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Effects of TiO2 and Ag Nanoparticles on Polyhydroxybutyrate Biosynthesis By Activated Sludge Bacteria John Priester, Laurie C. Van De Werfhorst, Yuan Ge, Adeyemi Adeleye, Shivira Tomar, Lauren M Tom, Yvette M. Piceno, Gary Andersen, and Patricia Ann Holden Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/es504117x • Publication Date (Web): 19 Nov 2014 Downloaded from http://pubs.acs.org on November 24, 2014

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Environmental Science & Technology

Effects of TiO2 and Ag Nanoparticles on Polyhydroxybutyrate Biosynthesis By

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Activated Sludge Bacteria

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John H. Priester1, Laurie C. Van De Werfhorst1, Yuan Ge1, Adeyemi Adeleye1, Shivira

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Tomar1, Lauren M. Tom2, Yvette M. Piceno2, Gary L. Andersen2, Patricia A. Holden1*

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1Bren

School of Environmental Science & Management, Earth Research Institute, and

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UC Center for the Environmental Implications of Nanotechnology (UC CEIN), University of

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California, Santa Barbara, Santa Barbara, CA, 93106

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2Lawrence

Berkeley National Laboratory, Earth Sciences Division, Berkeley, CA 94720

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*Corresponding author: [email protected]; (805)- 893-3195; 3508 Bren Hall,

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University of California, Santa Barbara, CA, 93106-5131

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ACS Paragon Plus Environment


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TOC Art

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ABSTRACT

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Manufactured nanomaterials (MNMs) are increasingly incorporated into consumer products that

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are disposed into sewage. In wastewater treatment, MNMs adsorb to activated sludge biomass

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where they may impact biological wastewater treatment performance, including nutrient

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removal. Here, we studied MNM effects on bacterial polyhydroxyalkanoate (PHA), specifically

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polyhydroxybutyrate (PHB), biosynthesis because of its importance to enhanced biological

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phosphorus (P) removal (EBPR). Activated sludge was sampled from an anoxic selector of a

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municipal wastewater treatment plant (WWTP), and PHB-containing bacteria were concentrated

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by density gradient centrifugation. After starvation to decrease intracellular PHB stores, bacteria

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were nutritionally augmented to promote PHB biosynthesis while being exposed to either MNMs

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(TiO2 or Ag) or to Ag salts (each at a concentration of 5 mg L-1). Cellular PHB concentration

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and PhyloChip community composition were analyzed. The final bacterial community

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composition differed from activated sludge, demonstrating that laboratory enrichment was

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selective. Still, PHB was synthesized to near-activated sludge levels. Ag salts altered final

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bacterial communities, although MNMs did not. PHB biosynthesis was diminished with Ag (salt

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or MNMs), indicating the potential for Ag-MNMs to physiologically impact EBPR through the

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effects of dissolved Ag ions on PHB producers.

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INTRODUCTION Over 1600 consumer products1 reportedly contain manufactured nanomaterials (MNMs),

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yet associated risks to humans and environmental receptors are mostly unknown.2 MNMs in

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impermanent goods such as coatings, textiles, cosmetics, and personal care products enter

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sewage and therefore wastewater treatment plants (WWTPs). In WWTPs, MNMs adsorb to

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activated sludge biomass3-5 where MNMs may transform6, and accumulate in biosolids.7

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As previously reviewed, MNMs could interfere with biological wastewater treatment via

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impacts on associated bacteria.8 For example, Ag-MNMs inhibited bacterial respiration,9, 10 and

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damaged nitrifying bacterial populations11 including ammonia oxidizers.12, 13 Graphene oxide

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inhibited organic matter biodegradation, nitrification, and phosphorous uptake by activated

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sludge microbial communities14. Nanoparticulate SiO2 15 and Al2O3 5 inhibited enzymes involved

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in denitrification, resulting in elevated effluent nitrate concentrations. Similar effects of ZnO

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MNMs were observed, owing to nanoparticle dissolution and oxidative stress to denitrifying

