Temporal Dynamics of Bacterial and Fungal Colonization on Plastic

Surprisingly, none of the plastics exposed to offshore conditions displayed the typical signature of a late stage biofilm, suggesting that biofilm for...
0 downloads 0 Views 4MB Size
Subscriber access provided by Binghamton University | Libraries

Article

The temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea Caroline A. De Tender, Lisa Inès Devriese, Annelies Haegeman, Sara Maes, Jürgen Vangeyte, André Cattrijsse, Peter Dawyndt, and Tom Ruttink Environ. Sci. Technol., Just Accepted Manuscript • Publication Date (Web): 31 May 2017 Downloaded from http://pubs.acs.org on June 4, 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 31

Environmental Science & Technology

1

The temporal dynamics of bacterial and fungal colonization on plastic debris in the North Sea

2

Caroline De Tender1,2,*, Lisa I. Devriese1, Annelies Haegeman1, Sara Maes1, Jürgen Vangeyte1, André

3

Cattrijsse3, Peter Dawyndt2, Tom Ruttink1

4 5

1

6

9820 Merelbeke, Belgium

7

2

8

281 S9, 9000 Ghent, Belgium

9

3

Institute of Agricultural, Fisheries and Food Research (ILVO), Burgemeester Van Gansberghelaan 92,

Ghent University, Department of Applied Mathematics, Computer Sciences and Statistics, Krijgslaan

Flanders Marine Institute, InnovOcean site, Wandelaarkaai 7, 8400 Oostende, Belgium

10 11

*Corresponding author:

12

Caroline De Tender

13

Address:

14

Phone number: +3292722473

15

Fax:

16

Email address: [email protected]

Ankerstraat 1, 8400 Oostende, Belgium

+32 (0) 9 272 24 29

17 18 19

1 ACS Paragon Plus Environment

Environmental Science & Technology

Page 2 of 31

20

Abstract

21

Despite growing evidence that biofilm formation on plastic debris in the marine environment may be

22

essential for its biodegradation, the underlying processes have yet to be fully understood. Thus far,

23

bacterial biofilm formation had only been studied after short-term exposure or on floating plastic, yet a

24

prominent share of plastic litter accumulates on the seafloor. In this study, we explored the taxonomic

25

composition of bacterial and fungal communities on polyethylene plastic sheets and dolly ropes during

26

long-term exposure on the seafloor, both at a harbor and an offshore location in the Belgian part of the

27

North Sea. We reconstructed the sequence of events during biofilm formation on plastic in the semi-

28

enclosed harbor environment and identified a core bacteriome and subsets of bacterial indicator species

29

for early, intermediate, and late stages of biofilm formation. Additionally, by implementing ITS2

30

metabarcoding on plastic debris, we identified and characterized for the first time fungal genera on

31

plastic debris. Surprisingly, none of the plastics exposed to offshore conditions displayed the typical

32

signature of a late stage biofilm, suggesting that biofilm formation is severely hampered in the natural

33

environment where most plastic debris accumulates.

2 ACS Paragon Plus Environment

Page 3 of 31

Environmental Science & Technology

34

Introduction

35

Global production of plastics has reached an annual production of 322 million tons in 2015.1

36

Polyethylene (PE) is the most commonly produced plastic in the world and accounts for 64% of plastics

37

discarded shortly after use.2 A substantial share of plastics eventually ends up in the oceans and seas,

38

with an estimated 5.25 trillion plastic particles currently scattered throughout the oceans.3 In the Belgian

39

part of the North Sea, PE plastic sheets and dolly ropes are the dominant types of plastic debris (PD).4

40

The hard, hydrophobic surface of PD is an ideal environment for colonization by various biota, including

41

diatoms, bacteria, and invertebrate species.4,5,6,7,8 Merely one week exposure to the marine environment

42

is sufficient for a microbial biofilm to form on PD.9 Polymer type, environmental conditions and season

43

appear to affect the composition of the microbial biofilm.10,11 Furthermore, the bacterial community of

44

the biofilm significantly differs from that of the surrounding environment, e.g. seawater and sediment.4

45

Based on these unique characteristics, the assemblage of taxa inhabiting the outer surface of PD is often

46

referred to as the “plastisphere”.8

47

As for most thermoplastics, degradation of PE is extremely slow, and it is therefore expected to persist

48

in the marine ecosystem.12 Micro-organisms may contribute to the degradation of PD in the marine

49

environment. So far, only a few marine bacterial strains have been identified as potential PE degraders.

