Identification of bisphenol-A assimilating microorganisms in mixed

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Environmental Processes

Identification of bisphenol-A assimilating microorganisms in mixed microbial communities using C-DNA stable isotope probing 13

Sandeep Sathyamoorthy, Catherine Hoar, and Kartik Chandran Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b01976 • Publication Date (Web): 24 Jul 2018 Downloaded from http://pubs.acs.org on July 26, 2018

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Environmental Science & Technology

Identification of bisphenol-A assimilating microorganisms in mixed microbial communities using 13C-DNA stable isotope probing Sandeep Sathyamoorthy‡, Catherine Hoar‡, Kartik Chandran*

Columbia University, Department of Earth and Environmental Engineering, 500 West 120th Street, Room 1045 Mudd Hall, New York, NY 10027. *corresponding author: [email protected]. ‡These authors contributed equally.

1 ACS Paragon Plus Environment


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1

ABSTRACT

2

A wide range of trace organic contaminants (TOrCs), including the endocrine disrupting

3

compound bisphenol-A (BPA), are subject to microbial transformations during biological

4

wastewater treatment.

5

capable of assimilating emerging contaminants.

6

(DNA-SIP) was used to investigate biodegradation and assimilation of BPA by mixed microbial

7

communities collected from two full-scale wastewater treatment plant bioreactors in New York

8

City and subsequently enriched under two BPA exposure conditions. The four enrichment modes

9

(two reactors with two initial BPA concentrations) resulted in four distinct communities with

10

different BPA degradation rates. Based on DNA-SIP, bacteria related to Sphingobium spp. were

11

dominant in the assimilation of BPA or its metabolites. Variovorax spp. and Pusillimonas spp.

12

also assimilated BPA or its metabolites. Our results highlight that microbial communities

13

originating from wastewater treatment facilities harbor the potential for addressing not only

14

human-derived carbon, but also BPA, a complex anthropogenic TOrC. While previous studies

15

focus on microbial biodegradation of BPA, this study uniquely determines the ‘active’ fraction of

16

microorganisms engaged in assimilation of BPA-derived carbon. Ultimately, information on

17

both biodegradation and assimilation can facilitate better design and operation of engineered

18

treatment processes to achieve BPA removal.

However, relatively little is known about the identity of organisms Here,

13

C-DNA stable isotope probing

19


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INTRODUCTION

21

The ecological impacts of trace organic contaminants (TOrCs) elicit growing concern,

22

particularly in light of research suggesting that chronic exposure to TOrCs may be deleterious to

23

the reproduction and development of aquatic species.1-3 Wastewater treatment plants (WWTPs)

24

are essential barriers for the influx of TOrCs into the environment. Although WWTPs are not

25

currently designed for attenuating TOrCs, reports suggest partial removal and transformation of a

26

wide range of TOrCs within the biological treatment process through both abiotic and biotic

27

reactions.4-6 However, relatively little is known about the identity of microbial species able to

28

assimilate TOrCs.

29

The overall objective of this study was to identify bacteria capable of assimilating the

30

endocrine disrupting compound bisphenol-A (BPA). The release of BPA into the environment is

31

primarily from industrial facilities and WWTPs.7 BPA mimics estrogen and has both agonist and

32

antagonist effects.8 In the environment, chronic exposure to BPA at concentrations as low as

33

10 µg/L can result in transcriptional level changes in the reproductive systems of certain fish

34

species.9 However, other studies have shown no effect of certain BPA exposure treatments on

35

other aquatic species.10 Furthermore, the effects of BPA on different species are varied and not

36

well understood, especially in relation to long-term exposure and implications of BPA mixed

37

with other chemicals.11 Despite its elimination from a wide range of consumer products and

38

industrial formulations, over one million pounds of BPA are released into the environment

39

annually.12 In 2014, BPA was included on the list of chemicals for assessment under the Toxic

40

Substances Control Act by the United States Environmental Protection Agency.12 The removal

41

of BPA in WWTPs is highly variable, ranging from 10% to >99%, with reported WWTP influent



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concentrations of BPA ranging from nondetectable to ~40 µg/L and effluent concentrations

43

ranging from nondetectable to ~20 µg/L.13

44

We evaluated BPA biodegradation and assimilation by two enriched microbial communities

45

originating from an urban WWTP in New York City through substrate consumption and DNA

46

stable isotope probing (DNA-SIP) assays, respectively. DNA-SIP relies on the incorporation of a

47

stable isotope (e.g.,

48

compound in batch substrate depletion assays.14, 15 Isopycnic ultracentrifugation is used to isolate

49

the labeled DNA, and bacteria that carry the

50

Generation Sequencing.16 A variety of SIP techniques have been used to study the

51

biodegradation of a wide range of contaminants, including TOrCs.17-19 In this study, we expand

52

the application of SIP to elucidate the assimilation of BPA or its metabolites by enriched

53

microbial communities originating from a full-scale urban WWTP. Future efforts to achieve or

54

enhance BPA removal could take advantage of the presence of these protagonists in WWTPs

55

through engineering strategies aimed at reconciling specific microbial community structure with

56

biodegradation or treatment function.