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bacteria.16

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Coupled nitrification and denitrification underpin enhanced biological nutrient removal

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(EBNR) of nitrogen from wastewater, and thus MNM impacts on nitrifying and denitrifying

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bacteria, as above, are of special concern with regards to meeting N treatment objectives; similar

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concerns surround MNM impacts on P removal. In enhanced biological phosphorus removal

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(EBPR), polyphosphate accumulating organisms (PAOs) anaerobically synthesize and

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accumulate polyhydroxyalkanoates (PHAs, e.g. including polyhydroxybutyrate or PHB, and

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polyhydroxyvalerate or PHV), then aerobically metabolize intracellular PHAs during

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intracellular polyphosphate synthesis.17 In previous studies, neither Cu,18, SiO2,15 Al2O3,5 nor

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TiO2 19 MNMs impacted EBPR in activated sludge sequencing batch reactors (SBRs). Yet Zn 4


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ions released from ZnO MNMs,16 or Ag ions,20 interfered with either PHA synthesis or

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metabolism, respectively, resulting in relatively poor EBPR. The effects of Ag ions on PHA

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metabolism and concomitant polyphosphate accumulation were short term, owing to

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extracellular polymeric substances (EPSs) sequestering toxic Ag ions during a “recovery”

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period.20 However, Zn ions from ZnO MNMs favored PHV biosynthesis, which could have

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advantaged competing glycogen accumulating organisms (GAOs) that are not involved in

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EBPR.16

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Thus PHA biosynthesis, an important stage of EBPR, may be sensitive to MNMs. The

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consequences of MNMs deleteriously affecting microbes performing EBPR in WWTPs could

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include compromised treatment and thus discharge of poor quality, even eutrophying

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effluent, into receiving streams. In this study, we investigated the relative effects of either TiO2

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or Ag-MNMs, versus Ag ions, on PHB biosynthesis. We targeted PHB-biosynthesizing bacteria

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by first recovering PHB-rich bacteria from activated sludge using density gradient centrifugation.

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The recovered PHB-rich bacteria were starved to deplete PHB stores, and then fed nutrients to

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facilitate PHB biosynthesis while being exposed to MNMs. We also studied the bacterial

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community composition to understand if PHB biosynthesis changes were from bacterial

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community shifts, versus physiological impacts to PHB-biosynthesizing bacteria. The results of

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our research inform how MNMs could, through effects on PHB biosynthesis, impact EBPR in

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WWTPs.

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MATERIALS AND METHODS

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Nanomaterials, chemicals, positive control bacterial strain, and microbiological media

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TiO2 MNMs (P25; Evonik, Parsippany, NJ) were previously characterized as having

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mean diameters of 20 – 30 nm and a crystal structure of 81% anatase and 19% rutile.21, 22 The

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cysteine-capped Ag-MNMs (mean diameter 9 nm) were synthesized as before.23 AgNO3 was a

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positive control for Ag ions from dissolving Ag-MNMs. Unless otherwise noted, all other

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chemicals were reagent grade or better (Sigma Chemical, St. Louis, MO; and Fisher Scientific,

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Hampton, NH).

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Azotobacter vinelandii UWD (ATCC 53799) was a positive control PHB-biosynthesizing bacterial strain24-26 for validating the growth medium C:N molar ratio and the PHB analysis

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methods. To prepare A. vinelandii UWD, a sample from frozen stock (maintained at -80 oC in

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70% Luria Bertani (LB) broth plus 30% glycerol) was aseptically transferred onto solid growth

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medium (LB with 1.5% w/v agar) and incubated (30 oC, 18 h). A bacterial colony was dispersed

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into defined aqueous medium (4 mL) as the inoculum for culturing A. vinelandii UWD.

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The defined aqueous medium for bacterial culturing was based on Burk’s medium27 (pH

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7.1; per 1 L H2O: 0.2 g MgSO4.7H2O, 0.079 g CaSO4, 0.005 g FeSO4.7H2O, 0.00024 g

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NaMoO4.2H2O, 0.25 g KH2PO4, 0.85 g K2HPO4), modified as before28 (per 1 L H2O: 0.012 g

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ferric citrate, 1.16 g ammonium acetate) and using sodium acetate (49.13 g per 1 L H2O) as the C

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source to favor PHB biosynthesis.29 Several variations on Burk’s medium were tested to

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determine how the molar C:N ratio affected the proportion of cells that were PHB-positive. The

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final C:N molar ratio of 82:1 resulted in a relatively high proportion of A. vinelandii UWD cells

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that were PHB-positive (data not shown) and therefore was used. The N within AgNO3, where

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used, amounted to less than 0.5% (mass basis) of the Burk’s medium N.

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The modified Burk’s medium (30 mL, in 50 mL sterile culture tubes) was inoculated with

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suspended A. vinelandii UWD (50 µL), and incubated aerobically in the dark (30 oC, 200 rpm).

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After 24 h, intracellular PHB was visually confirmed by staining and microscopy.

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Activated sludge sampling and processing

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Activated sludge (AS, 2 L) was collected from an anoxic selector of one aeration basin at

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the El Estero WWTP in Santa Barbara, CA. The anoxic selector solids retention time (SRT) was

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1.6 d, and the dissolved oxygen (DO) and temperature were 0.3 mg L-1 and 23 oC, respectively.

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The AS was maintained on wet ice or refrigerated before sub-sampling (several, 20 µL each,

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within 2 hours) for staining and microscopically assessing PHB-positive bacteria. Triplicate AS

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sub-samples (30 mL each) were centrifuged (12,000 × g, 10 min., 4 oC); the supernatants were

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discarded, and the pellets were frozen (-80 oC) for later DNA extraction. Separate triplicate sub-

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samples (10 mL) were centrifuged (10,000 × g for 10 min. at 4 oC) and the supernatants

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discarded; the pellets were used immediately for PHB quantification. The remaining AS (approx.

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1 L) was capped and stored briefly (dark, 4 oC, 0.05) from pre-starvation

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cell counts (Table 1), indicating that starvation did not result in significant population loss.

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Thus, the AS contained PHB-rich bacteria that were concentrated by density gradient

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centrifugation, since cells in AS were fluorescent upon Nile Blue A staining, and their

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concentration was higher in the density gradient-centrifuged, pre-starvation biomass. Then, as

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expected, starvation decreased the proportion of Nile Blue A-positive cells.

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PHB Producing Bacterial Taxa were Identified by PhyloChip PhyloChip analysis provided further evidence for PHB-producing taxa in AS and in the

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post-starvation biomass. Identifying PHB-producing taxa was essential, as these may be

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susceptible to MNM exposure in WWTPs, causing deleterious changes to EBPR processes. The

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initial AS sample had an average relative richness of 2312 OTUs in 223 taxa, which was not

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significantly different from the post-starvation sample (Table S2), indicating that richness,

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similar to total cell counts, was unaffected by density gradient centrifugation and biomass

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starvation. Potential PHB producers were initially identified by examining the taxa that were in

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common between the AS and post-starvation samples, since those would comprise the subset

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recovered during density gradient centrifugation. By the qualitative analysis approach, 106 of the

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490 total taxa were present in all replicates of the AS and post-starvation samples (Table S3),

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meeting our criteria for identifying potential PHB producers. Of these, at least eight correspond

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to genera that were previously discussed in the PHA literature: Rhodobacter,52

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Novosphingobium,53 Thauera,54 Aeromonas,55, 56 Acinetobacter,54, 57, 58 Rhodocyclus,56

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Pseudomonas,54, 56 Sphingomonas.56

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From the quantitative statistical approach using the OTU level results, 203 of the 4615

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total OTUs were present in both the AS and post-starvation samples. These 203 OTUs were from

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seventy-one different taxa and originally had a significantly higher relative abundance in the

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post-starvation sample than in the AS, but after the P-value adjustment there was no significant

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difference (Table S4). However, despite the lack of significance after adjustment, it is still

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worthwhile to examine the taxa identified. Of these seventy-one taxa, forty overlap with

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previously-identified potential PHB producers using the taxa level results (Table S3). The OTU

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level results match most of the genera discussed with the taxa level results above, including

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Rhodobacter, Novosphingobium, Thauera, Aeromonas, Acinetobacter, and Sphingomonas. Two

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genera were identified in the taxa level results but not in the OTU level results: Rhodocyclus and

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Pseudomonas. One additional genus was identified at the OTU level but not in the taxa level

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results: Paracoccus, which is also discussed in the PHA literature.59

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Potential PHB producers were also identified by examining the taxa that were present in

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all replicates of the post-starvation samples, but absent or below the cutoff in all replicates of the

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initial AS sample. Using the qualitative approach, five additional taxa were identified using these

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criteria. These taxa are: order Chlorobiales, Isophaera spp., Burkholderia spp., order

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Myxococcales, and Aeromonas spp. The statistical approach using the OTU level results did not

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yield any additional taxa using these criteria, as it focused only on the taxa that were present in

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both the AS and post-starvation samples.

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PHB Was Biosynthesized under Laboratory Conditions The goal in augmenting (Burk’s media) post-starvation biomass and amending (with or without MNMs or silver salt) was to assess MNM impacts to PHB production. Assuming a 18


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bacterial dry mass of 2.8 × 10-13 g cell-1 60, and given that each PHB biosynthesis reaction (25

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mL) contained approximately 1.47 × 1012 cells L-1 (from 5.85 mL of post-starvation biomass at

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0.63 ± 0.04 ×1010 cells L -1, Table 1), the initial biomass concentration used in the PHB synthesis

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experiments was approximately 410 mg L-1. This estimated biomass concentration is well

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below the ca. 3000 mg L-1 mixed liquid suspended solids (MLSS) concentrations in other

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laboratory PHB biosynthesis studies,5, 61 and thus the amount of C relative to biomass was likely

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favorable (i.e. in excess) for PHB biosynthesis.

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By Nile Blue A staining and microscopy, PHB was produced in all final samples (Table

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1). Samples co-stained with SYBR Gold and Nile Blue A fluoresced green and red, respectively

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(Figure S3, c-f). Filamentous, bacillus, and coccus morphologies for PHB-rich cells were

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observed across the final samples; PHB-deficient cells were also observed (Fig. 2). The

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percentages of Nile Blue A-positive cells were much lower in the final versus the pre-starvation

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samples (Table 1). Still, there were differences in the final proportions of PHB-rich cells: while

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two treatments, the control and TiO2, resulted in measureable PHB biosynthesis, both treatments

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involving Ag (Ag salt or MNM) yielded final proportions of PHB-rich cells that were no

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different than the post-starvation biomass (Table 1). This indicated that little PHB biosynthesis

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occurred in the Ag salt or Ag-MNM treatments.

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Quantitatively, cellular PHB masses (as crotonic acid equivalents) for the control, TiO2

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and AgNO3 treatments did not significantly differ from that of the AS sample (t-test, p > 0.05).

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Cellular PHB mass was lower in the AgNO3 treatment as compared to the control and TiO2

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treatments, although the differences were not significant (t-test, p > 0.05, Table 1). Cellular PHB

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in the Ag-MNM treatment, however, was significantly lower (t-test, p < 0.05) than the other

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treatments and the AS sample. 19



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Cells exposed to TiO2 did not accumulate Ti at levels above background. Final Ag-

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MNM and AgNO3 biomass samples had mean Ag masses of 1.3 ± 0.18 × 10-4 pg cell-1 and 2.1 ±

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0.30 × 10-4 pg cell-1, respectively. There was no measureable Ti or Ag in the control biomass

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samples.

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Thus, similarly to a prior report,62 acetate amendment resulted in consistent but overall

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low PHB biosynthesis. However, there were treatment effects from amending with Ag-MNMs.

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Since the Ag-MNMs and the Ag salts similarly decreased the Nile Blue A-positive cell

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proportions (Table 1), the effects of Ag-MNMs on visualized and quantified PHB equivalents

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were likely due to Ag ions released from Ag-MNM dissolution. While we did not study Ag-

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MNM dissolution in Burk’s medium, these Ag-MNMs were previously shown to inhibit

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Escherichia coli and Pseudomonas aeruginosa population growth from reactive oxygen species

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(ROS)-mediated membrane damage due to even small percentages of MNM dissolution.23 Given

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that the prior study23 was at the same temperature (30 °C) and over similar exposure times (ca. 0

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to 24 hr) when compared to this study, the implication is that PHB biosynthesis by AS

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communities is sensitive to Ag ions dissolved from Ag-MNMs.

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PHB Biosynthesizing Enrichments Were Selective The sensitivity of PHB biosynthesis to Ag-MNMs, apparently from small amounts of Ag

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ions released during MNM dissolution, could have been from PHB-producing community

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compositional shifts, from cellular physiological effects, or both. To assess bacterial community

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compositional shifts, PhyloChip data were first analyzed to determine community shifts due to

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laboratory enrichment (i.e. between the control and post-starvation); subsequent analyses

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assessed treatment effects from MNMs or Ag salts, in comparison to the control. 20


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The average relative richness decreased significantly (P < 0.05, ANOVA, Tukey’s HSD)

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between the post-starvation and control samples, both at the taxa and OTU levels (Table S2),

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indicating biological selection during PHB biosynthesis. There was also a clear separation in the

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microbial community between the post-starvation and control samples, using either the taxa or

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OTU level results (Figures 3 & S4). This indicates that the control community was very different

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from the post-starvation community.

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In comparing the post-starvation and control communities, there were taxa that stayed in-

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common or increased in abundance. These taxa (or OTUs) were therefore viable and possible

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PHB producers under the experimental conditions. SIMPER analysis of the taxa level results

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revealed an average dissimilarity of 65% between the post-starvation and control treatments

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(Table S8). SIMPER also identified 84 taxa that were driving the separation on the NMDS plot

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(Figure S4A) between the post-starvation replicates and the control replicates (summed total of

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41% difference between groups; Table S5). These taxa were either present or absent in all

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replicates of the post-starvation sample and exactly the opposite in all replicates of the control

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sample. Of these eighty-four taxa, fifty-two were among the identified potential PHB producers

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(Table S3).

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There were twenty-eight taxa identified as potential PHB producers in the taxa level

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results that were present in all replicates of the post-starvation and control samples (Table S3).

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Of these twenty-eight taxa that were in common, five corresponded to potential PHB producer

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genera discussed above (Novosphingobium, Thauera (2), Aeromonas, Pseudomonas). From the

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initial subset of potential PHB producer OTUs identified (Table S4), four OTUs significantly

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increased between the post-starvation and control samples, even when evaluating using the

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adjusted P-values (two OTUs from Aeromonas, one each from the Enterobacteriales order and

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Vibrio) (Table S7).

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The taxa that disappeared and the OTUs that reduced in abundance, when comparing the

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post-starvation and control samples, are the taxa and OTUs that were obviously not enriched

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under these experimental conditions. There were eighty-three taxa identified as potential PHB

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producers in the taxa level results that decreased or disappeared when comparing the post-

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starvation sample to the control sample (Table S3). Of these eighty-three taxa, seven correspond

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to potential PHB producer genera discussed above (Rhodobacter, Sphingomonas, Rhodocyclus,

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Aeromonas, Acinetobacter (2), Pseudomonas). From the initial subset of potential PHB

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producer OTUs identified (Table S4), 125 OTUs significantly decreased between the post-

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starvation and control samples, even when they were evaluated using the adjusted P-values

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(Table S7). Of these 125 OTUs, thirty-eight correspond to potential PHB producer genera

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discussed above (five OTUs in Paracoccus, thirteen Rhodobacter, six Novosphingobium, six

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Thauera, six Acinetobacter). In summary, the control bacterial communities were less diverse

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subsets of the post-starvation microbial community (Fig. S1)—which was comprised of

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originally-dense (i.e. PHB-rich) bacteria from AS. Similarly to what others have observed using

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other phylogenetic analysis approaches (TRFLP and DGGE),54 laboratory enrichment for PHB

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biosynthesis resulted in taxonomic selection from more diverse AS communities.