50

Arthrobacter sp. and Pseudomonas sp. were isolated from high-density PE (HDPE) debris from the Gulf

51

of Mannar, a marine coastal area.13 Furthermore, Kocuria palustris, Bacillus pumilis and Bacillus

52

subtilis strains were isolated from low-density PE debris originating from the Arabian Sea.14 Recently

53

it was shown that also a marine fungus, Zalerion maritimum, has the potential to actively degrade PE.15

54

Furthermore, several microbial strains, including bacteria and fungi, were isolated from PD in different

55

types of soil environment, and described as potential PE degraders.16 Most studies on PE degradation

56

are based on growth on medium with plastic polymers as the sole carbon source, PE mass loss and size

57

reduction, and the screening of changes in functional groups by FT-IR. Only few studies described the

58

actual degradation, e.g. based on enzyme production by the bacterial strain.17

59

The formation of a biofilm, a structured system which facilitates metabolic interaction between cells,18

60

can be important in terms of biodegradation. Biofilm formation can increase degradation efficiency of

61

pollutants such as diesel oil, and destruction of the biofilm architecture can disrupt interspecies 3 ACS Paragon Plus Environment

Environmental Science & Technology

Page 4 of 31

62

cooperation and interfere with degradation efficiency.19,20 To date, little is known both about the

63

temporal dynamics of colonization and biofilm formation on PD and about the microbial interactions

64

underlying the resulting biodegradation process. Previous studies on biofilm formation on plastics under

65

controlled conditions focused on short-term processes only (< 8 weeks),9,10,21,22,23 or did not include

66

taxonomic classification of the bacterial communities.24,25,26,27,28 Most of those studies focused on

67

floating plastic litter, whereas the predominant part of PD is located on the seafloor.29,30 Reconstructing

68

the temporal dynamics of microbial colonization based on randomly sampled PD proved difficult by

69

comparing independent pieces of plastic with unknown history, travel pattern, and duration of exposure

70

to the marine environment to each other.4

71

We established a long-term exposure time-series experiment in which two types of PE were exposed to

72

the Belgian part of the North Sea at two different locations. First, plastics were exposed and sampled at

73

the harbor of Ostend, a semi-enclosed environment, with low influence of currents. The presence of

74

anthropogenic activity, e.g. waste-pipes, land run-off and oil discharges, and the small median grain size

75

of the sediment makes this environment more susceptible to environmental pollution.31 Second, plastics

76

were exposed and sampled at the Thornton windmill park, which we will further refer to as the

77

“offshore” environment. In this area, currents are stronger, but pollution is less pronounced.

78

The aims of this research are fourfold. First, an in-depth study of the biofilm formed on plastic exposed

79

to the marine environment on fixed locations was done. Previous research focused on bacterial

80

communities,4,8,10,21,23,27 whereas the fungal community has been less intensively studied.23 Therefore,

81

we used 16S rDNA and ITS2 metabarcoding in parallel to study the taxonomic composition of bacterial

82

and fungal communities, respectively. To study possible factors that affect biofilm formation, we

83

compared two types of plastic (sheet or dolly rope) in two different environments (harbor or offshore).