13

C or

15

N) into the DNA of bacteria capable of assimilating the labeled

13

C label are subsequently identified using Next

57 58

MATERIALS AND METHODS

59

BPA Degradation Experiments

60

Batch experiments were employed to evaluate the biodegradation and assimilation of BPA by

61

microbial communities initially originating from two separate biological treatment reactors at a

62

full-scale WWTP in New York City. One of these reactors is used for primary effluent treatment

63

(PET reactor) and the other is used to treat reject water from anaerobic digestion after

64

centrifugation (separate centrate treatment – SCT reactor). Samples were collected from aerobic


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zones of both reactors; a simplified process flow diagram of the reactors is shown in Figure S-1

66

(Supplementary Information). The PET and SCT reactors were selected in order to assess the

67

influence of two distinct wastewater treatment processes (Supplementary Information Table S-1)

68

on the microbial community structure and the resulting BPA biodegradation and assimilation

69

potential of these communities.

70

Enrichment of the microbial communities on BPA prior to conducting batch DNA-SIP

71

experiments allowed for the elucidation of microbes with a higher potential for BPA assimilation

72

than the parent activated sludge population along with an assessment of the effect of different

73

BPA exposure conditions on biodegradation. Preliminary experiments indicated that BPA was

74

slowly biodegraded by microbial communities from the PET and SCT reactors after a long lag

75

period (~50-80 h, Supplementary Information Section S-2), conditions that are not ideal or

76

recommended for DNA-SIP experiments.20,

77

conditions were applied to lab scale fed-batch reactors (SI, Section S-3). The first condition was

78

a high concentration (HC) exposure at a dose of 100 mg-BPA/L, which corresponded to the BPA

79

concentration used in the subsequent DNA-SIP experiments. The second condition was a low

80

concentration (LC) dose exposure at 500 µg/L. BPA was added to the fed-batch reactors using a

81

stock solution of BPA in methanol (600 mg-BPA/g and 7,000 µg-BPA/g for HC and LC

82

exposure treatments, respectively) upon >99% removal of BPA. We hypothesized that the

83

selected BPA concentration in the HC and LC-treatments would give rise to distinct diversity

84

and identity of BPA-assimilating microorganisms. Samples from high concentration exposure

85

treatments (PET-HC and SCT-HC) and low concentration exposure treatments (PET-LC and

86

SCT-LC) were collected for SIP experiments after 35 and 40 days, respectively. The resulting

21

During enrichment, two separate exposure



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total BPA loading through the HC and LC exposure treatments was 1,000 mg and 6 mg BPA,

88

respectively.

89 90

13

C-DNA Stable Isotope Probing

91

Biomass samples from each of the four treatments were centrifuged (3,500 xg, 15 min, 4°C

92

using a Beckman Coulter Avanti J-6 XPI centrifuge and JA 25.15 rotor) and the biomass pellets

93

were twice washed and re-suspended in a BPA-free medium for DNA-SIP experiments (see SI,

94

Section S-4 for starting biomass concentrations and Table S-2 for medium details). For each

95

treatment, batch experiments included duplicate

96

experiments. Inclusion of the 12C-BPA control samples allowed us to differentiate the fraction of

97

biomass capable of BPA assimilation from the fraction of the microbial consortium capable of

98

BPA biodegradation. The 12C-BPA SIP controls and

99

in open 20 mL glass scintillation vials (Kimble-Chase, Vineland, NJ) with a liquid volume of 5

100

mL. The vials were sampled at the time of >99.99% removal of BPA to maximize

101

biodegradation and assimilation of

102

metabolites, similar to the approach applied in previous studies, e.g. Baytshtok, et al.22 An

103

additional parallel time-course evaluation (TC) using

104

appropriate time at which to terminate the SIP experiments. The TC experiments were conducted

105

in 40 mL glass scintillation vials (VWR, Radnor, PA) with a liquid volume of 20 mL. A

106

no-biomass control (NB) was also included. All experiments were conducted in duplicate in an

107

orbital shaker (New Brunswick Scientific, Enfield, CT) operating at 260 RPM at 21 ± 1 oC. The

108

target initial BPA concentration in the DNA-SIP experiments, controls and the TC reactors was

109

100 mg/L. Control and TC reactors were spiked with unlabeled BPA (99+%, Sigma-Aldrich,

13

12

C-BPA controls and duplicate

13

13

C-BPA

C-BPA SIP experiments were conducted

C-BPA while minimizing cross-feeding of

12

13

C-BPA

C-BPA was included to determine the


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Saint Louis, MO), and experimental reactors were spiked with labeled Bisphenol-A-(diphenyl-