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Silver Nitrate Altered PHB Producer Community Composition Final community differences across the treatments were analyzed to assess whether

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differences in PHB production—between the Ag (salt and MNM treatments) versus the other

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treatments including the control (Table 1)—were due to further community shifts or, by default, 22


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physiological effects. Neither the MNMs (TiO2 or Ag) nor the silver salt (AgNO3) treatments

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appeared to significantly alter community structure in the enrichment culture, as there was no

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separation between these treatments and the control on the PCoA or NMDS plots using the OTU

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level results (Figures 3b, S4b). Additionally, there was no significant difference for average

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relative richness between the Ag treatments and the control at the species or OTU level (Table

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S2). Using the adjusted P-values from the initial subset of potential PHB producer OTUs

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identified (Table S4), no significant difference was detected between the control and either of the

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Ag treatments (data not shown).

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However, the taxa level results revealed a slight separation of the AgNO3 treatment from

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the control and the MNM treatments using both PCoA and NMDS (Figures 3a, S4a). SIMPER

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analysis on the taxa level results revealed an average dissimilarity of 44% between the control

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and AgNO3 treatments (Table S8). In comparison, SIMPER analysis reported an average

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dissimilarity of 29% between the control and Ag-MNM treatments, and 33% between the control

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and the TiO2 treatments (Table S8). SIMPER also identified twenty-three taxa that were driving

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the separation on the NMDS plots (Figure S4A) between the control and the AgNO3 replicates

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(summed total of 21% difference between the groups) (Table S6). These taxa were either present

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or absent in all replicates of the control and exactly the opposite in all replicates of the AgNO3

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sample. Of these taxa, six are among the potential PHB producers already identified, with two of

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these absent in all replicates of the AgNO3 sample but present in all replicates of the control

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(Trichococcus spp.); four were absent in all replicates of the control sample but present in all

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replicates of the AgNO3 sample (order Sphingobacteriales, order Myxococcales, Pseudomonas

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spp., order Erysipelotrichales).

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Comparing the taxa level-identified potential PHB producers, the same Trichococcus

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taxa mentioned above were present in all replicates of the control and absent in all replicates of

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the AgNO3 treatment (Table S3). These taxa were also among those identified in SIMPER as

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driving the difference between the control and AgNO3 samples on the NMDS plot, as described

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above. Two additional taxa also showed a decrease, but not with all replicates (Table S3). When

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comparing the control and the Ag-MNM samples, fourteen taxa decreased in some, but not all,

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replicates (Table S3).

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Ag-MNMs Reduced PHB Biosynthesis Through Physiological Effects In summary, we observed that Ag-MNMs impacted PHB biosynthesis. We conclude

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that, because the impact to PHB biosynthesis by Ag salts was similar to that of Ag-MNMs, Ag-

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MNMs likely dissolved enough to inflict Ag ion-related toxicity. While bacterial community

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compositional shifts occurred with Ag ions, they did not occur from Ag-MNMs, thus further

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implying that the change in PHB production was from Ag ion effects on PHB producers’

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physiologies. We previously reported that these cysteine-capped Ag-MNMs, although slowly

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dissolving, released sufficient Ag ions to impair E. coli and P. aeruginosa growth.23 An Ag ionic

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effect thus also occurred here, resulting in decreased PHB biosynthesis. By comparison, Chen et

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al. reported, with sequencing batch reactors performing EBPR, that 1 mg L-1 Ag ions impaired

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short term (over 6 hours) PHA production and P removal, and the microbial community

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composition did not change.20 They also assessed effects at the Ag concentration studied here (5

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mg L-1), but within a longer operating reactor (up to 130 d), finding that continuous Ag exposure

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resulted in no measureable effects to either PHA production or P removal.20 Since our Ag-

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MNMs likely dissolved enough to cause Ag ion-related toxicity to PHB biosynthesis, our 24


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findings support that varying additions of Ag-MNMs—as could occur with fluctuating influent

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flows or loads—could have important process-level implications for PHB production and,

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therefore, EBPR in WWTPs.

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AKNOWLEDGEMENTS This research was primarily funded by the National Science Foundation (NSF) and the

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Environmental Protection Agency (EPA) under Cooperative Agreement DBI-0830117 (to

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P.A.H,). Additional funding was through a Lindberg Foundation Grant (gifted to P.A.H. at

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UCSB from awardee Dr. ChiHua Ho). Any opinions, findings, and conclusions expressed in this

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material are those of the author(s) and do not necessarily reflect those of either the NSF or EPA.

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This work has not been subjected to EPA review and no official endorsement should be inferred.

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We thank the staff of the El Estero WWTP for their assistance in AS sample collection.

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SUPPORTING INFORMATION AVAILABLE

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Supporting Information includes methods for evaluating the sucrose density centrifugation

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method for separating PHB-rich bacteria from activated sludge, an analysis of dissolved oxygen

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availability in the enrichment, a schematic of the experimental procedures, and other figures and

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data as referred to in the manuscript. This information is available free of charge via the internet

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at http://pubs.acs.org/.

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TABLES Table 1. Total (SYBR Gold-positive) and PHB–rich (Nile Blue A-positive) bacterial cell counts, the percentage of PHB–rich cells, and cellular PHB content (crotonic acid equivalents). The standard error of the mean was calculated for triplicate samples for each treatment. Like letters in each column indicate no significant difference (t-test, p > 0.05). Crotonic acid equivalents were not measured for the pre-starvation and post-starvation biomass. *NA; data not available Treatment

Activated Sludge Pre Starvation Post Starvation Control TiO2 AgNO3 Ag-MNM

Total counts per mL (/1010)

Nile Blue A counts per mL (/1010)

Percent Nile Blue A-positive cells

Cellular PHB (crotonic acid equivalents, pg cell-1)

1.14 ± 0.07a,d

0.13 ± 0.01a,c

11.1 ± 0.6a

10.5 ± 0.2a

0.55 ± 0.23b,c

0.52 ± 0.22a,b

91.9 ± 3.5b

NA*

0.63 ± 0.04b

0.06 ± 0.01b

10.1 ± 0.8a

NA

1.20 ± 0.09a,d 0.99 ± 0.05a,c 0.97 ± 0.09a,c 1.21 ± 0.07d

0.19 ± 0.03a 0.16 ± 0.02a,c 0.08 ± 0.03b,c 0.13 ± 0.03a,b

15.5 ± 1.5c 15.7 ± 1.4c 8.0 ± 2.8a 10.2 ± 1.8a

9.0 ± 1.1a 9.2 ± 1.4a 8.1 ± 1.4a 6.7 ± 0.3b

745 746 747 748 749 750 751 752 753 754 755

31



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FIGURES

Figure 1. Nile Blue A-stained activated sludge (AS), showing red-orange fluorescence in a fluorescent micrograph, overlapping a phase contrast image. PHB (granular, red-orange) is visible in some cells (triangles). Smaller, non–stained cells are also visible (arrows).

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Figure 2. Final samples, following incubation with modified Burk’s medium and staining with Nile Blue A. Red-orange fluorescence indicates the presence of PHB. The treatments are (a) control, (b) TiO2, (c) AgNO3 salts and (d) Ag-MNM.

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Figure 3. Principal coordinates analysis (PCoA) plots of the PhyloChip taxa level (a) and OTU level abundance (b) results. Analysis was performed on triplicate samples for each treatment. While the activated sludge and post-starvation samples appear similar, there is a clear separation between the sample replicates, and the sample replicates from the final treatments (control, TiO2 or Ag-MNMs, or AgNO3). At the taxa level, there is also a separation of the AgNO3 treatment replicates from the control treatment replicates.

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