84

Second, the temporal dynamics of bacterial and fungal colonization of PD are reconstructed, and we

85

identified signature species for early, intermediate or late phases of long-term exposure in the harbor

86

environment. This series of microbial colonization in the harbor was used to evaluate biofilm formation

87

stages in the offshore samples of the exposure series. In addition, these were compared to bacterial

88

communities of previously described4 randomly collected samples exposed to similar offshore

89

conditions. Third, possible sources of microorganisms were studied by comparing taxonomic profiles 4 ACS Paragon Plus Environment

Page 5 of 31

Environmental Science & Technology

90

of plastic to those of sediment and seawater. Fourth, we investigated if bacterial and fungal species

91

previously identified as potential PE degraders were also present in the biofilm to thus assess whether

92

microbial biodegradation in the marine environment may take place.

93 94

Materials and Methods

95

EXPERIMENTAL SETUP AND SAMPLE COLLECTION

96

From September 2015 until July 2016, PE samples were exposed to the marine environment at two

97

different locations in the Belgian part of the North Sea: at the harbor of Ostend (51°13’N, 2°56’E) and

98

offshore, at the Thornton windmill park (51° 34’N, 2° 58’ E) (Supplementary (S) Figure S1). These

99

locations are characterized by different features (Table S1). Environmental properties (seawater

100

temperature, conductivity, pH, oxygen, salinity and density) were measured using the CTD SBE-19plus

101

on each sampling date. Sediment organic matter or total organic carbon (TOC) of the upper sediment

102

layer (0-5 cm) was measured at the first sampling date for both locations, using the “dichromate method”

103

as previously described.32 In addition, concentrations of pollutants in the sediment were compiled from

104

previous studies and are listed in Table S1.33,34

105

Two types of PE, with different color and shape, were exposed to the two environments: transparent

106

plastic sheets (A4 size) (RKW Hyplast, Hoogstraten, Belgium) and orange-colored dolly ropes (ø 1 cm,

107

length 20 cm; ø single monofilament 1 mm). Three pieces of each type of plastic, representing three

108

biological replicates, were attached to a wooden block that sinks to the seafloor, and which was secured

109

in a construction (total length: +/- 60 m, weight +/- 75 kg) comprising a buoy, ropes (ø 16 mm), chains,

110

an anchor and concrete weights (Figure S2). At the start of the experiment, sets of thirteen and five

111

identical constructions were placed on the seafloor in the harbor and offshore, respectively. Handling of

112

these constructions was always done with plastic gloves to avoid contamination. In addition, from three

113

randomly chosen constructions a piece of PE sheet and dolly rope were cut with sterilized scissors to

114

study the microbial load of plastics at the onset of the experiment, using metabarcoding. This was done

115

for both locations.

116

At the harbor, one construction per week was pulled up and removed during the first month, and from

117

then on one construction per month. This led to thirteen collection dates: 1 (September 2015), 2, 3, 4, 9, 5 ACS Paragon Plus Environment

Environmental Science & Technology

Page 6 of 31

118

14, 18, 22, 27, 31, 35, 40 and 44 (July 2016) weeks after placing the constructions. Offshore, the

119

constructions were pulled up and removed on four collection dates: 4 (October 2015), 14, 18 and 22

120

weeks (February 2016) after placing them. Upon collection of a sample, which was done with sterile

121

forceps, scissors and gloves, half of the plastic was immediately stored at -80 °C for DNA extraction

122

and the other half was air-dried and stored at room temperature for the biofilm assay. Offshore, three

123

replicate seawater and sediment samples were collected on the same date and location as the

124

constructions were sampled, as described in a previous study.4 Per replicate, 1 l seawater was filtered

125

through a 0.22 μm Millipore membrane filter (Merck Millipore, Billerica, MA). After collection,

126

sediment samples and the membrane filters were stored at -80 °C until further use.

127 128

BIOFILM ASSAY

129

The quantitative biofilm assay was used to measure biofilm formation on the plastic sheets.9 Briefly,

130

plastic samples (4x5 cm, n=2 per time point x location x replicate, resulting in n=6 per time point x

131

location) were rinsed three times with sterile water and air-dried for at least 45 min in sterile Petri dishes.

