp., *Maritimibacter* sp., and *Filomicrobium* sp., which represented nearly 48% of the total OTUs at D40 + 94 but less than 2% at D40. These four genera explained 27.8% of the difference between the two sampling dates based on SIMPER analysis.

The OTUs selected at D40 + 94 for Bioplast were dominated by Microtrichaceae (12%), which belonged to the rare biosphere at D40 (<0.05%). Other genera emerged (> 9%) including *Aliiglaciecola* sp., *Filomicrobium* sp., and *Flexitrhrix* sp., while representing low abundance at D40 (<0.4%). On the contrary, *Alteromonas* sp. and *Oleibacter* sp. were abundant (8.8%) at D40 and became rare at D40 + 94 (<0.05%).

Selection on minimum medium with PHBV as sole carbon source showed the emergence of *Sphingorhabdus* sp., *Lewinella* sp.
<figure><img src="image_105.png" /><figcaption>**FIGURE 3** | Comparison between hierarchical clustering based on the Bray–Curtis similarity between the temporal dynamic of bacterial communities growing on the eight material types (top part) and their taxonomic affiliation by cumulative charts comparing relative class of abundances (bottom part).</figcaption></figure>

sp., *Pseudahrenzia* sp., and *Marinobacter* sp. (> 21% after D40 + 94 and <0.6% at D40), which explained 17.26% of the difference between D40 and D40 + 94 according to SIMPER analysis (Figure 4), whereas the dominant *Sedimentitalea* sp., *Thalassotalea* sp., and *Lentibacter* sp. at D40 (20% of the total OTUs) represented less than 0.1% at D40 + 94.

Cellulose selected *Parvularcula* sp., *Oleiphilus* sp., *Pirellula* sp., and *Phycisphaera* sp. at D40 + 94 (21.5% of the total OTUs at D40 + 94 compared with <1.5% at D40), which explained 16% of the difference between the two sampling dates. *Hirschia* sp. remained relatively stable between D40 (11.4%) and D40 + 94 (7.32%).

We finally focused on 35 different genera present on all the plastics and that may be considered as potential HCB. In this functional group, Gammaproteobacteria were overrepresented (40% of putative hydrocarbon degraders) followed by Bacteroidia (31%) and Alphaproteobacteria (23%). The most abundant

genera potentially involved in hydrocarbon degradation were *Neptuniibacter* sp., *Sulfitobacter* sp., *Balneola* sp., *Lentibacter* sp., and *Marinobacter* sp., which represented 52% of the total OTUs. Overall, putative HCB were abundant during the growth and maturation phases of the biofilms in seawater (between 8.4 and 38.0% of the total communities), with higher abundance generally found during the growth phase for all plastics (Table 2). Interestingly, the putative HCB decreased drastically until the end of the incubation in minimum medium with the plastics as sole carbon sources (D40 + 94). One exception was found for PP, which showed the highest percentage of putative HCB on the mature biofilm (38%), with even higher percentage found after 30 days in minimum medium (63.6%) mainly composed of *Balneola* sp., *Winogradskyella* sp., and *Marinobacter* sp. Other putative HCBs, such as *Polycylovorans* sp., *Neptuniibacter* sp., *Croceibacter* sp., and *Colweillia* sp. (Figure 4), were found in seawater during the colonization phase of most of
<figure><img src="image_106.png" /><figcaption>**FIGURE 4** | Evolution of the major OTUs (> 4%) on the different materials according to the date of incubation in seawater for 40 days (D40) and in minimum medium for 94 days (D40 + 94). OTUs, operational taxonomic units.</figcaption></figure>

<table><caption>**TABLE 2** | Percentage of OTU selected as putative HCB on each material (PP, PLA, PBS, PBAT, Mater-Bi, Bioplast, PHBV, and cellulose) according to the date of incubation in seawater during 7, 15, and 40 days (D40) followed by the transfer in minimum medium during 3 (40 + 3), 7 (40 + 7), 15 (40 + 15), 30 (40 + 30), and 94 days (40 + 94).</caption><thead><tr><th rowspan="2">Days</th><th colspan="3">Natural seawater</th><th colspan="5">Minimum medium</th></tr><tr><th>7</th><th>15</th><th>40</th><th>40 + 3</th><th>40 + 7</th><th>40 + 15</th><th>40 + 30</th><th>40 + 94</th></tr></thead><tbody><tr><td>Bioplast</td><td>21.1</td><td>21.6</td><td>12.1</td><td>55.1</td><td>7.1</td><td>9.6</td><td>5.8</td><td>6.6</td></tr><tr><td>Mater-Bi</td><td>23.0</td><td>14.5</td><td>13.3</td><td>37.3</td><td>19.5</td><td>6.2</td><td>10.7</td><td>13.4</td></tr><tr><td>PBAT</td><td>31.9</td><td>21.3</td><td>21.0</td><td>54.7</td><td>4.8</td><td>26.3</td><td>19.9</td><td>8.6</td></tr><tr><td>PBS</td><td>21.7</td><td>20.7</td><td>19.8</td><td>12.4</td><td>32.7</td><td>27.7</td><td>22.9</td><td>8.6</td></tr><tr><td>PHBV</td><td>16.0</td><td>18.5</td><td>8.4</td><td>33.4</td><td>15.8</td><td>8.3</td><td>46.1</td><td>11.1</td></tr><tr><td>PLA</td><td>35.1</td><td>22.3</td><td>22.7</td><td>1.4</td><td>20.1</td><td>NA</td><td>35.5</td><td>4.9</td></tr><tr><td>PP</td><td>32.5</td><td>26.6</td><td>38.0</td><td>21.3</td><td>37.8</td><td>60.2</td><td>63.6</td><td>26.6</td></tr><tr><td>Cellulose</td><td>17.4</td><td>16.3</td><td>11.2</td><td>14.1</td><td>13.9</td><td>3.3</td><td>0.9</td><td>12.3</td></tr></tbody></table>

OTU, operational taxonomic unit; HCB, hydrocarbonoclastic bacteria; PP, polypropylene; PLA, polylactic acid; PBS, poly(butylene succinate); PBAT, polybutylene adipate terephthalate; PHBV, polyhydroxybutyrate-valerate.

the materials but did not succeed in colonizing the plastics in minimum medium.

## DISCUSSION

### Formation of Mature Biofilms on all Plastic Types in Seawater

#### Aquarium With Direct Circulation to the Sea Mimics the Natural Seawater Conditions of the North Western Mediterranean Sea

The colonization of the different material types was feasible thanks to distinct incubation in aquariums with seawater renewal

every 30 min. These aquariums presented similar hydrological conditions to those found at the SOLA observatory in the Banyuls Bay (NW Mediterranean Sea, France). During the first 40 days of incubation in seawater, temperature (13°C ± 0.5°C), chlorophyll *a*, and nutrient values (data not shown) were similar to those usually observed at the long-term SOLA observatory station (Ghiglione et al., 2005; Lambert et al., 2019). Bacterial heterotrophic activity in seawater, measured by ³H-leucine incorporation, reached 22.3 ng C·L⁻¹·h⁻¹, which was similar to what was generally reported *in situ* in the NW Mediterranean Sea (Pulido-Villena et al., 2012). These results showed that incubation in our aquariums with direct circulation to the sea allowed to reproduce environmental conditions, thus making extrapolation of our results to natural seawater possible.
## Growth and Maturation Phases of the Biofilms on all the Plastic Types in Seawater

In agreement with patterns observed in another study (Salta et al., 2013), we found a succession of growth and maturation phases of the biofilms for all the material types during the first 40 days of incubation in natural seawater. Each phase was linked with bacterial communities that were similar during the growth phase but became specific to the material types when the mature biofilm formed. This result is in agreement with a previous study conducted in the same region (Dussud et al., 2018).

Bacterial heterotrophic activity peaked during the growth phase of the biofilm in all plastic types (either after 7 or 15 days). These bacterial communities were very similar regardless of the plastic types, except for Bioplast and cellulose, which clustered in another group. This result was surprising considering the distinct chemical material compositions and surface properties of the plastics used in our study. It is noteworthy that bacterial communities growing on Mater-Bi (about 60% corn starch and 40% PBAT) did not cluster with cellulose and Bioplast at this stage (about 40% potato starch and 60% PLA), probably because of different surface properties of the materials that may be related to distinct polymer compositions together with the unknown effect of potentially incorporated additives (not mentioned by the manufacturer). Indeed, early stages of biofilm formation have been depicted as strongly dependent on the surface properties of plastics, including hydrophobicity, structure, and roughness (Lorite et al., 2011). This was not obvious for immersed plastics in seawater, since our results were similar with other studies that did not find distinct communities during the first weeks of biofilm formation on different plastic types (Harrison et al., 2014; De Tender et al., 2017; Dussud et al., 2018). Regardless of the plastic type, the first growth phase was mainly dominated by Proteobacteria (84% of the total abundance on all materials), followed by Bacteroidetes (12.5%). The dominance of Proteobacteria during the first stage of colonization has already been described on the plastisphere (Jacquin et al., 2019). The main classes of the Proteobacteria were Alphaproteobacteria (47.14%, mainly Methyloligellaceae and Rhodobacteraceae) and Gammaproteobacteria (31.46%). The Methyloligellaceae family has already been isolated on other plastic types such as polyamide, polycarbonate, or PP (Feng et al., 2020). All studied plastics were mainly colonized by members of the Rhodobacteraceae during the colonization phase and the specialization phase. This family has been depicted as a key taxonomic group in marine biofilm formation on different material types (Elifantz et al., 2013; Kviatkovski and Minz, 2015; Roager and Sonnenschein, 2019), which explained its large presence on the tested plastics. During the colonization phase, the Flavobacteriaceae family and particularly the genus Neptuniibacter sp., generally able to degrade hydrocarbons, was found in abundance on PLA, PBS, and Mater-Bi (Gutierrez, 2019). This result is consistent with a previous study that showed its presence only during the primary colonization phase of plastics under similar conditions (Dussud et al., 2018). In all material types, the mature biofilm that formed after 40 days of incubation in seawater showed lower values of bacterial heterotrophic activity compared with the growth phase. This is

consistent with observations made in the unique study measuring bacterial heterotrophic activities on the plastisphere (Dussud et al., 2018). It can be explained by the equilibrium reached by a biofilm at its maturity, which is related to the decrease of available labile inorganic and organic matter and also to the complex interactions (competition, commensalism, etc.) between species coexisting on plastics (Lorite et al., 2011; Henson and Phalak, 2017). Diversity in mature biofilms became specific for each plastic type, which is consistent with observations made in the natural environment (Webb et al., 2012; De Tender et al., 2017; Pinto et al., 2019).

## Convergent Signs of Biodegradation for Some Plastic Types

As recommended in several standard tests (ISO 18830:2016, ISO 19679:2016, and ASTM D6691-09), the biodegradation of the studied plastics was tested after transferring the plastic pieces into minimum medium with no other carbon or energy source (ISO 18830:2016, ISO 19679:2016). As suggested in these tests, we used cellulose as a positive control, and we considered PP as a negative control since its biodegradation was previously depicted to take years or decades in the environment (Barnes et al., 2009). Standards allow the possibility of using two types of inoculum, i.e., artificial mixing of culture of microorganisms or bacterial communities from natural seawater. These methods have been criticized since they are not able to reproduce the mature biofilms usually found on plastics in the environment (Harrison et al., 2018; Jacquin et al., 2019). In our study, we followed the development of a natural mature biofilm on eight different material types (40 days' incubation in seawater) before using that biofilm as an inoculum for further biodegradation tests.

After 94 days in minimum medium, SEM observations showed no surface modifications for PP, PLA, and PBS, whereas PBAT, Bioplast, Mater-Bi, PHBV, and cellulose exhibited observable morphological alterations, with clear evidence of swelling and erosion. Visible alterations of plastic surfaces are considered to be suitable signs of biodegradation, even if these alterations alone are not sufficient (Zettler et al., 2013). Plastics, such as PHBV, are known to undergo biofragmentation by numerous exo-enzymatic actions (amylases, proteases, and nucleases), leading to weakening of the surface of the material (Deroiné et al., 2014a, 2015). Our results are consistent with a previous study showing evidence of faster degradation of PHAs in seawater compared with other plastics such as PBS, PBAT, or PLA (Sashiwa et al., 2018). In our conditions, PHBV showed more cracks, holes, and colonization features than PBAT, Bioplast, Mater-Bi, PHBV, and cellulose (Figure 1). In parallel, SEM observations at the end of the experiment showed dispersed and isolated bacteria for PP, PLA, and PBS, whereas dense biofilms with a three-dimensional structure and substantial extracellular matrix were found for all the other plastics. PLA was previously found to degrade very slowly in seawater and remain stable at room temperature (Deroiné et al., 2014b), while PBS degradation was found to be slower when salinity increased
(Silva Dutra et al., 2019), which are also consistent results with our observations. In our conditions, the radiolabeled leucine incorporation as a proxy of bacterial biodegradation has drawn the same conclusion as the SEM analysis. Interestingly, this is the first time that this technique has been used for this purpose, whereas it has been used for decades to measure the microbial heterotrophic activity and growth in seawater (Kirchman et al., 1985).

It has been successfully used to prove the biodegradability of several other substrates, such as antifouling biocides (Maraldo and Dahllöf, 2004), hydrocarbons (Peng et al., 2020), plasticizers, and flame retardants (Vila-Costa et al., 2019), but never for plastic materials in seawater, probably because of the lack of studies in marine environments. In seawater, this technique is generally preferred to respirometry measurements (O₂ consumption or CO₂ production) because its sensitivity is more suitable in oligotrophic waters (Briand et al., 2004; Pulido-Villena et al., 2012). Current standards are based on respirometry analyses and generally measure CO₂ by gas chromatography, total organic carbon analyzer, infrared analyzer (ISO 14855-1), or CO₂ capture by soda lime (ISO 14855-2). These experimental approaches can be automated but require a significant financial investment and large laboratory spaces, which can reduce the number of experimental replicates. This has already been described as one of the limitations of the current standards on plastic biodegradation (Harrison et al., 2018). Here, we found that other activity tests based on ³H-leucine incorporation were well suited for biodegradation tests of plastic materials in seawater. Our experiments showed that bacteria living in the mature biofilm can continue to be active for several days after transfer under minimum medium with no other carbon source, as it is the case for PP, PLA, and PBS, which showed bacterial heterotrophic activities during 1 or 2 weeks before falling off. Cross-feeding of metabolites synthetized by one species to support the growth of another species to maximize utilization of available resources, together with competition to limited resources, may explain the activity observed during several days and its further decline (Pande et al., 2014; Henson and Phalak, 2017). This is an important point because such an activity could be misinterpreted as a sign of plastic biodegradation if integrative measurements were only taken at the end of the experiment. For PBAT, Bioplast, Mater-Bi, PHBV, and cellulose, the bacterial heterotrophic activity remained rather stable after 15 days in minimum medium and until the end of the experiment (94 days). Taken together with congruent results from SEM observations, we consider that the radiolabeled leucine incorporation measurement was a good proxy of their biodegradation. Our results are consistent with previous findings showing the biodegradation in soil, compost, or seawater for PBAT (Saadi et al., 2013; Weng et al., 2013; Wang et al., 2021), Mater-Bi (Kim et al., 2000), Bioplast<fnref n="3" />, PHBV (Arcos-Hernandez et al., 2015; Deroiné et al., 2015; Wang et al., 2021), and cellulose (Salehpour et al., 2018; Kliem et al., 2020; Zambrano et al., 2020). It is interesting to note that PLA, a plastic considered to be compostable and biodegradable in soil and compost

(Ren et al., 2007), did not show any sign of biodegradation under our marine experimental conditions.