111

13

112

concentration relative to the typical BPA concentrations detected in wastewater influents was

113

utilized in the DNA-SIP experiments because the biomass yields for BPA for wastewater

114

microbial communities are unknown. Applying a high concentration of the labeled compound

115

ensures the availability of labeled

116

study of emerging contaminant degradation by enriched cultures.23

C12) (99 atom %

13

C, 98%, Sigma-Aldrich, ISOTEC INC, Miamisburg, OH). A high BPA

13

C-DNA, an approach that has been used previously in the

117 118

Analytical Methods

119

BPA was quantified using high performance liquid chromatography with UV detection and

120

mass spectrometry (MS). Separation was achieved using a 3µm analytical Acclaim PA2 column

121

(2.1 x 150 mm; Thermo Scientific) at 30 oC with an isocratic mobile phase consisting of

122

methanol (70 vol %) and water at 0.20 mL/min. The injection volume was 50 µL. Quantification

123

of BPA was based on UV detection at 230 nm (DAD-3000, Thermo Scientific), with a method

124

detection limit of 1.8 µg-BPA/L. Calibration standards were prepared using serial dilutions in

125

water of a stock solution of BPA dissolved in methanol. Identification of BPA was confirmed

126

using MS (MSQ Plus single quadrupole mass spectrometer, Thermo Scientific) using

127

atmospheric pressure charge ionization (APCI) in negative mode with a cone voltage of -65V,

128

source temperature of 550oC and nitrogen pressure of 50 psi.

129 130

DNA Extraction and Density Gradient Ultracentrifugation

131

DNA from biomass samples was extracted with the DNeasy Blood & Tissue Mini Kit protocol

132

using the Qiacube robotic workstation (Qiagen, Valencia, CA) following the manufacturer’s



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protocol. The DNA concentration in the extracts was measured using a NanoDrop lite UV

134

spectrophotometer (Thermo Scientific, Waltham, MA) and the extracted DNA was stored at -

135

20oC. Equal masses of DNA from the duplicate

136

achieve a mass of 5,000 ng DNA for utilization in ultracentrifugation of labeled DNA fractions.

137

This mass of DNA was sufficient for detection and quantification of separated DNA based on

138

preliminary optimization of ultracentrifugation (data not shown) and consideration of the

139

unknown adbundance of and resulting biomass yields for those organisms capable of

140

assimilating BPA or its metabolites. Equal masses of DNA from the 12C-BPA SIP controls were

141

similarly combined.

13

C-BPA SIP experiments were combined to

142

The gradient solution for ultracentrifugation was prepared in a 15 mL centrifuge tube

143

(Corning, Tewksbury, MA) following the method previously described, with minor

144

adjustments.22,

145

1,200 µl, to which 4,800 µl CsCl solution (64 wt%) was added. The density of the gradient

146

solution was determined using a refractometer (Reichert Model Brix/RI-Chek, Reichert

147

Industries, Depew, NY). The density was adjusted using either gradient buffer or CsCl solution

148

to 1.72 g/ml. The gradient solution was loaded into 5 mL polyalomer ultracentrifuge tubes

149

(Beckman Coulter, Jersey City, NJ) and the tubes were balanced to within 10 mg as

150

recommended. Ultracentrifugation (40,000 RPM or ~177,000 xgav, 20 oC, 69 h) was performed

151

at using a vTi 65.2 rotor in a L8-M ultracentrifuge (Beckman Coulter, Jersey City, NJ). Ten

152

gradient fractions (500 µL each) were removed from the ultracentrifuge tube using a syringe

153

pump and bromophenol blue solution.22, 24 The density of each fraction was determined using a

154

refractometer. DNA was recovered from each fraction using ethanol precipitation and

155

resuspended in 100 µL DNA-free water.

24

Gradient buffer24 was added to the combined DNA to a total volume of


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156

Quantitative real-time polymerase chain reaction (qPCR) was used to measure the abundance

157

of the total bacterial 16S rRNA gene in each fraction using EUB primers25 (forward primer

158

1055f (5′-ATGGCTGTCGTCAGCT-3′) and reverse primer 1392r (5′-ACGGGCGGTGTGTAC-3′),

159

Integrated DNA Technologies, Coralville, IA). qPCR analyses were conducted on an iQ5

160

real-time PCR thermal cycler (BioRad, Hercules, CA) using SYBR® green chemistry in a 25 uL

161

reaction mixture with the specific components of the reaction mixture as follows: mastermix 12.5

162

µL (iQ SYBR® Green Supermix), forward primer 1.0 µL, reverse primer 1.0 µL, 5 µL target

163

DNA and nuclease-free water 5.5 µL. Primer stock solutions used in the reactions were made at

164

a concentration of 5 µM. The reactions were performed in triplicate for each sample with a set of

165

triplicate standards and no-template controls included in each plate. The standards used in each

166

assay were generated through serial decimal dilutions of a stock standard of plasmid DNA with

167

the targeted 16S rRNA gene inserts. Absence of primer dimers was confirmed for each reaction

168

based on inspection of the melt-curves. Information for the qPCR standard curve and qPCR

169

results is included in SI Section S-5.