132

These plastics were stained with crystal violet (1% w/v) for 45 min and washed three times with sterile

133

seawater. Stained samples were air-dried for another 45 min, cut into four pieces of similar size and

134

placed into a 2 ml Eppendorf tube to which 1 ml ethanol (95% v/v) was added. The ethanol was then

135

diluted 100-fold in ethanol and transferred to a cuvette to measure the optical density at 595 nm using

136

an UV-VIS spectrophotometer (UV-1700 Pharmaspec, Shimadzu, Brussel, Belgium). The optical

137

density is directly proportional to the amount of biofilm per surface area on the plastic.

138 139

DNA EXTRACTION

140

DNA was extracted from the sediment and plastic samples using the Powersoil DNA isolation kit

141

(MOBIO Laboratories, Carlsbad, CA) according to the manufacturer’s instructions. In total, 250 mg of

142

sediment, a piece of 2 cm by 2.5 cm (total surface area of ± 10 cm2) of the plastic sheet, or 10 individual

143

monofilaments with a length of 2.5 cm of the dolly rope (surface area of ± 10 cm2) were used for DNA

144

extraction. Before extraction, plastic was rinsed three times with sterile water to remove sediment

145

particles and loosely attached organisms. DNA was extracted from the Millipore filters containing the 6 ACS Paragon Plus Environment

Page 7 of 31

Environmental Science & Technology

146

micro-organisms of seawater as described in a previous study.4 The DNA extracts of all samples were

147

stored at -20 °C until further use for amplicon sequencing.

148 149

16S rDNA AND ITS2 AMPLICON SEQUENCING AND SEQUENCE READ PROCESSING

150

Amplicon sequencing of the V3-V4 fragment of the 16S rRNA gene and the ITS2 gene fragment using

151

Illumina technology (Illumina, San Diego, CA, USA) was done to study both the bacterial and fungal

152

communities on PD. DNA fragments were amplified and extended with Illumina specific index adaptors

153

using an amplification PCR followed by a dual-index PCR, as previously described.32 Detailed

154

information on the library preparation can be found in the supplementary materials. Each PCR reaction

155

product was purified using the CleanPCR reagent kit (MAGBIO, Gaithersburg, MD, USA). Libraries

156

were quality-controlled using the Qiaxcel Advanced, with the Qiaxcel DNA High Resolution kit

157

(QIAGEN, Germantown, MD, USA), and concentrations were measured using the Quantus double-

158

stranded DNA assay (Promega, Madison, WI, USA). The indexed libraries of each sample were diluted

159

to 10 nM and pooled in a 2:1 ratio for bacterial and fungal libraries, respectively. Resulting libraries

160

were sequenced using Illumina MiSeq v3 technology (2 x 300 bp) by Macrogen, South-Korea, using

161

30% PhiX DNA as spike-in.

162

Demultiplexing of the amplicon dataset and removal of the barcodes was done by the sequencing

163

provider. The raw sequence data is available in the NCBI Sequence Read Archive under the accession

164

number Bioproject ID SUB2179884 for the bacterial sequences and SRP094681 for the fungal

165

sequences. Processing the sequence reads to Operational Taxonomic Unit (OTU) tables was done as

166

previously described 32 and described in detail in the supplementary materials.

167

The bacterial load of three PE sheets and three ropes was checked at the onset of the experiment before

168

exposure to the marine environment. Samples were analyzed by metabarcoding in parallel to all other

169

samples. Because the PCR product concentration was low, all DNA was used for sequencing. For each

170

sample, the number of reads was < 100.