## Identification of Putative Biodegraders of the Tested Materials

According to SEM and heterotrophic bacterial activity measurements, we focused on the OTUs selected at the end of the incubation in minimum medium on the plastic materials that showed clear signs of biodegradation under our experimental conditions: PBAT, Bioplast, Mater-Bi, PHBV, and cellulose. The presence of complex communities living under such strict conditions strongly suggest that they are putative biodegraders that may interact with one another in order to perform all or part of the degradation until the ultimate mineralization in CO₂ (Dussud and Ghiglione, 2014). We are aware that part of the identified OTUs may not be involved in the plastic degradation but rather opportunists that may survive in the biofilm (Sauret et al., 2014). Our original procedure allowed to present here a list of putative OTUs selected from a mature biofilm grown under natural seawater conditions, which may be involved in biodegradation of the plastics.

On the PBAT, we found *Thalassobius* sp. as dominant OTUs after 94 days in minimum medium with the mature biofilm and its plastic as sole carbon source. This genus has already been suggested as potentially capable of degrading aromatic hydrocarbons in seawater (Rodrigo-Torres et al., 2017) and found in the oil polluted waters of the Gulf of Mexico (Liu et al., 2017). *Filomicrobium* sp. were also dominant at the end of the experiment and have already been observed in other marine environments (Wu et al., 2009), as well as *Prevotella* sp. and *Streptococcus* sp. known as potential pathogenic genera (Fittipaldi et al., 2012; Larsen, 2017). None of them were previously identified as potential PBAT degraders.

The bacteria growing with Mater-Bi as sole carbon source was dominated by *Filomicrobium* sp., already identified on the plastisphere of PE and PET (Muthukrishnan et al., 2018). Other groups were *Winogradskyella* sp., *Maritimibacter* sp., *Aquimarina* sp., and *Halomonas* sp., which were previously described as putative HCB (Mnif et al., 2009; Liu et al., 2017). None of them were previously identified as potential Mater-Bi (starch with PBAT) degraders.

Bacteria selected on Bioplast at the end of the experiment included *Lewinella* sp., already identified on the plastisphere of fossil-based PP, PE, and PET, as well as on biobased PHBH films (Morohoshi et al., 2018; Roager and Sonnenschein, 2019). Other potential OTUs belonging to putative HCB were *Croceitalea* sp., *Hirschia* sp., *Maribacter* sp., and *Cellulophaga* sp. None of them were previously identified as being potentially able to degrade Bioplast (starch with PLA).

Microbial communities selected after 94 days on PHBV as sole carbon source such as *Marinobacter* sp. and *Alteromonas* sp. were already known for their ability to degrade PHBV (Mergaert et al., 1995; Martinez Tobon, 2019). *Marinobacter* sp. genera contain well-known HCB capable of degrading benzene, toluene,

<footnote n="3">³https://fr.biotec.de/</footnote>
ethylbenzene, and xylene (BTEX) (Li et al., 2012), long-chain alkanes (Grimaud et al., 2012), polycyclic aromatic hydrocarbons (Gao et al., 2013; Cui et al., 2016), chlorinated solvents (Inoue et al., 2020), phenol (Zhou et al., 2020), and PHA (Kasuya et al., 2000; Martinez Tobon, 2019).

Finally, it is noteworthy that putative HCB were less abundant during the biodegradation phase (from 6 to 12% of the total OTUs) than during the colonization phase (from 8 to 35% of the total OTUs). These results were unexpected since several authors proposed that the HCB are certainly involved in plastic biodegradation since they are always found as dominant members of the microbial communities growing on fossil-based plastics recovered from the sea surface (Bryant et al., 2016; Pollet et al., 2018; Amaral-Zettler et al., 2020). Here, we showed that some HCB may actually be involved in biodegradation, but they were not a majority under our experimental conditions. HCB generally present a high ability to attach to particles and to live in a biofilm (Grimaud et al., 2012), two factors that may be more important than their ability to degrade plastics.

## Concluding Remarks

The decisions taken by the European government to reduce the use of single-use plastics and promote biodegradable materials lead to finding substitutes for conventional plastics. Here, we emphasized that materials such as PHBV, Mater-Bi, Bioplast, PBAT, and cellulose are good alternatives to banned cotton swabs generally made of non-biodegradable PP. It is noteworthy that the bio-based polymer PLA showed no signs of biodegradation under our marine conditions, whereas the fossil-based polymer PBAT showed significant evidence of biodegradation. This reinforced the distinction that has to be made between biobased and biodegradable polymers when considering alternative to conventional fossil-based plastics.

Our approach, coupling bacterial diversity and activity measurements, brought new insights on the ability of natural biofilms to biodegrade plastic materials in seawater, with this information being essential for our understanding of the microbial communities involved in the plastosphere and for future alternative biodegradation tests of plastic items in the marine environment. Our study also opens new perspectives by identifying putative bacteria potentially capable of biodegrading the tested plastic materials. These results require further studies to better understand the still largely unexplored functional processes involved in biodegradation in natural seawater.

## REFERENCES

* Amaral-Zettler, L. A., Zettler, E. R., and Mincer, T. J. (2020). Ecology of the plastosphere. *Nat. Rev. Microbiol.* 18, 139–151. doi: 10.1038/s41579-019-0308-0

* Arcos-Hernandez, M., Montano-Herrera, L., Janarthanan, O. M., Quadri, L., Anterrieu, S., Hjort, M., et al. (2015). Value-added bioplastics from services of wastewater treatment. *Water Pract. Technol.* 10:546.

* Auta, H., Emenike, C., and Fauziah, S. H. (2017). Distribution and importance of microplastics in the marine environment: a review of the sources, fate, effects, and potential solutions. *Environ. Inter.* 102, 165–176.

## DATA AVAILABILITY STATEMENT

The datasets presented in this study can be found in online repositories. The names of the repository/repositories and accession number(s) can be found in the article/ Supplementary Material.

## AUTHOR CONTRIBUTIONS

J-FG, JJ, and NC conceived and designed the study. NC, JJ, CG, JC, and YG carried out all the experiments and acquired the data. SB and CD provided the equipment. NC, JJ, and J-FG analyzed and interpreted the data. JJ, J-FG, NC, CG, and VB drafted the manuscript. MP-P, PC, and A-LM revised the manuscript and approved the version of the manuscript to be published. All authors contributed to the article and approved the submitted version.

## FUNDING

This project was funded by the Ministry of Ecological and Solidarity Transition of the French Republic (project BIOTIGE). This work was also supported by the European project JRA-ASSEMBLE+, the French National Research Agency (project ANR-OXOMAR), the Tara Ocean Foundation (project Microplastic 2019), and the CNRS (project PEPS’I-PHABIO).

## ACKNOWLEDGMENTS

We are grateful to Valerie Bayo and Charlene Odobel for their kind laboratory support and to Guigui PA and VF for insightful comments on the manuscript. This work is part of the PhD thesis of JJ supported by the French Ministry of Education.

## SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmicb.2021.604395/full#supplementary-material

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**Conflict of Interest:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

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# Bioleaching of E-Waste: Influence of Printed Circuit Boards on the Activity of Acidophilic Iron-Oxidizing Bacteria

Juan Anaya-Garzon<fnref n="1" />,², Agathe Hubau<fnref n="1" />*, Catherine Joulian<fnref n="1" /> and Anne-Gwénaëlle Guezennec<fnref n="1" />

Bioleaching is a promising strategy to recover valuable metals from spent printed circuit boards (PCBs). The performance of the process is catalyzed by microorganisms, which the toxic effect of PCBs can inhibit. This study aimed to investigate the capacity of an acidophilic iron-oxidizing culture, mainly composed of *Leptospirillum ferriphilum*, to oxidize iron in PCB-enriched environments. The culture pre-adapted to 1% (w/v) PCB content successfully thrived in leachates with the equivalent of 6% of PCBs, containing 8.5 g L⁻<fnref n="1" /> Cu, 8 g L⁻<fnref n="1" /> Fe, 1 g L⁻<fnref n="1" /> Zn, 92 mg L⁻<fnref n="1" /> Ni, 12.6 mg L⁻<fnref n="1" /> Pb, and 4.4 mg L⁻<fnref n="1" /> Co, among other metals. However, the inhibiting effect of PCBs limited the microbial activity by delaying the onset of the exponential iron oxidation. Successive subcultures boosted the activity of the culture by reducing this delay by up to 2.6 times under batch conditions. Subcultures also favored the rapid establishment of high microbial activity in continuous mode.

**Keywords:** iron-oxidizing bacteria, *Leptospirillum ferriphilum*, printed circuit boards, bioleaching, pre-oxidation phase, subculturing

## OPEN ACCESS

**Edited by:**
Mechthild Schmitt-Jansen,
Helmholtz Centre for Environmental
Research (UFZ), Germany

**Reviewed by:**
Sabrina Hedrich,
Freiberg University of Mining
and Technology, Germany

Eva Pakostova,
Coventry University, United Kingdom

**\*Correspondence:**
Agathe Hubau
A.Hubau@brgm.fr

**Specialty section:**
This article was submitted to
Microbiotechnology,
a section of the journal
Frontiers in Microbiology

**Received:** 19 February 2021

**Accepted:** 12 July 2021

**Published:** 18 August 2021

**Citation:**
Anaya-Garzon J, Hubau A, Joulian C and Guezennec A-G (2021) *Bioleaching of E-Waste: Influence of Printed Circuit Boards on the Activity of Acidophilic Iron-Oxidizing Bacteria*. *Front. Microbiol.* 12:669738.
doi: 10.3389/fmicb.2021.669738

## INTRODUCTION

Printed circuit boards (PCBs) are the main processing unit for the electronic devices that have become essential to our daily lives in recent decades. Their disposal at the end of their useful life is a major challenge for the circular economy and sustainable development. Improper handling of waste PCBs is widespread, with incineration and landfilling as the easiest but polluting solutions. Waste PCBs contain heavy metals (Pb, Hg, As, Cd, Cr), polychlorinated biphenyls, polybrominated biphenyls, and epoxy resins, which are highly toxic for the environment (Kaya, 2019). Likewise, spent PCBs are a valuable source of various strategic metals, including Au, Ag, Pd, Ti, Ga, Ta, Co, Cu, Fe, Al, Zn, Sn, Pb, and Ni (Hubau et al., 2019; Kaya, 2019).

The dissolution of metals assisted by microorganisms, also known as bioleaching, is a biotechnological process that is now established worldwide. This technology is already applied at an industrial scale for the extraction of Au and Cu, and marginally for Ni, Co, and U, from primary ores (Rawlings and Johnson, 2007; Wassink and Asselin, 2019). It is now attracting considerable attention for the extraction of metals from secondary sources such as mining waste, contaminated river sludge (Zhang et al., 2018), fly ash (Xu et al., 2014), batteries (Liu et al., 2020), and spent PCBs (Bryan et al., 2015, 2020; Guezennec et al., 2015; Işıldar et al., 2016; Wu et al., 2018; Hubau et al., 2020). Some of the advantages of biotechnologies compared to conventional pyrometallurgical

<footnote n="1">¹ Bureau de Recherches Géologiques et Minières, Orléans, France, ² Chimie ParisTech, PSL Research University, CNRS, Institut de Recherche de Chimie Paris, Paris, France</footnote>