Fate of Eight Different Polymers under ... - ACS Publications

Subscriber access provided by UNIV OF CALIFORNIA SAN DIEGO LIBRARIES

Article

Fate of eight different polymers under uncontrolled composting conditions: relationships between deterioration, biofilm formation and the material surface properties Anne Mercier, Kevin Gravouil, Willy Aucher, Sandra Brosset-Vincent, Linette Kadri, Jenny Colas, Didier Bouchon, and Thierry Ferreira Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.6b03530 • Publication Date (Web): 23 Jan 2017 Downloaded from http://pubs.acs.org on January 28, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Environmental Science & Technology is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 32

Environmental Science & Technology

1

Fate of Eight Different Polymers under Uncontrolled Composting Conditions:

2

Relationships Between Deterioration, Biofilm Formation and the Material Surface

3

Properties

4 5 6 7 8 9 10 11 12 13 14 15 16

Anne Mercier†, Kevin Gravouil†, Willy Aucher†, Sandra Brosset-Vincent†, Linette Kadri†, Jenny Colas†, Didier Bouchon†,‡, Thierry Ferreira†*

17

ABSTRACT

18

With the ever-increasing volume of polymer wastes and their associated detrimental impacts

19

on the environment, the plastic life-cycle has focused increasing attention. Here, eight

20

commercial polymers selected from biodegradable to environmentally-persistent materials, all

21

formulated under a credit card format, were incubated in an outdoor compost to evaluate their

22

fate over time and to profile the microbial communities colonizing their surfaces. After 450

23

days in compost, the samples were all colonized by multispecies biofilms, these latest

24

displaying different amounts of adhered microbial biomass and significantly distinct bacterial

25

and fungal community compositions depending on the substrate. Interestingly, colonization

26

experiments on the eight polymers revealed a large core of shared microbial taxa,

27

predominantly composed of microorganisms previously reported from environments

28

contaminated with petroleum-hydrocarbons or plastics debris. These observations suggest that

29

biofilms may contribute to the alteration process of all the polymers studied. Actually, four

30

substrates, independently of their assignment to a polymer group, displayed a significant

31

deterioration, which might be attributed to biologically-mediated mechanisms. Relevantly, the

32

deterioration appears strongly associated with the formation of a high-cell density biofilm

†

Laboratoire coopératif ThanaplastSP - Carbios Bioplastics - Ecologie et Biologie des Interactions, Centre National de la Recherche Scientifique, UMR 7267, Université de Poitiers, Poitiers, France ‡ Equipe Ecologie Evolution Symbiose, Ecologie et Biologie des Interactions, Centre National de la Recherche Scientifique, UMR 7267, Université de Poitiers, Poitiers, France

1 ACS Paragon Plus Environment


33

onto the polymer surfaces. The analysis of various surface properties revealed that roughness

34

and hydrophilicity are likely prominent parameters for driving the biological interactions with

35

the polymers.

36 37

ABSTRACT ART

38 39 40

Keywords: polymer, biofilm, biodegradable, surface property, plastic, deterioration

41 42

INTRODUCTION

43

Due to their unique chemical properties, both natural and synthetic polymers are widely used

44

worldwide for a large variety of applications including the formulation of plastics, which offer

45

many advantages over other materials, such as high performance and durability, ease of

46

processing, high productivity and low production costs.1 Constant innovations in the plastic

47

industry have resulted in technical breakthroughs in medicine, agriculture or transportation in

48

particular.1 Nevertheless, this widespread and increasing use of plastics is also directly

49

responsible for the serious public and environmental issues related to their accumulation as

50

wastes, which have emerged within the last 40 years.2-5 Accidental disposal of solid polymers,

51

the major components of plastics, are visible as litter in the environment, including open

52

landfills, coastal sediments, oceans and rivers and can induce harmful environmental

53

damages.4,6 Without preventive actions, such as implementation of remediation or 2 ACS Paragon Plus Environment

Page 2 of 32

Page 3 of 32


54

introduction of new environmentally compatible and friendly polymers, this anthropogenic

55

contamination and the ensuing long-term threats to living organisms are expected to last for

56

decades to centuries.2

57

Polymer degradation is achieved mostly through scission of the main or side-chains to

58

transform polymers into oligomers, dimers and finally monomers. These initial cleavage steps

59

are usually catalysed by abiotic factors such as UV-radiation, thermal activation or/and

60

chemical treatments.7-9 Also, polymers may undergo biodegradation through microbial

61

hydrolysis and oxidation. This process occurs via several sequential steps, including bio-

62

deterioration, bio-fragmentation, bio-assimilation and bio-mineralisation.4,8,10 Due to their

63

high molecular weights and their poor solubility in water, long-chain polymers are not easily

64

captured and assimilated by surrounding microorganisms.11 To circumvent this limitation,

65

secretion of extracellular enzymes and free-radicals occurs first to depolymerise the polymer

66

chain(s) outside the cells. Subsequently, when the molecular weight of the polymer is

67

sufficiently reduced, intermediate products can be transported into the cytoplasm of the cell

68

and introduced into the microorganisms’ metabolic pathways. Biodegradation requires a

69

crucial initial step that is the formation and development of a microbial biofilm, either at the

70

surface of the polymer or directly into the polymer matrix itself following abiotic alterations,

71

in a process which is influenced by the chemical and the physical properties of the polymer,

72

as well as by the environmental conditions.11-15 Surprisingly, data presently available on

73

polymer-associated microbial communities are scarce compared to the large amount of studies

74

on biofilm formation and development on other physical supports.16 To date, the few studies

75

on the microbial communities inhabiting the so-called “plastisphere” are mostly related to the

76

micro- to macro- plastic debris collected from sea environments, and are mainly focused on

77

the potential toxicity of persistent organic pollutants adsorbed to plastic particles and the risk

78

of pathogen dispersion with plastic transport.5,16-19 Nevertheless, the bio-mineralization



79

(resulting in excretion of completely oxidized metabolites: H2O, CO2, N2, CH4) appears as a

80

suitable long-term answer for the management of polymer wastes once they have reached the

81

environment, since it can result in the complete elimination of the primary and secondary

82

intermediate products and therefore avoid their leaching and/or accumulation in environments

83

with unknown and/or ignored effects on biodiversity and health. In this context, more

84

investigations are required to profile the microbial communities colonizing each type of

85

polymer, and to identify the main parameters driving microbial attachment onto polymer

86

surfaces, in relation with the polymer degradation efficiency.

87

The present study falls within this context. Here, cards obtained by compression moulding of

88

commercial granules from eight polymers of various natures were all incubated under the

89

same environmental conditions at a fixed location (an outdoor compost system) for the same

90

period of time, in order to investigate the fate of the substrates and to address the relationships

91

between the attached bacterial and fungal taxa, and the surface properties of the polymer

92

samples.

93 94

MATERIALS AND METHODS

95

Polymers, card preparation and surface properties. Eight different polymer types were

96

selected from biodegradable (or claimed as such) to environmentally-persistent materials

97

(namely polyesters, polyolefins and polyamide) and with the highest purity available from the

98

manufacturers (Table 1, see the chemical structures in Fig. S1, Supporting Information SI1).

99

The raw polymers were supplied as granules and used to form card (850 x 550 x 20 mm; see

100

SI2 for detail on card preparation).

101

The degree of crystallinity of each polymer card was determined using a Mettler Toledo

102

DSC1 (differential scanning calorimeter) by Technopolym (Toulouse, France). Contact angles

103

with three liquid probes were determined at the Alençon Institute of Plastics (ISPA, Alençon,


Page 4 of 32

Page 5 of 32


104

France), using a GBX Digidrop goniometer. The values were used to estimate the energy of

105

surface to the card surface as well as its dispersive and polar components. Additionally, the

106

surface micro-structures of each polymer card were observed using a Scanning Electron

107

Microscope JEOL JSM 5600LV (see SI2 for detail on surface property analyses).

108

Compost-incubated experiment. Fifteen clean cards of each polymer type were positioned

109

vertically in an outdoor compost (see SI2 for detail on compost site preparation). The natural

110

microorganisms found in the compost were regarded as the source of colonizers of polymer

111

cards and deterioration was based on naturally occurring composting reactions. Hence, no

112

microbial bio-augmentation and additional organic amendment were conducted over the

113

experiment. The experimental plot was subjected to the oceanic and temperate climate of the

114

region characterized by mild temperatures (on average 12.5-13°C), average rainfall of 800–

115

900 mm/year (Poitiers), and narrow seasonal ranges (http://www.infoclimat.fr/).

116

Compost-incubated experiments were conducted for 450 days. Ten replicate cards of each

117

type of polymer were weighed using an OHAUS TS120 Precision Standard analytical balance

118

(Bioblock Scientific) after washing, thoroughly scraping off to remove cells and particles,

119

rinsing and drying the cards to a constant weight was obtained. The polymer card weight loss

120

was calculated using the following formula: Percentage of weight loss = [(initial weight−final

121

weight after compost incubation) / initial weight] × 100.

122

DNA extraction from polymer card associated- and soil microbial communities. Three

123

compost-incubated cards of each polymer type were washed 4-times by complete immersing

124

and moderate movements in sterile water to remove non-adherent cells and particles to the

125

card surface. Both sides of the card were thoroughly scraped off for 5 min using a sterile cell

126

scraper perpendicular to the surface and rinsed with a total of 50 mL of sterile water.

127

Extractable fraction of biofilm was centrifuged 12 800 g, 20 min at 4 degrees C. The

128

supernatant was discarded and samples were stored at -20°C until DNA extraction.



Page 6 of 32

129

Additionally, the soil from 5 cores was taken in compost area surrounding the cards and

130

pooled to yield one single composite sample, sieved to 2 mm and portioned into 500 mg (dry

131

weight) aliquots used for analysis.

132

DNA extraction from the polymer card associated- and soil microbial communities were

133

performed using the same procedure based on the ISO 1106321 (see SI2 for detail on DNA

134

extraction and purification). Quality and size of the crude and purified extracted DNA were

135

checked by electrophoresis. DNA quantifications were estimated with the BIO-1D image

136

analysis software (Vilber Lourmat) using a standard curve of HindIII-digested λ DNA

137

fragments (Promega). Aliquots of 0.5 ng/µL diluted DNA were stored at -20°C ready for

138

molecular applications.

139

Pyrosequencing of 16S and 18S rRNA gene sequences. The microbial communities

140

inhabiting the surface of each type of polymer card were analysed by pyrosequencing of

141

ribosomal genes. Each sample was prepared from the same DNA input quantity in order to

142

normalize the libraries and achieve even representation of each library in the pyrosequencing

143

results. The V3-V4 region of the bacterial 16S rRNA genes was amplified using the primers

144

F479 (5′-CAGCMGCYGCNGTAANAC-3′) and R888 (5′-CCGYCAATTCMTTTRAGT-3′)

145

as described.21 Additionally, a fungal 18S rRNA gene fragment was amplified using the

146

primers

147

CGATAACGAACGAGACCT-3′)22 as described21 (see SI2 for detail on amplicon preparation

148

and purification). The two pyrosequencing runs were conducted out in a GS Junior Sequencer

149

(Roche Applied Science) following manufacturer's recommendations.

150

Bioinformatic analysis of 16S and 18S rRNA gene sequences. Pyrosequencing data were

151

analysed using the GnS-PIPE (version 1.1.13) pipeline21 (see SI2 for detail on bioinformatic

152

analysis). Normalization by random selection was conducted to obtain the same number of

153

reads for each sample and to avoid biased bacterial or fungal community comparisons.

FR1

(5′-ANCCATTCAATCGGTANT-3′)


and

FF390

(5′-

Page 7 of 32


154

All the taxonomy-based analyses using similarity approaches against SILVA r114 reference

155

database obtained from the GnS-PIPE were then post-processed using R packages and

156

BioVenn.23,24 The complexities of the bacterial and fungal communities adhered onto the eight

157

polymer surfaces and collected from the surrounding compost were investigated by measuring

158

within-sample diversity (α-diversity) including richness (number of genus-level Operational

159

taxonomic units (OTUs)), and the Shannon and Evenness indices, using the defined OTU

160

composition. Bray-Curtis dissimilarity distances were calculated from the normalized OTU

161

tables for each fraction to construct a dissimilarity matrix and Principal Coordinate Analysis

162

(PCoA) was carried out with the Vegan package in R.25 Datasets are available on the EBI

163

database system in the Sequence-Read Archive (SRA), under study accession numbers

164

PRJEB14758.

165

Statistical analysis. Comparisons in card weight were performed using Mann-Whitney test

166

(significance assessed at the level of p < 0.05). The Spearman’s rank correlation test was used

167

to measure the strength of the linear association between values from the microbial biomass

168

of the biofilm developed at the card surface and the compost-incubated card weight loss.

169

Comparison of multiple groups was performed using the R Vegan software program and the

170

analysis of similarity (ANOSIM).25 Data from card properties (Table 1) were fit to the

171

distance matrix generated from sequencing using the R Vegan software program and the

172

envfit function25, producing coefficient of determination r2 and combined p-values, in order to

173

evaluate the direct contributions of variables to the microbial community patterns (i.e.

174

bacterial and fungal β-diversity analysis). The significance of the variable fitting was

175

determined using 999 permutations.

176 177

RESULTS

178

Polymer card properties. In this study, the fate in compost of five polyesters (namely PBAT,



179

PBS, PET, PLLA and PHA; Table 1), four of them being considered as biodegradable, at least

180

under industrial, controlled composting conditions (PBAT, PBS, PLLA and PHA26), of two

181

polyolefins (EVOH and PP) and of one polyamide (PA66) was evaluated. Characteristic

182

surface properties of the polymer cards, with potential relations with biomass recruitment

183

and/or degradation were determined. A first observation was that the degree of crystallinity

184

varied greatly from one polymer to the other, with values ranging from 9.8 to 60.1%. PBS

185

cards appeared to be the most crystalline polymer whereas PET was the most amorphous

186

material (Table 1). Surface energy determinations revealed that PHA displayed the highest

187

surface energy with 60.0 mJ/m2, whereas values ranged from 32.1 to 48.9 mJ/m2 for the other

188

polymers. The dispersive component was quite homogenous among the various samples,

189

ranging from 31.8 to 44.6 mJ/m2. By contrast, the polar component was significantly higher

190

on the surface of PHA, with 22.7 mJ/m2, whereas this parameter ranged from 0.3 to 6.9 mJ/m2

191

for the other polymers (Table 1). Finally, the surface of the eight polymer cards displayed

192

different micro-structures (SI1 Fig. S2). The cards synthetized from PBS, PHA, PBAT and

193

PA66 showed a rough appearance, with micron-sized depressions and protuberances covering

194

their surface, while EVOH, PET, PP and PLLA cards appeared rather smoother and more

195

homogenous.

196

Compost-incubated polymer cards. In a next step, the polymer cards were removed from

197

compost after 450 days of incubation and their relative deterioration was assessed by weight

198

loss. The initial and final weights of the various samples are displayed in Fig. 1A.

199

Interestingly, the results of this study demonstrated that deterioration took place in our

200

compost simulation and showed that the weight loss did not occur to a similar extent for the

201

eight substrates studied. Two of the five polyesters and the polyamide polymers showed a low

202

but significant weight loss (%), with a decrease reaching 7.9 (± 0.7)%, 5.5 (± 0.3)% and 1.1

203

(± 0.2)% for PHA, PBS and PA66, respectively. Whereas weight loss was not significant (Fig.


Page 8 of 32

Page 9 of 32


204

1A), surface (SI1 Fig. S3) and mechanical alterations (i.e. increased flexibility) were

205

noticeable for compost-incubated PBAT cards. Therefore, for this specific polymer, an

206

additional Gel Permeation Chromatography (GPC) analysis was conducted to evaluate a

207

potential reduction of the average length of the PBAT chains after incubation in compost.

208

Relevantly, both Mw and Mn were decreased after compost incubation for this polymer (from

209

39 400 Da to 36 000 Da for Mw, and from 35 000 Da to 18 400 Da for Mn, respectively), and

210

the polydispersity index was increased (from 1.125 to 1.957 Da), showing a clear-cut

211

reduction of the PBAT chain length (SI1 Table S1).

212

Microbial biomass colonizing the polymer card surface. Relative DNA amounts were used

213

as estimates of the microbial biomass colonizing the surface of the polymer cards27. Results

214

revealed that a microbial colonization occurred for all the compost-incubated cards studied,

215

however to a variable extent depending on the polymer considered (Fig. 1B). Indeed, the

216

relative amounts of DNA recovered ranged from 28.2 (± 2.6) to 238.2 (± 27.0) ng DNA/cm2

217

of card. Remarkably, the formation of a high cell-density biofilm was quantified on both PHA

218

and PBS polyester cards, which displayed the most important weight loss (see above). By

219

contrast, the microbial colonization appeared to be lesser on PET (Mann-Whitney test

220

between values from PET and PHA; p < 0.05), which is also assigned to the polyester group,

221

but which remained unaltered under compost incubation. In compost, the microbial biomass

222

was estimated to 8.7 (±1.6) DNA/g of dry soil, in agreement with literature.27

223

Interestingly, a significant Spearman coefficient value of 0.738 (p < 0.05) revealed a positive

224

correlation between the microbial biomass adhered at the polymer surface and the card weight

225

loss after compost incubation.

226

Bacterial and fungal OTUs richness, Shannon and Evenness diversity indices. A high

227

throughput sequencing was used to characterize both bacterial and fungal communities

228

colonizing each polymer card surface. A total of 5 400 (with a mean length of 371 bases) and



229

3 000 reads (with a mean length of 317 bp) for bacteria and fungi respectively, were obtained

230

after sequencing quality filtering and homogenization steps, for each type of polymer card and

231

for the compost samples.

232

The compost-incubated PHA, PBS and PBAT polymer cards displayed a lower bacterial

233

richness on their surfaces than the one initially present in the compost, but also than the one

234

adhered to the surface of the other polymers studied (Table 2). Between 250 to 290 bacterial

235

OTUs were identified at 95% identity in the biofilm communities onto the surface of PHA,

236

PBS and PBAT, while more than 550 bacterial OTUs were assigned to the biofilm collected

237

from the surface of the PET card, for example. The Shannon index ranged from 3.82 to 5.56,

238

indicating a bacterial diversity on the surface of all the polymer cards. Nevertheless, the

239

Shannon index was also significantly lower for PBS, PHA and PBAT than for the other

240

polymers. The Evenness index of all the polymer samples showed few differences, ranging

241

from 0.69 to 0.88, suggesting there was not a discrepancy in the repartition of OTU

242

abundance in favor of a few, but highly dominant, bacterial OTUs on the polymer cards.

243

Interestingly, regarding fungal diversity metrics, compost-incubated PBS, PHA and PBAT

244

polymer cards displayed also a lower fungal richness and Shannon index than for the other

245

polymers (SI3 Table S1).

246

Bacterial and fungal community structures. A principal coordinate analysis (PCoA) from

247

Bray-Curtis distance matrix was performed to examine the overall variations among bacterial

248

and fungal communities colonizing the polymer cards after incubation in compost (Fig. 2A).

249

PCoA analysis showed that the three replicate cards from each type of polymer clustered

250

closely, underscoring the reproducibility of the bacterial and fungal community structures and

251

the robustness of the molecular characterization of these microbial communities. Moreover,

252

PCoA analysis also revealed that each polymer displayed the ability to support distinct

253

microbial communities (Fig. 2A and SI3 Fig. S1; ANOSIM permutation test, R = 0.918 and


Page 10 of 32

Page 11 of 32


254

0.728 for bacterial and fungal communities respectively, R significance = 0.001).

255

Interestingly, the bacterial communities colonizing the PHA, PBS and PBAT cards were

256

clearly distinct from those identified on the five other polymers, but also from the

257

autochthonous bacterial community of the surrounding compost: they clustered together,

258

along the first principal coordinates (PCoA1), which accounts for 34.18% of the total

259

variability (Fig. 2A). Along the PCoA2 axis (16.76% of the variability), the bacterial

260

community associated to the PHA card surface appeared clearly separated from those attached

261

to PBS and PBAT. Similar results were obtained from PCoA analysis of the fungal

262

communities colonizing the polymer cards (see SI3 Fig. S1 for description).

263

Thus, according to the two PCoA analyses, the microbial communities on the 8 polymers can

264

be clustered into three groups: PHA, [PBS, PBAT] and [PLLA, PA6.6, PP, EVOH, PET].

265

Bacterial and fungal community composition associated to polymer cards. Taxonomic

266

classification at the phylum level of the OTU representative sequences revealed the

267

dominance of Proteobacteria and Bacteroidetes in compost soil and at the surface of all the

268

incubated polymer cards, with 71.54 to 93.76% of the total pyrosequencing sequences being

269

assigned to these two phyla (SI4 Table S1). Members of the Planctomyces (< 5.26% of the

270

total pyrosequencing sequences) and Acidobacteria (< 4.89%) were also found. Notably, the

271

Firmicutes phylum reached 11.90% of the total pyrosequencing sequences at the surface of

272

the PET cards, a value higher than for the other polymers (0.79 to 8.32% only). The most

273

abundant bacterial classes were Alphaproteobacteria (9.81 – 23.60% of the total

274

pyrosequencing sequences at the class level), Gammaproteobacteria (8.01 – 16.66%),

275

Sphingobacteria (14.88 – 29.05%). Interestingly, the Betaproteobacteria class reached

276

15.14%, 23.98% and up to 25.19% of the total pyrosequencing sequences at the surfaces of

277

PHA, PBS and PBAT cards respectively, whereas their relative abundance remained in the

278

range of 4.06 to 7.18% for the other samples (SI4 Table S2). Differences between the



279

bacterial communities colonizing the polymer cards were also obvious at the family level (see

280

SI3 Fig. S5 for description and SI4 Table S3).

281

A similar fungal community composition was observed at the phylum level at the surface of

282

all the polymer cards, with a predominance of Ascomycota, Basidiomycota, Chytridiomycota

283

and Glomeromycota, their relative abundance varying with the type of polymer (SI4 table S4).

284

For example, the Ascomycota phylum reached 70.96 (± 2.48)% and 52.05 (± 4.57)% of the

285

sequences assigned at the surface of the PBAT and PHA cards, respectively, and only 8.64 (±

286

1.05)% for PET cards. Nevertheless, unknown fungi represented from 2.25% to 8.75% of the

287

total sequences assigned at the phylum level, unclassified fungi, 2.26% and 26.82%, and

288

environmental fungi, 8.09% and 44.71%. Differences in the compositions of the polymer-

289

associated communities were also highlighted at the fungal class level (SI4 Table S5). For

290

example, Dothideomycetes was the most represented fungal class observed at the surface of

291

PBAT cards (with 65.26 (± 4.03)% of the sequence assigned at the class level),

292

Sordariomycetes for PHA cards (33.48 ± 4.15%) and Agaricomycetes for EVOH cards (35.47

293

± 5.38%). Differences between the fungal communities colonizing the polymer cards were

294

also obvious at the family level (SI3 Fig. S2 for description and SI4 Table S6).

295

Comparison of bacterial and fungal communities colonizing polymers: unique and

296

shared OTUs. A three-set Venn diagram analysis was performed to represent the unique and

297

shared OTUs and sequences among the three bacterial and fungal community clusters

298

identified from the PCoA analyses (see above): this diagram was elaborated using the

299

communities colonizing either PHA, PET (as representative of the [PLLA, PA6.6, PP, EVOH,

300

PET] cluster) and PBS (as representative of the [PBS, PBAT] cluster). A total of 130 bacterial

301

OTUs were shared by the three community clusters and accounted for 87.53%, 78.66% and

302

94.47% of total classified sequences for PHA, PET and PBS, respectively (Fig. 2B).


Page 12 of 32

Page 13 of 32


303

Regarding the fungal communities (SI3 Fig. S3), a high proportion of the total sequences

304

were also common to the three defined polymer clusters. From 1.65% to 13.97% of total

305

bacterial and fungal classified sequences were identified in only one cluster and from 1.29%

306

to 21.60% in two clusters. This analysis revealed a common core within the bacterial and

307

fungal communities colonizing the surface of polymers (core microbiome). Hence, the

308

differences in the bacterial and fungal community structures observed as a function of the

309

polymer studied (as revealed by the PCoA analyses) resulted mainly from variable

310

abundances of dominant microbial OTUs, rather than from the emergence of specific yet rare

311

OTUs. Among the sequences of the core microbiome, the most represented OTUs belonged to

312

the bacterial phyla Bacteroidetes (with the families: Chitinophagaceae, Bacteroidaceae,

313

Cytophagaceae, Flavobacteriaceae) and Proteobacteria (with the families: Pseudomonadaceae,

314

Sphingomonadaceae), and to the fungal family Mortierellaceae.

315

Drivers of microbial colonization: relationships between material surface properties and

316

microbial taxa of biofilms. A combined analysis of the card-surface properties and of the

317

microbial community structure was conducted to evaluate the contribution of these physico-

318

chemical parameters (Table 1) to the polymer-associated microbial community patterns (Fig.

319

3 and SI3 Fig. S4). In the ordination plots, the length of the arrow corresponding to a given

320

surface parameter indicates the strength of the relationship between this parameter and the

321

community composition. From this analysis, it appeared that the best correlations were

322

obtained for the bacterial composition dissimilarity matrices from PHA and the polar

323

component (r2 = 0.907) and with the energy of surface (r2 = 0.665). The PBS and PBAT cards

324

were associated with the dispersive component (r2 = 0.347). All associated p-values were

325

below 0.01, indicating that these results can be considered as robust (Fig. 3). Interestingly,

326

crystallinity did not appear to be an important determinant of microbial community structure

327

on polymer surface (r 2 = 0.2657 for bacterial communities) since the correlation did not reach



328

a statistical significance (p > 0.01). Similar results were obtained from the fungal

329

communities (see SI3 Fig. S4 for description). Whereas the scanning electron microscopy

330

(SEM) observations were not converted into quantitative data, the biological interactions with

331

a given polymer appeared strongly dependent on its surface roughness, as suggested by the

332

presence of micron-sized irregularities covering the surface of the PHA, PBS, PBAT and

333

PA66 polymer cards (SI1 Fig. S2), providing favourable microniches for microbial

334

attachment.

335 336

DISCUSSION

337

Most of the current investigations on the biomass and diversity in microbial communities

338

colonizing the surface of plastics focus on marine debris, randomly collected from different

339

locations after variable submersion periods.18,19,28-32 Under these conditions, the samples

340

correspond almost exclusively to polypropylene and polyethylene-based plastic wastes, the

341

most prominent polymers used worldwide, and originate from different and unknown

342

production sites. As a consequence, they display, for a given type of polymer, different

343

molecular weight distributions and oxidation levels, and are likely to contain various levels of

344

adjuvants and prooxidant additives. Moreover, the exposition of the debris to abiotic stresses

345

such as UV-radiations, remain uncontrolled and is likely to vary greatly from one sample to

346

the other, making difficult a clear-cut correlation between the intrinsic properties of a given

347

polymer and the recruitment of a selective microbial community.19 Considering these

348

foregoing observations, and to avoid as much as possible some of these bias and allow direct

349

comparisons, eight different commercial raw polymer types, ranging from polyolefin,

350

polyester to polyamide groups, all formulated under the same credit-card format, were

351

incubated in the present study at a fixed outdoor location under uncontrolled conditions that

352

resemble home-composting conditions. This approach allowed us replicating the samples to


Page 14 of 32

Page 15 of 32


353

profile and compare the microbial communities forming a biofilm at the surface of a specific

354

polymer.

355

Whatever the substrate studied (i.e. claimed as biodegradable or environmentally-persistent,

356

chemical structure; Table 1), all compost-incubated cards were colonized by a multispecies

357

biofilm. Interestingly, some supports, independently of their belonging to a specific polymer

358

group, developed biofilms displaying significant variations in both bacterial and fungal

359

community compositions as compared to the surrounding compost soil, indicating that these

360

substrates represent distinct environmental niches. However, microspatial variations of the

361

soil microbial community cannot be completely excluded. Microbial richness and diversity

362

metrics confirmed that each polymer card develops a heterogeneous and complex ecological

363

system.

364

The microorganisms detected at the surface of polymer belonged mostly to the bacterial phyla

365

Proteobacteria and Bacteroidetes. The prevalence of these phyla has already been reported

366

from petroleum-hydrocarbon contaminated environments and some of their members have

367

been directly implicated in hydrocarbon degradation.16,33-35 Dominance of Proteobacteria

368

(mainly Alpha and/or Gammaproteobacteria) and Bacteroidetes has also been reported on

369

plastic debris collected from marine environments.19,31 Their relative abundance was used in

370

several studies as putative signatures of biofilm formation stages, Proteobacteria and

371

Bacteroidetes behaving as primary and secondary biofilm colonizers, respectively.18,36,37

372

Filamentous fungi were also detected at the polymer card surfaces, as from hydrocarbon-

373

contaminated soils.38-40 Some of these organisms have been reported to be potent hydrocarbon

374

bio-degraders, but fungi are also suspected to increase bacterial access to hydrophobic organic

375

substrates.41,42

376

The colonization experiments performed in this study also revealed that the different

377

polymers share a large core of microbial taxa, mainly composed of members of the



378

Bacteroidetes phylum. Members of this phylum, common in ecosystems, have been reported

379

to be fast-acting decomposers of the organic matter and are known to degrade high-molecular-

380

weight organic compounds, including petroleum hydrocarbons.43,44 Altogether, these

381

observations strongly support the presence of a biodiversity displaying potent microbial

382

polymer-degrading capacities on virtually all the card surfaces, most polymers being formed

383

from hydrocarbon molecules. Nevertheless, the direct involvement of all these

384

microorganisms in the alteration process should be now demonstrated. Relevantly, three of

385

the five polyesters (PHA, PBS and PBAT) and the polyamide (PA66) showed a significant

386

weight loss and/or visual surface alterations that might be attributed to biologically-mediated

387

processes based on our experimental conditions, in which abiotic factors such as mechanical

388

constraints and exposition to UV radiations are likely to be moderate (i.e. in a buried static

389

system) in comparison with aquatic systems. However, the weight loss remained low after

390

450 days in the compost, confirming the expected long–term persistence of polymeric

391

material contaminations in the environment. These results sustain that actions such as

392

preventing plastic waste from reaching the environment, optimizing plastic management

393

(reducing, reusing, recycling), or developing new environmentally compatible and friendly

394

polymers (i.e. truly degradable under natural conditions) should be promoted.

395

The most significant weight loss was observed with PHA, which is in good agreement with

396

previously published data on this hydrolysable polyester produced from renewable

397

resources.45 The hydrolysis of PHA can take place in different environments. However, the

398

degradation rate depends on several parameters, including the duration of the experiment, the

399

temperature and the soil pH, but also on the material thickness, experiments being often

400

conducted with thin films of few micrometers.45 Many microorganisms isolated from various

401

environments such as the model bacteria Pseudomonas putida KT 2440, belonging to the

402

family Pseudomonadaceae identified here on the card surfaces, are able to intracellularly


Page 16 of 32

Page 17 of 32


403

hydrolyse PHA in a wide range of hydroxyalkanoic acids and thus may indirectly contribute

404

to the polymer deterioration.45-47

405

Altogether, our results do not suggest that some of the eight polymers studied are non-

406

degradable per se, but rather that they differ for their ability to support a high cell-density

407

biomass from the surrounding environment, with effective deterioration capacities. Indeed, if

408

all the polymers studied in this work recruit potent degraders on their surface, we could notice

409

that the deterioration potential for a given polymer could be related with the amounts of

410

fungal and bacterial biomass. The results suggest a cell-density dependent polymer

411

degradation process. Another important bottleneck in polymer degradation is very likely the

412

environmental conditions, which have to be optimal to initiate the first cleavage steps that are

413

required to generate the pre-substrates compatible with their further assimilation by the

414

recruited microorganisms. Such conditions may be quite difficult to achieve when the samples

415

are buried in the soil. In this context, microorganisms may also preferentially degrade simpler

416

carbon sources that are available from the environment, rather than initiate a switching of

417

their metabolism to access the more complex carbon-sources offered by the polymer cards.

418

In the present study, under uncontrolled composting conditions, the property of

419

biodegradability was not confirmed for PHA, PBAT, PBS and PLLA, suggesting a

420

discrepancy with environmental marketing claims. This is explained by the test methods used

421

to determine the degree and rate of aerobic biodegradation of plastic materials, which are

422

based on exposure to a controlled-composting environment under laboratory conditions, at

423

thermophilic temperatures.48 Clearly, most of the plastic packaging that claims to be

424

“biodegradable” will only weakly or partially deteriorate under conditions typical of most

425

home-compost piles and will probably not be degraded at all in natural soil environments, in

426

which they often end up.

427

An important conclusion of the present study is also that the surface properties of the



428

polymers may strongly influence the formation and the microbial composition of the biofilms.

429

An intriguing observation was that, among the various surface properties tested in this study,

430

crystallinity appeared to be poorly correlated with biofilm formation. Indeed, intuitively, one

431

may suspect that microorganisms may more easily adhere and consume a less organised (i.e.

432

less crystalline or more amorphous) material. Accordingly, previous data showed that

433

enzymes indeed mainly attack the amorphous domains of a polymer.49-52 A clear

434

counterexample here was PBS, which displayed the highest crystallinity index, but recruited

435

biofilms very efficiently and was the second most degraded polymer under our experimental

436

conditions.

437

By contrast to crystallinity, a high surface energy, in conjunction with a high surface

438

roughness, may represent a driving parameter for biofilm formation. Obviously, a hydrophilic

439

surface, as observed for PHA cards, is prone to favour microbial adherence, as previously

440

reported.53-54 However, the present results also showed that relatively hydrophobic polymeric

441

surfaces, as observed with the PBAT and PBS cards, are also efficiently colonized and

442

degraded by surrounding microorganisms. A common parameter to the most degradable

443

polymers of this study (namely, PHA, PBS, PBAT and PA66) clearly appeared to be their

444

surface micro-structures: they all displayed a quite rough surface as compared to the other

445

materials. These micron-sized irregularities provide sites for microbial attachment, due to

446

their dimensions similar to those of bacteria, and thus favour microorganisms-polymer

447

contacts that may accelerate polymer degradation by the microbial community from the

448

compost. Therefore, the present results tend to confirm that the surface wettability (water

449

contact angle) could be a less important parameter for microbial attachment to polymer

450

surface than the surface roughness, as also suggested in the literature.55 However, a set of

451

material properties may act synergistically to sustain optimal biofilm formation and therefore

452

facilitate further degradation, as is the case of PHA. Altogether, our study points at the


Page 18 of 32

Page 19 of 32


453

modulation of surface properties as a crucial determinant to consider in order to enhance the

454

recruitment of potent microbial polymer degraders from the surrounding environment. In this

455

context, surface irregularities and wettability appear as the most promising parameters to

456

target. Future studies, aiming at evaluating the impacts of these properties on polymer

457

degradation in soil (with a main focus on the most persistent polymers) will undoubtfully

458

bring new perspectives for a better control of the fate of plastics in the environment and for a

459

significant reduction of their detrimental impacts on living organisms.

460 461

ASSOCIATED CONTENT

462

Supporting Information

463

Supporting Information1 (SI1, Fig. S1-S3 and Table S1) includes data on surface properties of

464

the polymer cards, SI2 provides a detailed experimental procedure, SI3 (Table S1 and Fig. S1-

465

S5) provides results on fungal (Part1) and bacterial (Part2) communities colonizing the

466

surface of the compost-incubated polymer cards and SI4 (Tables S1-S6) provides the relative

467

abundances of the microbial taxonomic groups colonizing the card surface.

468

This material is available free of charge via the Internet at http://pubs.acs.org.

469 470

AUTHOR INFORMATION

471

Corresponding author

472

*Thierry Ferreira, Laboratoire coopératif ThanaplastSP- Carbios Bioplastics - Ecologie et

473

Biologie des Interactions, Centre National de la Recherche Scientifique, UMR 7267,

474

Université de Poitiers, UFR SFA, Pôle Biologie Santé, 1 rue Georges Bonnet, bât B37, 86073

475

Poitiers, cedex 9, France, E-mail address: [email protected], phone: +33 (5) 49

476

45 40 04; fax:+33(5) 49 45 40 14

477

Notes



478

The authors declare no competing financial interest.

479 480

ACKNOWLEDGMENTS

481

Authors are grateful to the laboratory of Ecology and Biology of Interactions (University of

482

Poitiers) and Carbios (Biopôle Clermont-Limagne) for collaboration and support. We want to

483

acknowledge the staff of Parks Department at the University of Poitiers, E. Leroy (Valagro-

484

Carbone Renouvelable, Poitiers) and E. Beere (ImageUP platform, Poitiers).

485

Authors are sincerely thankful for the scientific support and helpful discussions provided by

486

S. Terrat and the technical recommandations from M. Lelièvre and V. Novak (GenoSol

487

platform, INRA Dijon, France, www2.dijon.inra.fr/plateforme_genosol/) for the development

488

of the pyrosequencing. A. M. dedicates this work to J. Lesobre.

489 490

References

491

(1) Andrady, A. L.; Neal, M. A. Applications and societal benefits of plastics. Phil. Trans. R.

492

Soc. B. 2009, 364 (1526), 1977−1984

493

(2) Barnes D. K. A.; Galgani, F.; Thompson R. C.; Barlaz, M. Accumulation and

494

fragmentation of plastic debris in global environments. Philos. Trans. R. Soc. B: Biol. Sci.

495

2009, 364 (1526), 1985−1998.

496

(3) Law, K. L.; Morét-Ferguson, S.; Maximenko, N. A.; Proskurowski, G.; Peacock, E. E.;

497

Hafner, J.; Reddy, C. M. Plastic accumulation in the North Atlantic Subtropical Gyre. Science

498

2010, 329, 1185−1188.

499

(4) Lambert, S.; Sinclair, C.; Boxall, A. Occurrence, degradation, and effect of polymer-based

500

materials in the environment. Rev. Environ. Contam. Toxicol. 2014, 227, 1−53.

501

(5) Teuten, E. L.; Saquing, J. M.; Knappe, D. R.; Barlaz, M. A.; Jonsson, S.; Björn, A. et al.

502

Transport and release of chemicals from plastics to the environment and to wildlife. Philos.


Page 20 of 32

Page 21 of 32


503

Trans. R. Soc. Lond. B Biol. Sci. 2009, 1526, 2027−2045.

504

(6) Boerger, C. M.; Lattin, G. L.; Moore, S. L.; Moore, C. J. Plastic ingestion by

505

planktivorous fishes in the North Pacific Central Gyre. Mar. Pollut. Bull. 2010, 60,

506

2275−2278.

507

(7) Singh B.; Sharma, N. Mechanistic implications of plastic degradation. Polym. Degrad.

508

Stabil. 2008, 93, 561–584.

509

(8) Lucas, N.; Bienaime, C.; Belloy, C.; Queneudec, M.; Silvestre, F.; Nava-Saucedo, J. E.

510

Polymer biodegradation: mechanisms and estimation techniques. Chemosphere 2008, 73 (4),

511

429−442.

512

(9) Arkakar, A.; Arutchelvi, J.; Bhaduri, S.; Uppara, P.V.; Doble, M. Degradation of untreated

513

and thermally pretreated polypropylene by soil consortia. Int. Biodeter. Biodegrad. 2009, 63

514

(1), 106−111.

515

(10) Dussud, C.; Ghiglione, J. F. Bacterial degradation of synthetic plastics. CIESM

516

Workshop Monograph n°46 on Marine Litters. In Marine litter in the Mediterranean and

517

Black Seas, Briand, F. (ed) 180 p., Monaco, CIESM Publisher, 2014, pp. 43 – 48.

518

(11) Artham, T.; Doble, M. Biodegradation of aliphatic and aromatic polycarbonates.

519

Macromol. Biosci. 2008, 8 (1), 14−24.

520

(12) Albertsson, A. C.; Karlsson, S. Aspects of biodeterioration of inert and degradable

521

polymers. Int. Biodeter. Biodegrad. 1993, 31 (3), 161−170.

522

(13) Lugauskas, A.; Levinskait, L.; Peciulyte, D. Micromycetes as deterioration agents of

523

polymeric materials. Int. Biodeterior. Biodegradation 2003, 52 (4), 233−242.

524

(14) Kaczmarek, H.; Bajer, K.; Galka, P.; Kotnowska, B. Photodegradation studies of novel

525

biodegradable blends based on poly(ethylene oxide) and pectin. Polym. Degrad. Stabil. 2007,

526

92 (11), 2058−2069.



527

(15) Fotopoulou, K. N.; Vakros, J.; Karapanagioti, H. K. Surface properties of marine

528

microplasctics that affect their interaction with pollutants and microbes. CIESM Workshop

529

Monograph n°46 on Marine Litters. In Marine litter in the Mediterranean and Black Seas,

530

Briand, F. (ed) 180 p., Monaco, CIESM Publisher, 2014, pp. 55 – 60.

531

(16) Keswani, A; Oliver, D. M.; Gutierrez, T.; Quilliam, R. S. Microbial hitchhickers on

532

marine plastic debris: human exposure risks at bathing waters and beach environments. Mar.

533

Env. Res. 2016, 118, 10−19.

534

(17) Zarfla, C.; Matthies, M. Are marine plastic particles transport vectors for organic

535

pollutants to the Arctic? Mar. Pollut. Bull. 2010, 60 (10), 1810−1814.

536

(18) Zettler, E. R.; Mincer, T. J.; Amaral-Zettler, L.A. Life in the "plastisphere": microbial

537

communities on plastic marine debris. Environ. Sci. Technol. 2013, 47 (13), 7137−46.

538

(19) De Tender, C. A.; Devriese, L. I.; Haegeman, A.; Maes, S.; Ruttink, T.; Dawyndt, P.

539

Bacterial community profiling of plastic litter in the Belgian part of the North Sea. Environ.

540

Sci. Technol. 2015, 49, 9629−9638.

541

(20) ISO Standard method 11063. Soil quality - Method to directly extract DNA from soil

542

samples. 2010, 20 pp.

543

(21) Terrat, S.; Plassart, P.; Bourgeois, E.; Ferreira, S.; Dequiedt, S.; Adele-Dit-De-

544

Renseville, N. et al. Meta-barcoded evaluation of the ISO standard 11063 DNA extraction

545

procedure to characterize soil bacterial and fungal community diversity and composition.

546

Microb. Biotechnol. 2015, 8 (1), 131–142.

547

(22) Chemidlin Prévost-Bouré, N.; Christen, R.; Dequiedt, S.; Mougel, C.; Lelièvre, M.;

548

Jolivet, C.; Shahbazkia, H. R.; Guillouet, L. et al. Validation and application of a PCR primer

549

set to quantify fungal communities in the soil environment by real-time quantitative PCR.