171 172

DOWNSTREAM DATA ANALYSIS AND STATISTICS

7 ACS Paragon Plus Environment

Environmental Science & Technology

Page 8 of 31

173

OTU tables of the 16S V3-V4 and ITS2 amplicon sequencing were analyzed using the QIIME software

174

package (v1.9.0).35 Taxonomy was assigned with the script “assign_taxonomy.py” using the Uclust

175

method considering maximum 3 database hits, with the Silva v119 97% rep set (as provided by QIIME)

176

as reference for the bacterial sequences and UNITE v7 (dynamic) for fungal sequences.36,37,38

177

A part of the fungal sequences could not be classified using the UNITE database. These sequences were

178

extracted from the total data set and their taxonomy was assigned using BLAST for sequence

179

comparison with the non-redundant nucleotide database of NCBI.39 We kept the best hit per query using

180

an e-value cut-off of 1e-5 and a minimal percent identity of 98.5%.

181

Rarefaction analysis was done using the “alpha_rarefaction.py” script of QIIME. A plateau was reached

182

at 10,000 sequences for the bacterial and fungal OTUs (Figure S3 and Figure S4). Richness of the

183

bacteria and fungi was determined on rarefied data, for which the number of sequences and the

184

calculation of the Chao1 richness index was set on the plateau.

185

A core microbiome was calculated separately for the bacterial and fungal communities. Only plastics

186

exposed for at least four weeks were considered to calculate a core microbiome to account for some lag-

187

time for biofilm build-up. This core microbiome was calculated separately for the two environments.

188

OTUs were denoted as core organisms if their relative abundance contributed at least 0.1% to the total

189

community of a sample in at least 90% of the plastic items per environment. Calculations for the core

190

microbiome were done in R.40 In addition, the genera that were most abundant at the last time point of

191

sampling were calculated by selecting those genera with a minimal mean abundance of 1% over the

192

replicates for sheets or dolly ropes.

193

The multivariate analysis was done using the R package vegan (version 2.0-10).41 The OTU tables of

194

bacterial and fungal sequences, as generated by Usearch, were normalized by calculating relative

195

abundances. Next, OTUs with a low count number were removed by only retaining the OTUs which

196

had a minimal relative abundance of 0.01 % in at least three samples. The dissimilarity matrix, based on

197

the Bray-Curtis dissimilarity index, was calculated from this normalized and filtered OTU table, for both

198

the bacterial and fungal sequences. The homogeneity of the variances was checked on this dissimilarity

199

matrix using the betadisper function. The significance of the factors environment, type of plastic, and

200

time, and their various interaction effects were analyzed using PERMANOVA analysis (number of 8 ACS Paragon Plus Environment

Page 9 of 31

Environmental Science & Technology

201

permutations = 1,000) using the Bray-Curtis dissimilarity index matrix as input. Factors were considered

202

significantly different if p-value < 0.05.

203 204

Results

205

Bacterial and fungal colonization in the harbor environment

206

From the first week of exposure onwards, a coating comprising a microbial biofilm, sediment particles,

207

algae and macro-fouling (e.g. mussels) was formed on the plastic sheets at the harbor (Figure S5). Using

208

a quantitative biofilm formation assay,9 at least part of the coating on the plastic sheets could be

209

attributed to a microbial biofilm (Figure 1). This biofilm was already detected after one week of

210

exposure, and increased slightly until week 27, followed by a period of stronger growth until week 40.

211

The taxonomic composition of the bacterial and fungal community on plastic at the harbor was analyzed

212

in detail by 16S and ITS2 metabarcoding. Richness of the samples was studied by estimating the number

213

of observed OTUs and the Chao1 index. Both measures showed that the richness of sheets is similar to

214

that of dolly ropes; both for bacterial OTUs (Figure S6 and S7 - Harbor) and for fungal OTUs (Figure

215

S6 and S7 - Harbor). The number of bacterial OTUs is only slightly higher on dolly ropes compared to

216

sheets in the first few weeks of exposure to the harbor environment. At each time point, the bacterial

217

richness of plastic (mean around 1500 OTUs) was markedly higher than fungal richness (mean around

218

500 OTUs).

219

The bacterial community of plastic sheets and dolly ropes at the harbor displayed a gradual change in

220

taxonomic composition during the period of exposure (Figure 2A and 2B). This temporal gradient, more