170 171

DNA Sequencing and Analysis of Sequencing Data

172

Ion Torrent sequencing was employed to identify BPA assimilating bacteria in each of the

173

microbial communities. Equal volumes of DNA from each of the independently obtained

174

duplicate heavy-DNA gradient fractions (density > 1.737 g/mL) from a given ultracentrifugation

175

tube were pooled to produce a single heavy-DNA sample. The light-DNA gradient fractions

176

(density ≤ 1.737 g/mL) from the ultracentrifugation tube were similarly pooled for each tube.

177

Therefore, four pooled DNA samples were sequenced from each of the four SIP experiments

178

(12C-SIP controls and

13

C-SIP experiments from PET and SCT reactors). These sixteen heavy



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and light pooled DNA samples, in addition to the DNA from the four microbial consortia at the

180

start of the SIP experiments and the PET and SCT reactors at the start of the exposure treatments,

181

were sequenced via amplicon sequencing of the 16S rRNA gene using the Ion Torrent Personal

182

Genome Machine (PGM) platform (Thermo Fisher Scientific).

183

Amplification for preparation of the sequencing libraries was carried out using the fusion

184

method

(Thermo

Fisher

Scientific)

with

the

forward

primer

1055f

185

(5′-ATGGCTGTCGTCAGCT-3′) and reverse primer 1392r (5′-ACGGGCGGTGTGTAC-3′) that

186

were linked to unique 6-nucleotide multiplex barcodes (Ion Xpress barcode adapters, Thermo

187

Fisher Scientific). The amplification consisted of 30 cycles of 94 °C for 30 s, 55 °C for 30 s, and

188

68 °C for 90 s. Amplification reactions were performed in a 25 µL reaction mixture with the

189

following components: mastermix 12.5 µL (iQ Sybrmix, BioRad, Hercules, CA), forward fusion

190

primer 1.0 µL, reverse fusion primer 1.0 µL, nuclease-free water 5.5 µL and template DNA

191

5.0 µL. The amplified DNA was purified using the Qiaquick PCR Purification protocol using the

192

Qiacube robotic workstation (Qiagen, Valencia, CA).

193

Sequencing libraries were quantified using the KAPA library quantification kit (KAPA

194

biosystems, Wilmington, MA) following the manufacturer’s protocol with minor modifications.

195

DNA libraries were diluted (1:2000) in nuclease-free water. The reactions were performed in a

196

20 µL reaction mixture with the following components: mastermix 10 µL (KAPA SYBR®

197

FAST qPCR Master Mix), forward/reverse primer mix 2 µL (10x Ion Torrent primer mix),

198

nuclease-free water 6 µL and diluted library DNA 2 µL. Sequencing libraries were each prepared

199

at a concentration of 100 pM and combined. A diluted library at a concentration of 8 pM was

200

subsequently enriched using the Ion PGM™ Template OT2 400 Kit (Thermo Fisher Scientific).


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The enriched samples were loaded onto a 316 Chip v2 and sequenced using the 400-bp

202

sequencing kit (Thermo Fisher Scientific).

203

Raw sequence reads from the Torrent Suite server were converted to fastq format and

204

processed using tools available in Mothur26 within the Galaxy platform.27, 28 Raw sequence reads

205

were trimmed using Mothur to remove those sequences with (i) a sequence length shorter than

206

280 or longer than 380 nt and (ii) an average quality score of 99.99% in all four SIP experiment batch incubation

218

reactors (PET-HC, SCT-HC, PET-LC and SCT-LC), over a course of 20 h, 36 h, 48 h and 58 h,

219

respectively (Figure 1). For each reactor, initial BPA-degradation was characterized by a lag

220

phase, followed by an approximately linear decrease in BPA concentration. Biomass-normalized

221

biodegradation coefficients for this period of linear degradation were estimated to be ~6 µg-

222

BPA/(mg-pCODh) for PET-HC, ~3 µg-BPA/(mg-pCODh) for SCT-HC, ~8 µg-BPA/(mg-

223

pCODh) for PET-LC, and ~4 µg-BPA/(mg-pCODh) for SCT-LC, (SI, Section S-7, Table S-5).



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Therefore, for the same exposure treatment, BPA biodegradation by the PET microbial

225

community occurred more rapidly than biodegradation by the SCT community, likely linked to

226

differences in structure and function of the two communities. Such differences may have

227

originated from different WWTP process conditions (e.g. diversity and availability of carbon

228

substrates) of the source biomass, which persisted through enrichment, though an assessment of

229

these factors was beyond the scope of this work. Nonetheless, BPA exposure indeed resulted in

230

distinct enriched community structures, as indicated by 16S rRNA gene-based phylogenetic

231

analysis (see details in the following section: Exposure Conditions Influence Microbial

232

Community Structure).

233

During all four experiments, a biodegradation metabolite was produced (as detected using

234

HPLC-UV-MS) and subsequently consumed. The UV spectrum of the metabolite exhibited a

235

single broad peak at 277 - 278 nm. Analysis of the mass spectrum identified dominant ions with

236

m/z ratios of 226 and 241 in negative-ion mode, and 149 and 243 in positive-ion mode,

237

respectively (SI, Section S-8). Positive identification of this and any other metabolites in future

238

studies would be useful in further understanding relevant biodegradation pathways, though such

239

analysis was beyond the scope of this work.

240 241

Analysis of SIP Gradient Fractions

242

The EUB gene-copy density profile for gradient fractions from the

12

C-BPA experiments

243

indicated a single peak with the center of mass in the range of 1.71 – 1.73 g/mL (Figure 2 (left

244

panel) and SI, Figure S-7 (Section S-9)). For the

245

density (~1.74 – 1.77 g/mL) was present in the EUB gene-copy density profiles from all four

246

13

13

C-SIP experiments, a second peak of higher

C-SIP experiments (SI, Figure S-7). The proportion of heavy-DNA (i.e., density >1.737 g/mL)


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13

247

gene copies present in the

248

high concentration BPA exposure) were 0.15 and 0.35, respectively (Table 1 and SI, Figure S-7,

249

calculated as the ratio of gene copies in the heavy fraction to the total gene copies in the sample).

250

In contrast, the proportion of heavy-DNA in the PET-LC and SCT-LC experiments were 0.51

251

and 0.63, respectively (Table 1). The higher proportion of heavy-DNA from the two SIP

252

incubations using the biomass previously exposed to a lower BPA concentration (i.e., PET-LC

253

and SCT-LC) reflected more effective assimilation of 13C-BPA or 13C-BPA metabolites over the

254

longer duration of BPA biodegradation observed for the ‘LC’ incubations (see Figure 1).

255

C-BPA samples from the PET-HC and SCT-HC experiments (i.e.,

Bacteria affiliated with the phylum Proteobacteria were preponderant in the heavy-DNA pool 13

256

from each of the four

257

reads, and α-Proteobacteria comprised between 65% and 98% of the bacterial reads from the

258

heavy-DNA pool (Figure 2, right panel and SI, Figure S-8 – S-11). The relative abundance of

259

α-Proteobacteria in the heavy-DNA pool samples from all four of the

260

significantly higher (z-test with α = 0.05) than that measured in the enriched samples collected

261

for SIP experiments (e.g., sample SCT-HC in Figure 2) or the initial samples collected from the

262

WWTP (e.g., sample SCT in Figure 2). This suggests that α-Proteobacteria may be preferentially

263

involved in the biodegradation and assimilation of BPA and its metabolites. Indeed, the

264

microbial communities derived from the heavy-pools of the 13C-BPA experiments are dominated

265

by relatively few bacterial genera as evidenced by their low Shannon-Wiener diversity indices

266

(SI, Table S-6: H’ = 0.09 – 1.87).

C-BPA SIP experiments accounting for greater than 95% of bacterial

13

C experiments was

267 268

Identification of Bisphenol-A Assimilating Bacteria



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Based on 16S rRNA gene-based phylogenetic analysis, the 13C-DNA pools from the PET-HC,

270

PET-LC, SCT-HC and SCT-LC samples were highly enriched (79%, 98%, 57% and 96%,

271

respectively) in bacteria related to Sphingobium spp. (genus-level analysis, Figure 3). Other

272

genera, notably Variovorax spp. and Novosphingobium spp., were also identified in the

273

heavy-DNA pool of the

274

(Figure 3). Based on a comparison of the inferred microbial community structure in the

275

heavy-DNA pool from each 13C-BPA SIP experiment with the heavy-DNA pool of the 12C-BPA

276

control, we identified genera which assimilated the

277

metabolites. It should be noted that the heavy-DNA pool associated with the

278

represents those microorganisms with DNA of inherently high buoyant density due to a high

279

G+C content and not due to assimilation of

280

assimilators were those present with a representation ratio greater than one. We define the

281

representation ratio to mathematically describe the ratio of the relative abundance of a genus in

282

the heavy-DNA pool from the

283

genus in the heavy-DNA pool from the 12C-BPA control. Our analysis revealed that the heavy-

284

DNA pool from the SCT-HC SIP experiment had the largest number of genera with a

285

representation ratio of greater than one (17) while the heavy-DNA pools from the PET-HC,

286

PET-LC and SCT-LC experiments each had three or fewer genera with a representation ratio

287

greater than one. The most prominent potential BPA assimilating genus was Sphingobium, which

288

had high relative abundance (>50% from all experiments) and an average representation ratio of

289

2.3 ± 0.8 across all experiments. The SCT-HC SIP experiment resulted in the most diverse set of

290

potential BPA assimilating organisms, which, in addition to Sphingobium spp., included

291

Sphingomonas spp., Pusillimonas spp., Novosphingobium spp., and GKS98 spp. among others

13

C-BPA SIP experiments using the PET-HC and SCT-HC biomass

13

13

13

C-label from

13

C-BPA or 12

13

C-labeled

C-BPA control

C-labeled BPA. Identified potential BPA

C-labeled experiment to the relative abundance of the same


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(Figure 4). Pusillimonas spp. were identified as potential BPA assimilators only through the

293

SCT-HC SIP experiment.

294

Bacteria belonging to the genera Sphingobium and Sphingomonas (see SI, Table S-7 for

295

details) have been previously identified as capable of using BPA or other endocrine disrupting

296

compounds as a sole carbon source for growth.30-35 Identification of Sphingomonas spp. as

297

potential BPA assimilators in this work thus provides direct evidence of their previously reported

298

growth on BPA. Here we also report the hitherto undocumented ability of Variovorax spp. or

299

Pusillimonas spp. to assimilate BPA or its biodegradation products. Relevant enzymes and

300

pathways for the degradation of aromatic compounds, including BPA, have been proposed for

301

several species of Sphingomonads.35, 36 Future research aimed at obtaining isolates of these BPA

302

assimilating species would help provide a deeper understanding of BPA biodegradation kinetics

303

and metabolic pathways. While isolates were not studied in this work, insights gained from

304

DNA-SIP can form the first line of evidence towards an exploration of degradation pathways in

305

mixed cultures by newly identified degraders or assimilators. Representation ratios above one for

306

Variovorax spp. and Pusillimonas spp. (Figure 4) were observed only in either of the two

307

experiments in which a high BPA (100 mg/L) concentration was utilized during the exposure

308

treatment (PET-HC or SCT-HC), suggesting a lower BPA affinity coefficient for these bacteria

309

relative to Sphingobium spp. Variovorax paradoxus are able to degrade complex organic

310

compounds and are often found in highly polluted engineered and natural systems.37-40

311

β-Proteobacteria GKS98 spp. have previously been implicated as typical freshwater bacteria.41

312

The potential complexity of BPA biodegradation by mixed culture systems warrants further

313

investigation into the individual roles and functions of specific genera. If, for example,

314

assimilating bacteria identified here are unable to act directly on BPA, they may instead be



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involved in assimilating metabolites of BPA, suggesting cooperation in the biodegradation and

316

assimilation of BPA by multiple bacteria within mixed microbial communities.

317 318

Exposure Conditions Influence Microbial Community Structure

319

Exposure to BPA in the high-concentration exposure treatment (100 mg/L) resulted in

320

differing enrichment of Sphingobacteria in the PET reactor and β-Proteobacteria in the SCT

321

reactor (SI, Figure S-12), which highlights the effect of the BPA exposure treatment on microbial

322

community structure. The relative abundance of the potential BPA assimilating Sphingobium

323

spp. increased in both PET-HC and SCT-HC exposure treatments (Figure 5). The low

324

concentration exposure treatment (500 µg/L BPA) also resulted in an increase in relative

325

abundance of Sphingobium spp. An increase in the relative abundance of other potential BPA

326

assimilating genera identified in this research (e.g., Pusillimonas spp. and Comamonas spp.) was

327

also observed in the SCT-HC exposure treatment. Interestingly however, the relative abundance

328

of the ‘BPA non-assimilators’ Methylobacillus spp. and Castellaniella spp. increased by more

329

than 5% in the SCT-HC but not the PET-HC exposure treatment (Figure 5). Note that the relative

330

abundance of these two BPA non-assimilating genera was less 0.02% in the samples collected

331

from both the PET and SCT reactors. The growth of Methylobacillus spp., which are reportedly

332

obligate methylotrophs,42 may be explained by the addition of methanol used to dissolve BPA, to

333

the fed-batch reactors. However, it is interesting that the same increase in relative abundance of

334

Methylobacillus spp. or other methylotrophs was not observed in the PET-HC reactor despite the

335

fact that the same total methanol COD was added to both high-concentration exposure

336

treatments. Rather, the microbial community dynamics in the PET-HC reactor favored BPA

337

assimilating organisms. Therefore, enrichments resulted in distinct communities (SI, Figure S-


Page 17 of 30


338

12), even across identical BPA exposure conditions, which suggests a persisting influence of the

339

starting community conditions.

340 341

Assessing the potential for BPA biodegradation and assimilation in WWTP bioreactors

342

Using the results from our SIP experiments, we classified the microbial communities into three

343

categories – potential BPA assimilators, potential BPA degraders and potential BPA non-

344

degraders. Potential BPA assimilators are those bacteria in the microbial community that took up

345

the 13C BPA-derived carbon, identified based on a representation ratio greater than one. Potential

346

BPA degraders are those bacteria that are able to degrade but not assimilate BPA and/or

347

biodegradation products and were identified based on (1) a representation ratio less than or equal

348

to one and (2) increased abundance and relative abundance from the start to end of each SIP

349

experiment. Potential BPA non-degraders are those bacteria that neither assimilate nor degrade

350

BPA or its biodegradation product(s). Genera with (1) representation ratios less than or equal to

351

one and (2) decreased or unchanged abundances from the start to the end of each SIP experiment

352

were classified as potential BPA non-degraders. This classification distinguishing assimilating

353

and non-assimilating bacteria underscores the utility of DNA-SIP. This distinction could not be

354

made by examining enrichment alone, since bacteria from both the assimilating and non-

355

assimilating groups increased in abundance over the course of BPA exposure.

356

Despite the significant differences in the treatment process conditions in the PET and SCT

357

reactors (see SI, Table S-1), the distributions of microbial genera enriched from these two

358

reactors were not statistically different (Mann Whitney test with α = 0.05). Both PET and SCT

359

initiated microbial enrichments were dominated by α, β, γ-Proteobacteria and Sphingobacteria,

360

with each being represented in approximately the same proportion (SI, Figure S-12). Analysis of



Page 18 of 30

361

the relative abundance of bacteria within the PET and SCT reactors suggests that while the

362

reactors were composed primarily of potential BPA non-degraders (79% and 78%, respectively),

363

both the PET and SCT reactors had the capacity for BPA degradation and assimilation, with

364

comparable combined fractions of potential BPA degraders (18% and 11%, respectively) and

365

assimilators (3% and 11%, respectively). However, the analysis does indicate that the SCT

366

bioreactor had the potential for a higher fraction of BPA assimilating microorganisms. Similar

367

analysis through molecular methods such as DNA sequencing or qPCR targeting relevant

368

organisms could be used as a preliminary assessment of the potential for TOrC degradation or

369

enrichment in both engineered and natural systems.

370

In conclusion, our application of DNA-SIP elucidated numerous Proteobacteria potentially

371

capable of assimilating BPA or biodegradation metabolites. We identified genera which have

372

been previously reported as capable of growth on BPA (e.g., Sphingobium spp. and

373

Sphingomonas spp.) and elucidated novel assimilators such as Pusillimonas spp. and Variovorax

374

spp. Results from our research can help to guide the design and development of specific

375

biomarkers to evaluate the potential for primary BPA assimilation and secondary assimilation of

376

its metabolites, in engineered and natural environmental systems. Our results confirm that

377

wastewater treatment process operating conditions and BPA exposure history play a role in

378

shaping microbial community structure and function. Additional research is warranted to explore

379

links between microbial community structure, function and the metabolic capability to degrade

380

and assimilate additional emerging contaminants. Such research will beneficially inform the

381

design and operation of engineered environmental systems and enable better characterization of

382

the ability of natural systems to degrade an ever-growing suite of complex anthropogenic

383

chemicals being detected in global water bodies.


Page 19 of 30


384 385

ACKNOWLEDGMENTS

386

Funding sources

387

This work was supported by the National Science Foundation, (CBET 1438578) and the Water

388

Environment Research Foundation.

389

Supporting Information. SI Sections S-1 through S-9, Tables S-1 through S-7 and Figures S-1

390

through S-12 can be found in the Supplementary Information file.



Page 20 of 30

TABLES

Table 1. Fraction of EUB gene copies in light (ρ ≤ 1.737 g/mL) and heavy (ρ > 1.737 g/mL) fractions from 12C-BPA controls and 13C-BPA SIP experiments. Enrichment exposure BPA Conc. (µg/L) Sample

100,000

500

PET-HC

SCT-HC

PET-LC

SCT-LC

light

99.3%

97.6%

96.6%

99.2%

heavy

0.7%

2.4%

3.4%

0.8%

light

85%

65%

49%

37%

heavy

15%

35%

51%

63%

Fractions from 12C-BPA control

Fractions from 13C-BPA experiment


Page 21 of 30


FIGURES

PET-HC SIP sample

PET-LC SIP sample SCT-HC SCT-LC SIP sample SIP sample

B P A C on c en tration (m g/L )

120 100 80 60 P E T -HC P E T -L C S C T -HC S C T -L C HC -no biomas s L C -no biomas s

40 20 0 0

10

20

30

40

50

60

Tim e (h ) Figure 1. BPA concentrations for time course evaluation (TC) reactors from SIP experiments. Results are shown for microbial communities originally derived from the PET and SCT bioreactors following high concentration exposure (PET-HC and SCT-HC) and low concentration exposure (PET-LC and SCT-LC). Results are also shown for the no biomass controls. Sampling time points are indicated.



1.70

1.0

SCTSCT-HC 1.0

1.0

SCTSCT-HC 12C

1.0

0.8

0.8

0.6

0.6

0.4

0.4

0.2

0.2

0.0 1.0

0.0 1.0

0.8

0.8

0.6

0.6

0.4

0.4

0.2

0.2

0.0

0.0

0.8

0.8

1.72

Relative Abundance

Density (g/mL)

1.71

1.73 1.74 1.75 1.76

0.6

0.4

0.2

0.6

0.4

0.2

1.77 0.0

0.2

0.4

0.6

0.8

SCTSCT-HC 13C

Light Fractions

SCT-HC 13C SCT-HC 12C

Heavy Fractions

SCT

1.69

Page 22 of 30

Other Flavobacteria Spartobacteria Verrucomicrobiae Acidobacteria Chlamydiae Chlorobia Sphingobacteria Cytophagia Anaerolineae Actinobacteria Phycisphaerae Planctomycetacia Deltaproteobacteria Betaproteobacteria Gammaproteobacteria Alphaproteobacteria

1.0 0.0

0.0

Fraction of EUB copies Figure 2. (Left Panel) Post ultracentrifugation EUB-gene copy-density profile from SCT-HC 12C control and 13C experiment. Dashed line indicates density threshold of 1.737 g/ml delineating heavy and light gradient fractions. (Right Panel) Relative abundances of different bacteria classes in the biomass collected from (i) the SCT reactor (SCT) and (ii) sample collected at the start of the SIP experiment following the high concentration exposure treatment (SCT-HC), shown alongside relative abundance of bacteria classes from the light and heavy fractions from the SCT-HC 12C and SCT-HC 13C.


Page 23 of 30


Is os phaera

Planctomycetacia

L uteimonas

γ -Proteobacteria

Xanthomonas

P E T-HC P E T-L C S C T-HC S C T-L C

R hodanobacter P s eudoxanthomonas P s eudomonas β -Proteobacteria

Methylobacillus Variovorax Hydrogenophaga C omamonas C as tellaniella P us illimonas GK S 98 S phingomonas

α -Proteobacteria

S phingobium Novos phingobium R hodobacter Actinobacteria

Acidimicrobineae 0.00

0.05

0.10 0.60 0.80 1.00

R elativ e A bu n dan c e Figure 3. Relative abundance of genera which are present in the heavy fraction following ultracentrifugation of DNA from the

13

C-BPA SIP experiments for the PET-HC, PET-LC,

SCT-HC and SCT-LC SIP experiments. Only those genera present at ≥0.05% in any of the heavy fractions are shown.



5

SIP expt.

Representation Ratio

Order

PET-HC PET-LC SCT-HC SCT-LC

GKS98

4

Acidobacteria Actinobacteria Alphaproteobacteria Betaproteobacteria Deltaproteobacteria Gammaproteobacteria Planctomycetacia Verrucomicrobiae Other

Pusillimonas

3

Page 24 of 30

Sphingobium

Variovorax Afipia

2

Novosphingobium Sphingomonas

1

0 0.00

0.02

0.04

0.06

0.08 0.50

1.00

Relative abundance in heavy fraction 13 from C-DNA SIP experiment Figure 4. Evaluation of relative enrichment of genera based on the Representation Ratio (see text for details) and relative abundance of genera in the heavy DNA fraction from SIP experiments.


Page 25 of 30


Assimilators

γ -Proteobacteria

F ulvimonas P s eudomonas Azomonas P eredibacter Alicycliphilus C urvibacter C omamonas B ordetella P us illimonas P igmentiphaga Achromobacter GKS 98 Variovorax S phingomonas Novos phingobium B os ea Afipia S phingobium

δ -Proteobacteria β -Proteobacteria

P E T -HC P E T -L C S C T -HC S C T -L C

α -Proteobacteria

0.00

0.02

0.04

0.06

0.08

0.10

Non-Assimilators Planctomycetacia

Is os phaera R hodanobacter L uteimonas F luvicola Hydrogenophaga Methylovers atilis Methylobacillus C as tellaniella P arvibaculum Acidimicrobineae

γ -Proteobacteria Flavobacteria

α -Proteobacteria

α -Proteobacteria Actinobacteria 0.00

1

0.02

0.04

0.06

0.08

0.10

C han ge in R elativ e A bun dan c e th rou gh E x pos u re

2

Figure 5. (top panel) Enrichment of BPA assimilators identified in this research (i.e., genera

3

with a representation ratio greater than 1) through the high and low concentration exposure 25 ACS Paragon Plus Environment


Page 26 of 30

4

treatments for those taxa with a relative abundance greater than 0.05%.

(bottom panel)

5

Enrichment of genera not able to assimilate BPA or metabolites but which exhibited an increase

6

in relative abundance of 1% or greater as a result of the exposure treatment.


Page 27 of 30

7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50


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ABSTRACT ART

ACS Paragon Plus Environment

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Identification of bisphenol-A assimilating microorganisms in mixed

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