550

PLoS One 2011, 6 (9), e24166.

551

(23) R Development Core Team. R: A language and environment for statistical computing. R


Page 22 of 32

Page 23 of 32


552

Foundation for Statistical Computing, Vienna, Austria. ISBN 3-900051-07-0, URL.

553

http://www.R-project.org, 2011. Accessed 10 December 2015.

554

(24) Hulsen, T.; de Vlieg, J.; Alkema, W. BioVenn - a web application for the comparison

555

and visualization of biological lists using area-proportional Venn diagrams. BMC Genomics

556

2008, 9, 488.

557

(25) Oksanen, J.; Blanchet, F. G.; Kindt, R.; Legendre, P. ; O'Hara, R. B. ; Simpson, G. L. et

558

al. vegan: community ecology package. R package version 2.2-1. http://cran.r-

559

project.org/packages=vegan, 2015. Accessed 10 December 2015.

560

(26) Karlsson, S.; Albertsson, A. C. Biodegradable polymers and environmental interaction.

561

Polym. Eng. Sci. 1998, 38 (8), 1251−1253.

562

(27) Dequiedt, S.; Saby, N. P. A.; Lelievre, M.; Jolivet, C.; Thioulouse, J.; Toutain, B.;

563

Arrouays, D.; Bispo, A. et al. Biogeographical patterns of soil molecular microbial biomass as

564

influenced by soil characteristics and management. Global Ecol. Biogeogr. 2011, 20 (4), 641–

565

652.

566

(28) Harrison, J. P.; Sapp, M.; Schratzberger, M.; Osborn, A. M. Interactions between

567

microorganisms and marine microplastics: a call for research. Mar. Technol. Soc. J. 2011, 45,

568

12−20.

569

(29) Harrison, J. P.; Schratzberger, M.; Sapp, M.; Osborn, A. M. Rapid bacterial colonization

570

of low-density polyethylene microplastics in coastal sediment microcosms. BMC Microbiol.

571

2014, 14, 232.

572

(30) Carson, H. S.; Nerheim, M. S.; Carroll, K. A.; Eriksen, M. The plastic-associated

573

microorganisms of the North Pacific Gyre. Mar. Pollut. Bull. 2013, 75, 126−132.

574

(31) Oberbeckmann, S.; Loeder, M.G.; Gerdts, G.; Osborn, A.M. Spatial and seasonal

575

variation in diversity and structure of microbial biofilms on marine plastics in Northern

576

European waters. FEMS Microbiol. Ecol. 2014, 90 (2), 478−92.



577

(32) Amaral-Zettler, L. A.; Zettler, E. R.; Slikas, B.; Boyd, G. D.; Melvin, D. W.; Morrall, C.

578

E.; Proskurowski, G.; Mincer, T. J. The biogeography of the Plastisphere: implications for

579

policy. Front. Ecol. Environ. 2015, 13 (10), 541−546.

580

(33) Jurelevicius, D.; Alvarez, V.M.; Marques, J.M.; Lima, L.; Dias, F.D.; Seldin, L. Bacterial

581

community response to petroleum hydrocarbon amendments in freshwater, marine, and

582

hypersaline water-containing microcosms. Appl. Environ. Microbiol. 2013, 79, 5927−5935.

583

(34) Matsui, T.; Yamamoto, T.; Shinzato, N.; Mitsuta, T.; Nakano, K.; Namihira, T.

584

Degradation of oil tank sludge using long-chain alkane-degrading bacteria. Ann. Microbiol.

585

2014, 64, 391−395.

586

(35) Røberg, S.; Østerhus, J.; Landfald, B. Dynamics of bacterial community exposed to

587

hydrocarbons and oleophilic fertilizer in high-Arctic intertidal beach. Polar Biol. 2011, 34,

588

1455−1465.

589

(36) Lee, J. W.; Nam, J. H.; Kim, Y. H.; Lee, K. H.; Lee, D. H. Bacterial communities in the

590

initial stage of marine biofilm formation on artificial surfaces. J. Microbiol. 2008, 46 (2),

591

174−182.

592

(37) Elifantz, H.; Horn, G.; Ayon, M.; Cohen, Y.; Minz, D. Rhodobacteraceae are the key

593

members of the microbial community of the initial biofilm formed in Eastern Mediterranean

594

coastal seawater. FEMS Microbiol. Ecol. 2013, 85 (2), 348−357.

595

(38) Leahy, J. G.; Colwell, R. R. Microbial degradation of hydrocarbons in the environment.

596

Microbiol. Rev. 1990, 54 (3), 305–315.

597

(39) April, T. M.; Foght, J. M.; Currah, R. S. Hydrocarbon-degrading filamentous fungi

598

isolated from flare pit soils in Northern and Western Canada. Can. J. Microbiol. 2000, 46 (1),

599

38−49.

600

(40) Russell, J. R.; Huang, J.; Anand, P.; Kucera, K.; Sandoval, A. G.; Dantzler, K. W.;

601

Hickman, D.; Jee, J. et al. Biodegradation of polyester polyurethane by endophytic fungi.


Page 24 of 32

Page 25 of 32


602

Appl. Environ. Microbiol. 2011, 77 (17), 6076–6084.

603

(41) Wick, L. Y.; Furuno, S.; Harms, H. Fungi as transport vectors for contaminants and

604

contaminant-degrading bacteria, in Handbook of Hydrocarbon and Lipid Microbiology,

605

Timmis, K., McGenity, T., van der Meer, J.R., and de Lorenzo, V. (eds) New York, NY:

606

Springer-Verlag Berlin Heidelberg, 2010, pp. 1555–1561.

607

(42) Ferrari, B. C.; Zhang, C.; van Dorst, J. Recovering greater fungal diversity from pristine

608

and diesel fuel contaminated sub-antarctic soil through cultivation using both a high and a low

609

nutrient media approach. Front Microbiol. 2011, 2, 217.

610

(43) Mayrberger, J. M. Studies of genera cytophaga-flavobacterium in context of the soil

611

carbon cycle. Dissertation, Michigan state University, pp 121, 2011.

612

(44) Drury, B.; Rosi-Marshall, E.; Kelly, J. J. Wastewater treatment effluent reduces the

613

abundance and diversity of benthic bacterial communities in urban and suburban rivers. Appl.

614

Environ. Microbiol. 2013, 79, 1897−1905.

615

(45) Shah, A. A.; Hasan, F.; Hameed, A.; Ahmed, S. Biological degradation of plastics: a

616

comprehensive review. Biotechnol. Adv. 2008, 26 (3), 246−265.

617

(46) Steinbüchel, A.; Valentin, H. Diversity of bacterial polyhydroxyalkanoic acids. FEMS

618

Microbiol. Lett. 1995, 128, 219–228.

619

(47) Prieto, A.; Escapa, I.F.; Martinez, V.; Dinjaski, N.; Herencias, C.; de la Peña, F. et al. A

620

holistic view of polyhydroxyalkanoate metabolism in Pseudomonas pudita. Environ.

621

Microbiol. 2016, 18 (2), 341−357.

622

(48) ASTM D5338 – 98. Standard test method for determining aerobic biodegradation of

623

plastic materials under controlled composting conditions. 2003, 6 pp.

624

(49) Tsuji, H.; Miyauchi, S. Enzymatic hydrolysis of poly(lactide)s: effects of molecular

625

weight, L-lactide content, and enantiomeric and diastereoisomeric polymer blending.

626

Biomacromolecules 2001, 2 (2), 597−604.



627

(50) Tokiwa, Y.; Calabia, B. P.; Ugwu, C. U.; Aiba, S. Biodegradability of plastics. Int. J.

628

Mol. Sci. 2009, 10, 3722−3742.

629

(51) Yoshida, S.; Hiraga, K.; Takehana, T.; Taniguchi, I.; Yamaji, H.; Maeda, Y.; Toyohara,

630

K.; Miyamoto, K. et al. A bacterium that degrades and assimilates poly(ethylene

631

terephthalate). Science 2016, 351 (6278), 1196−1199.

632

(52) Bornscheuer, U. T. Feeding on plastic. Science 2016, 351 (6278), 1154−1155.

633

(53) Cunliffe, D.; Smart, C. A.; Alexander, C.; Vulfson, E. N. Bacterial adhesion at synthetic

634

surfaces. Appl. Environ. Microbiol. 1999, 65 (11), 4995–5002.

635

(54) Azeredo, J.; Oliviera, R. The role of hydrophobicity and exopolymers in initial adhesion

636

and biofilm formation. In Biofilms in Medecine, Industry and Environmental Biotechnology.

637

Lens, P., Moran, A. P., Mahony, T., Stoodley, P., and O’Flaherty, V. (eds). IWA Publishing,

638

2003, pp 16−31.

639

(55) Nauendorf, A.; Krause, S.; Bigalke, N. K.; Gorb, E. V.; Gorb, S. N.; Haeckel, M. et al.

640

Microbial colonization and degradation of polyethylene and biodegradable plastic bags in

641

temperate fine-grained organic-rich marine sediments. Mar. Poll. Bull. 2016, 103, 168−178.

642 643

Table legends

644

Table 1: Name, origin of the polymers used in this study and card properties (n = 1).

645

Table 2: Bacterial OTU richness and diversity indices from the compost and the different

646

polymers (mean ± standard error of the mean).

647 648 649

Figure captions

650

Fig. 1: Compost-incubated cards analysis. (A) Polymer card weight before and after

651

incubation in the compost (mean ± standard error of the mean). * indicates significant weight


Page 26 of 32

Page 27 of 32


652

loss (Mann-Whitney test, p < 0.05). The percentage of weight loss (mean ± standard error of

653

the mean) is given above the bars. (B) Amounts of DNA measured after direct extraction from

654

biofilms colonizing the compost-incubated cards (mean ± standard error of the mean).

655

Fig. 2: (A) Principal coordinate analysis (PCoA) of the bacterial communities associated with

656

the different polymer cards and the compost. Two-dimensional PCoA plot was based on the

657

Bray-Curtis distance matrix. Percentage of the diversity distribution explained by each axis is

658

indicated on the figure. (B) Three-set Venn diagram representing the unique and shared

659

bacterial OTUs at 95% identity (in bold) among polymers. Percentages indicate the

660

corresponding proportions of associated sequences.

661

Fig. 3: Polymer card properties (see Table 1) fitted onto bacterial community PCoA (see Fig.

662

2A) after pyrosequencing and taxonomic assignment by GnS-PIPE. Arrows represent the

663

contribution of each property to the discrimination of the bacterial diversity in contact with

664

polymer card surface. A long arrow shows a strong correlation. 95% confidence ellipses are

665

drawn for each plastic. Explained variations for the first five dimensions are indicated on the

666

figure (labels and bar plot).



Page 28 of 32

Table 1 Surface energy (mJ/m2) Crystallinity Polymer

Abbreviation Manufacturer Reference

dispersive

polar

(%)

total component

Polyolefin Ethylene-vinyl alcohol

EVOH

Kuraway

33.7

33.7

3.2

36.9

Poly(propylene)

PP

LyondellBasell Moplen HP 520H

36.9

31.8

0.3

32.1

Poly(butylene adipate terephthalate) PBAT

BASF

ecoflex® F Blend C1200 †, ‡

18.8

44.6

4.3

48.9

Poly(ethylene terephthalate)

PET

NaturePlast

PTI 001 ‡

9.8

40.2

5.4

45.6

Poly(butylene succinate)

PBS

NaturePlast

PBE 003 †, ‡

60.1

38.1

6.3

44.4

Poly(hydroxy alkanoate)

scl-PHA*

Mirel®

M4100 †, ‡

21.5

37.3

22.7

60.0

Poly(-L-lactide)

PLLA

NaturePlast

PLE 001 †, ‡

19.7

33.4

6.9

40.3

PA 66

DuPont™

Zytel® 101F NC010

29.8

37.8

3.8

41.6

Eval™ E105B

Polyester

Polyamide Poly(amide) 66

* short-chain-length PHA, † claimed as biodegradable polymer, ‡ bio-based ou partially bio-based polymer ACS Paragon Plus Environment

Page 29 of 32


Table 2

Number of OTU

Shannon

Evenness

507 (± 26)

5.27 (± 0.14)

0.85 (± 0.02)

442 (± 2)

4.90 (± 0.01)

0.80 (± 0.00)

511 (± 22)

5.37 (± 0.10)

0.86 (± 0.01)

289 (± 12)

3.96 (± 0.09)

0.70 (± 0.01)

PET

557 (± 7)

5.56 (± 0.02)

0.88 (± 0.00)

PBS

250 (± 6)

3.82 (± 0.10)

0.69 (± 0.02)

PHA

261 (± 13)

4.30 (± 0.05)

0.77 (± 0.00)

PLLA

459 (± 6)

4.99 (± 0.02)

0.81 (± 0.00)

483 (± 18)

5.04 (± 0.07)

0.82 (± 0.00)

Compost Polyolefin EVOH PP Polyester PBAT

Polyamide

PA 66

ACS Paragon Plus Environment


Page 30 of 32

A initial weight

weight after incubation in the compost

card weight (g)

10 9

5.5%

7.9%

(± 0.3%)

(± 0.7%)

*

8

1.1% (± 0.2%)

*

*

7 6 5 EVOH

PP

PBAT

PET

Polyolefin

PBS

PHA

PLLA

Polyester

PA66 Polyamide

B 300

ng DNA /cm2 of card

250 200 150 100 50 0 EVOH Polyolefin

PP

PBAT

PET

PBS Polyester

Fig. 1


PHA

PLLA

PA66 Polyamide

Page 31 of 32


PBS

B

A

214 OTUs 4923 sequences

26 OTUs 1.65%

38 OTUs PBS = 1.34% PET = 3.74%

150 OTUs 13.97%

PET


130 OTUs PHA = 87.53% PBS = 94.47% PET = 78.66%

20 OTUs PHA = 7.05% PBS = 2.54%

35 OTUs PHA = 1.97% PET = 3.63%

41 OTUs 3.45%

Sample Compost EVOH PA66 PET PP PBAT PBS PHA PLLA

PHA


Fig. 2A and 2B ACS Paragon Plus Environment


Fig. 3


Page 32 of 32

Fate of Eight Different Polymers under ... - ACS Publications

Recommend Documents