221

evident on the plastic sheets, is, at least in part, caused by shifts in abundance of particular bacterial

222

classes: an increase in the relative abundance of alpha- and betaproteobacteria and flavobacteria, and a

223

decrease in the relative abundance of gammaproteobacteria (Figure 2A and 2B). Alpha- and

224

gammaproteobacteria are characteristic for primary biofilm colonization, while bacteroidetes are known

225

secondary biofilm colonizers in the marine environment.42,43,44 On the plastic sheets, a gradual decrease

226

in the relative abundance of these primary colonizers and an increase in secondary colonizers was

227

observed (Figure S8A), suggesting that subsequent time points reflect progressive stages of biofilm

9 ACS Paragon Plus Environment

Environmental Science & Technology

Page 10 of 31

228

formation. This shift from primary to secondary colonizers, however, was not as clearly discernible for

229

the dolly ropes (Figure S8B).

230

Next, we defined a core bacteriome of plastic samples (see Materials and Methods). In total, 25 bacterial

231

core OTUs were identified both on plastic sheets and dolly ropes (Table 1). Based on their temporal

232

profile, these core members were classified into four groups: (1) OTUs without a clear period of high

233

relative abundance (neutral), e.g. Arenicella, Methylotenera; (2) OTUs with higher abundance in the

234

beginning (early stage; week 1-14) of the exposure period, e.g. Sulfurovum, Maritimimonas; (3) OTUs

235

with higher abundance in the middle (intermediate; week 14-35) of the exposure period, e.g.

236

Robiginitomaculum, and (4) OTUs with highest abundance at the end (late stage; week 35-44) of the

237

exposure period, e.g. Sulfitobacter, Psychroserpens (Table 1; Figure S9; Figure S10).

238

Next, the fungal community on the plastic sheets and dolly ropes in the harbor was studied. Strikingly,

239

the majority of the fungal sequences (28% to 97% of the reads per sample) could not be assigned using

240

the UNITE database (Figure 3A and 3B). Using NCBI Blast, some of those reads could be assigned to

241

fungi, others to other members of the eukaryotes, e.g. Paramoeba permaquidensis, Paramoeba

242

aestuarina, Pleurobrachia pileus, Sugiura chengshanense, Sagartia elegans, and Rhizostoma pulmo,

243

but the vast majority remained unassigned (Table S2). Within the share of the fungal sequences that

244

were assigned to a certain taxonomy, the Ascomycota were highly abundant, followed by a smaller

245

fraction of Basidiomycota (Figure 3A and 3B). Zygomycota were also identified, but only represented

246

a minor fraction. In addition, genera that were highly abundant on sheets or dolly ropes ( > 1%) sampled

247

at the last time point (44 weeks) were studied (see Table S3). Especially members of the

248

Lecanoromycetes, e.g. Physconia, Candelariella, and Caloplaca were abundant. No clear temporal

249

profile characterized by early, intermediate, and late stage abundance peaks could be identified,

250

essentially because the fungal community profile varied considerably, even between successive time

251

points (Figure 3A and 3B). In addition, no core group of fungal organisms could be identified,

252

illustrating the variability of the fungal community through time.

253 254

Bacterial and fungal colonization in the offshore environment

10 ACS Paragon Plus Environment

Page 11 of 31

Environmental Science & Technology

255

Biofilm formation occurred on plastic exposed to offshore conditions, but was much less pronounced

256

compared to the harbor environment as described above. For instance, the biofilm layer was hardly

257

visible by the naked eye even after 22 weeks of exposure (Figure S5), and the amount of biofilm that

258

had accumulated after 22 weeks of exposure offshore was similar to the amount that had already

259

accumulated at the harbor after 1 week of exposure (Figure 1). Until week 18 of the exposure period,

260

the number of unique bacterial OTUs and the Chao1 index, both representing the richness of the samples,

261

on the plastic sheets sampled offshore remained low ( < 1000 OTUs; Chao1: