Title: Silver nanoparticle inhibition of polycyclic aromatic hydrocarbons degradation by <i>Mycobacterium species </i> RJGII-135
Abstract: Letters in Applied MicrobiologyVolume 58, Issue 4 p. 330-337 Original ArticleFree Access Silver nanoparticle inhibition of polycyclic aromatic hydrocarbons degradation by Mycobacterium species RJGII-135 S.R. Mueller-Spitz, Corresponding Author S.R. Mueller-Spitz Department of Biology and Microbiology, University of Wisconsin Oshkosh, Oshkosh, WI, USA Correspondence Sabrina R. Mueller-Spitz, Department of Biology and Microbiology, University of Wisconsin Oshkosh, 800 Algoma Blvd, Oshkosh, WI 54901, USA. E-mail: [email protected] for more papers by this authorK.D. Crawford, K.D. Crawford Department of Chemistry, University of Wisconsin Oshkosh, Oshkosh, WI, USASearch for more papers by this author S.R. Mueller-Spitz, Corresponding Author S.R. Mueller-Spitz Department of Biology and Microbiology, University of Wisconsin Oshkosh, Oshkosh, WI, USA Correspondence Sabrina R. Mueller-Spitz, Department of Biology and Microbiology, University of Wisconsin Oshkosh, 800 Algoma Blvd, Oshkosh, WI 54901, USA. E-mail: [email protected] for more papers by this authorK.D. Crawford, K.D. Crawford Department of Chemistry, University of Wisconsin Oshkosh, Oshkosh, WI, USASearch for more papers by this author First published: 28 November 2013 https://doi.org/10.1111/lam.12205Citations: 16AboutSectionsPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat Abstract Polycyclic aromatic hydrocarbons (PAH) are a common environmental contaminant originating from both anthropogenic and natural sources. Mycobacterium species are highly adapted to utilizing a variety of PAH. Silver nanoparticles (AgNP) are an emerging contaminant that possess bactericidal properties, interferes with the bacterial membrane and alters function. Mycobacterium sp. strain RJGII-135 provided a model bacterium to assess changes in carbon metabolism by focusing on PAH degradation, which is dependent upon passive uptake of hydrophobic molecules into the cell membrane. A mixture of 18 PAH served as a complex mixture of carbon sources for assessing carbon metabolism. At environmentally relevant PAH concentrations, RJGII-135 degraded two-, three-, and four-ring PAH within 72 h, but preferentially attacked phenanthrene and fluorene. Total cell growth and PAH degradation were successively reduced when exposed to 0·05–0·5 mg 1−1 AgNP. However, 0·05 mg l−1 AgNP inhibited degradation of naphthalene, acenaphthylene and acenaphthalene. RJGII-135 retained the ability to degrade the methylated naphthalenes regardless of AgNP concentration suggesting that proteins involved in dihydrodiol formation were inhibited. The reduced PAH metabolism of RJGII-135 when exposed to sublethal concentrations of AgNP provides evidence that nanoparticle pollution could alter carbon cycling in soils, sediment and aquatic environments. Significance and Impact of the Study Silver nanoparticle (AgNP) pollution threatens bacterial-mediated processes due to their antibacterial properties. With the widespread commercial use of AgNP, continued environmental release is inevitable and we are just beginning to understand the potential environmental ramifications of nanoparticle pollution. This study examined AgNP inhibition of carbon metabolism through the polycyclic aromatic hydrocarbon degradation by Mycobacterium species RJGII-135. Sublethal doses altered PAH metabolism, which is dependent upon cell membrane properties and intracellular proteins. The changed carbon metabolism when exposed to sublethal doses of AgNP suggests broad impacts of this pollution on bacterial carbon cycling in diverse environments. Introduction Nanoparticle pollution is an emerging concern for aquatic and soil environments. Nanoparticles can enter the environment after release from a commercial product through disposal, weathering, application of sewage sludge or with sewage effluent waters (Kiser et al. 2009; Nowack et al. 2012). Both aquatic and soil environments are likely to receive increasing amounts of nanoparticle pollution. A concern is how nanoparticle pollution will affect bacterial-mediated processes because of their antibacterial activities. Silver nanoparticles (AgNP) are commonly used for these antibacterial properties. Key factors controlling nanoparticle toxicity to bacteria are size, shape, number of nanoparticles per cell, aggregation, elemental composition, capping agents and interaction with UV light (reviewed in Fabrega et al. 2011; Hajipour et al. 2012). Numerous modes of action have been proposed for the antibacterial activities including membrane destabilization, generation of damaging oxygen radical species, intracellular nanoparticle-DNA or protein interactions and release of metal ions (Hajipour et al. 2012; Schacht et al. 2013). Delayed growth has been commonly reported for various bacterial species on nanoparticle exposure (Sondi and Salopek-Sondi 2004; Li et al. 2010; Dimkpa et al. 2011), which may relate to decreased energy generation because of changes in electron transport (Anas et al. 2013). The ability to tolerate nanoparticles varies among different strains of bacteria based upon cell envelope composition or the production of extracellular polysaccharides (Dimkpa et al. 2011; Jin et al. 2013). Less is known about the physiological response stimulated by sublethal concentrations of nanoparticles. The potential for nanoparticle pollution to kill susceptible bacterial taxa and alter metabolism could have great impact upon bacterial processes such as carbon and nitrogen cycling. Decreased total bacterial counts and reduced diversity following nanoparticle exposure correspond to changes in bacterial community composition (Bradford et al. 2009; Doiron et al. 2012; Rodrigues et al. 2013). Subsequently, these composition changes altered bacterial community function following nanoparticle exposure (Das et al. 2012; Kumar et al. 2012). Choi and Hu (2009) illustrated that 1 mg l−1 AgNP inhibited bacterial nitrification. Similar effects have been seen in soils exposed to carbon nanotubes, where both soil biomass and extracellular enzymes essential for complex carbon utilization were reduced (Chung et al. 2011). These functional changes could reduce bacterial degradation of various recalcitrant organic pollutants. An important area of bacterial metabolism in certain polluted environments is the transformation of organic compounds to less toxic substances. Polycyclic aromatic hydrocarbons (PAH) are a major chemical class of concern because of their continual releases from a myriad of sources, detection in aquatic and terrestrial environments, and their negative impacts upon human health (Tobiszewski and Namiesnik 2012). These molecules are diverse in chemical structure and are classified as both low molecular weight (≤three-rings, LMW) and high molecular weight (≥four-rings, HMW). Regardless the source, the PAH are released as mixtures of LMW and HMW forms (Achten and Hofmann 2009; Tobiszewski and Namiesnik 2012). Environmental persistence is driven by low water solubility and hydrophobicity that increases with number of rings (Sikkema et al. 1995; Kanaly and Harayama 2010). Numerous soil and aquatic bacterial species can degrade PAH as a sole carbon and energy source, including Pseudomonas, Rhodococcus, Burkholderia, Arthrobacter, Polaromonas and Sphingomonas species (Kanaly and Harayama 2000, 2010). It has been suggested that the ability to degrade these recalcitrant molecules is a common metabolic trait of many bacteria taxa because the enzymes that carry out oxidation of the PAH to a dihydrodiol function against many natural aromatics (Kanaly and Harayama 2010). Yet, the ability to degrade both LWM-PAH and HMW-PAH commonly resides with Mycobacterium species (Cerniglia and Heitkamp 1990; Kanaly and Harayama 2010). Various Mycobacterium species isolated from sediments, and soils can breakdown pyrene, benzo(a)pyrene, fluoranthene, benzo(a)anthracene, naphthalene and anthracene, illustrating the wide metabolic activity of this genera (Cerniglia and Heitkamp 1990; Grosser et al. 1991; Rehmann et al. 1998, 2001; Kanaly and Harayama 2010). Mycobacterium species have mycolic acids that are very hydrophobic forming an impermeable cell wall (Niederweis et al. 2010), which aids in the uptake of hydrophobic PAH (Sikkema et al. 1995). Prior work has focused on characterizing breakdown pathway(s) for an individual PAH when provided as the sole carbon source allowing complex metabolic pathways to be generated. M. vanbaalenii PYR-1 degrades numerous aromatics as demonstrated by the presence of numerous ring-hydroxylating oxygenases, cytochrome P450 monooxygenases and other monooxygenases that are involved in the beginning step of aromatic degradation (Kweon et al. 2011). M. vanbaalenii PYR-1 activates complicated multiprotein degradation pathways to breakdown PAH with different metabolic networks for both LMW and HMW molecules (Kim et al. 2007, 2008, 2009; Kweon et al. 2007, 2011). These metabolic pathways have been detected in various aromatic degraders providing support that Mycobacterium species are highly evolved for PAH degradation (Kweon et al. 2011). The co-occurrence of organic and metal pollutants has long been a major concern for the feasibility of bacterial-mediated biodegradation as metal ions reduce bacterial diversity and function. In this study, we examined how sublethal AgNP concentrations would interfere with PAH degradation. Mycobacterium species strain RJGII-135 was used as a model PAH degrader (Grosser et al. 1991, 1995; Schneider et al. 1996; McLellan et al. 2002). RJGII-135 was grown in a mixture of PAH provided at low part per billion concentration in a low-carbon media to mimic natural systems in that carbon sources are somewhat limited but diverse (Arp et al. 2011). AgNP interactions with the cell could alter up-take or oxidation of the various PAH providing evidence for altered carbon utilization. Results and discussion In contaminated environments, PAH occur as complex mixtures, but little is known about RJGII-135 ability to utilize both LMW-PAH and HMW-PAH. The uninduced cells in the presence of alternative carbon sources degraded ten of the 18 PAH, which included all of the two- and three-ring molecules as well as the four-ring molecules pyrene and fluoranthene (Table 1). The HMW-PAH, specifically five- and six-ring molecules, were not degraded over the 72-h incubation period. RJGII-135 preferentially attacked 1-methylnaphthalene, 2-methylnaphthalene, fluorene and phenanthrene with less than 1% of the parent compounds remaining (Table 1). PAH degradation has been shown to be more effective in the presence of other carbon sources (Boldrin et al. 1993), which may account for the rapid loss of ten PAHs from an uninduced culture. However, if given a longer time, there may have been a decrease in the HMW-PAH. RJGII-135 is capable of degrading benzo[a]pyrene (BaP) and benz[a]anthracene (BaA; Schneider et al. 1996), although the bacterium more efficiently mineralizes pyrene and phenanthrene over BaP and BaA (McLellan et al. 2002). The related organism M. vanbaalenii PYR-1 also preferentially degrades phenanthrene over fluorene and pyrene (Kim et al. 2012). Pyrene can inhibit the degradation of benzo[a]pyrene (McLellan et al. 2002), which may account for why RJGII-135 was unable to degrade any of the HMW-PAH with LMW-PAH remaining. The degradation of HMW-PAH may only begin once the preferred PAH are removed due to activation of degradative pathways (Kweon et al. 2011), which relates to the substrate specificity of the rieske nonheme iron proteins (Kweon et al. 2010). Table 1. Concentration of parent polycyclic aromatic hydrocarbons (PAH) (average μg l−1 and standard deviation) in low strength media (0·1× R2B) with and without silver nanoparticles PAH Control 0·05 mg l−1 0·25 mg l−1 0·5 mg l−1 0 h 72 h 0 h 72 h 0 h 72 h 0 h 72 h NAPT 25 ± 2 5 ± 1aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 27 ± 3 11 ± 4aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 29 ± 4 45 ± 19bb P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. 25 ± 3 32 ± 15 1-MeNAPT 9 ± 3 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 11 ± 2 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 9 ± 3 1·7 ± 0·4aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 7 ± 3 4 ± 1aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 2-MeNAPT 17 ± 2 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 15 ± 2 2·3 ± 0·7aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 14 ± 3 11 ± 2bb P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. 12 ± 1 10 ± 1aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. ACNY 18·5 ± 0·9 3 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 17 ± 2 8 ± 3aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 18 ± 1 19 ± 3 17 ± 1 20 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. ACEN 16 ± 1 4 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 14 ± 1 9 ± 4aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 15 ± 2 13 ± 2bb P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. 14 ± 2 13 ± 2 FLUR 37 ± 4 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 36 ± 4 0·7 ± 0·7aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 37 ± 5 26 ± 6aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 35 ± 3 34 ± 4 PHEN 15 ± 2 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 13 ± 2 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 13 ± 3 NDaa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 12 ± 3 14 ± 2bb P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. ANTH 22 ± 3 7 ± 7aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 20 ± 3 6 ± 1aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 22 ± 10 16 ± 6 19 ± 5 19 ± 4 FLTH 10 ± 2 2 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 10 ± 2 1·4 ± 0·3aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 12 ± 3 15 ± 3bb P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. 15 ± 2 19 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. PYR 10 ± 2 2 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 10 ± 1 1·2 ± 0·3aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 11 ± 4 18 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. 13 ± 3 21 ± 2aa P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. NAPT: naphthalene, 1-MeNAPT: 1-methylnaphthalene, 2-MeNAPT: 2-methylnaphthalene, ACEN: acenaphthene, ACNY: acenaphthylene, FLUR: fluorene, PHEN: phenanthrene, ANTH: anthracene, FLTH: fluoranthene, PYR: pyrene. a P-values from paired t-test <0·001 comparing 0 and 72 h concentration of each PAH. b P-values from paired t-test <0·05 comparing 0 and 72 h concentration of each PAH. RJGII-135 tolerated double the concentrations of silver ion (Ag+; 1 mg l−1) as compared to AgNP (0·5 mg l−1) when grown in R2B media. It has been commonly reported that AgNP are more toxic than Ag+, although AgNP toxicity can be driven by the capping agent (Fabrega et al. 2011; Arnaout and Gunsch 2012). For AgNP capped with PVA, similar tolerance levels have been seen in other bacteria (Fabrega et al. 2011). The hydrophobic cell envelope of this fast-growing Mycobacterium species appeared not to increase its tolerance to AgNP. Tolerance to both forms of silver was reduced in 10% strength R2B, which may relate to the higher salt concentrations allowing the AgNP to aggregate reducing their toxicity in R2B. RJGII-135 tolerated up to 0·25 mg l−1 silver nanoparticles in 0·1 × R2B media, which reduced total viable cells from 6·99 × 109 colony-forming units ml−1 in the control to 3·19 × 107 colony-forming units ml−1 in the exposed samples after 72 h (data not shown). The 0·25 mg l−1 AgNP was therefore chosen as upper limit to maintain some viable cells with 0·5 mg l−1 as the concentration of AgNP that should be lethal to RJGII-135. The presence of AgNP in the PAH mixture reduced total cell growth and PAH degradation. There was a lag in growth for the 0·05 mg l−1 treatment at 24 h with lower final cell densities (Fig. 1). This indicates there was inhibition despite the presence of simple carbon sources such as pyruvate and glucose. Limited growth was detected at both of the higher AgNP concentrations indicating some cells remained viable. Reduced growth rates have been seen in Escherichia coli, Pseudomonas sp., Bacillus subtilis and Cupriadvius necator on exposure to various NP (Sondi and Salopek-Sondi 2004; Li et al. 2010; Dimkpa et al. 2011; Schacht et al. 2013). These reductions in growth suggest that AgNP interfered with various cellular processes in RJGII-135. Figure 1Open in figure viewerPowerPoint Mycobacterium species RJGII-135 growth in 0·1× R2B amended with the polycyclic aromatic hydrocarbons mixture and silver nanoparticles (AgNP). Circle (control), triangle (0·05 mg l−1 AgNP), diamond (0·25 mg l−1 AgNP) and square (0·5 mg l−1 AgNP). The loss of metabolic function was measured by reduced PAH degradation (Table 1). Changes in PAH degradation have been measured in the presence of other metals (Sokhn et al. 2001), so it makes sense that AgNP would possess similar properties. At all concentrations, RJGII-135 degraded 1-methylnaphthalene, 2-methylnaphthalene, acenaphthene and fluorene. Each of these PAH has a saturated carbon that would not be the target of various ring-hydroxylating oxygenases (Kweon et al. 2011), which may be why these molecules were continually degraded. At 0·05 mg l−1 AgNP, RJGII-135 degraded the same ten PAH as the control, but it less efficiently removed naphthalene, acenaphthene, 2-methylnaphthalene and acenaphthylene (Tables 1 and 2). The lag in growth did not correspond to decreased phenanthrene, fluorene and 1-methylnaphthylene, which remained unchanged (Table 2). In M. vanbaalenii PYR-1, both acenaphthene and acenaphthylene are part of different metabolic networks (Kweon et al. 2011), suggesting that various proteins involved in PAH degradation are inhibited by AgNP. When grown in the presence of 0·25 mg l−1 AgNP, RJGII-135 was unable to degrade either of the previously degraded four-ring molecules, naphthalene and acenaphthylene, but there was significant loss of five other PAH (Table 1). The consistent degradation of phenanthrene up to 0·25 mg l−1 AgNP provides support that this is the preferred PAH for RJGII-135. The 0·5 mg l−1 AgNP significantly reduced total PAH degradation (Fig. 2) with only the methyl-naphthalenes concentrations significantly reduced. The ability of the bacterium to remove the methyl naphthalenes indicates that transformation of these compounds likely occurs through a different metabolic pathway. The increased naphthalene concentration at the higher AgNP concentrations suggests that demethylation of the methyl naphthalenes may be occurring although this is not a metabolic conversion supported in the literature. Known metabolic pathways include the formation of methylsalicylic acid, methanolnaphthalene and carboxynaphthalenes (Selifonov et al. 1996; Dutta et al. 1998; LeBlond et al. 2000). Even though total cell density of the 0·25 and 0·50 mg l−1 treated culture was the same (Fig. 1), it is possible that the remaining viable cells had further reduced overall metabolic activity preventing further PAH degradation (Fig. 2). To represent the effect of AgNP on reduced metabolism, total PAH concentrations for each sample were compared (Fig. 2). The cumulative loss of 2-, 3-, and 4-ring PAH demonstrates that the RJGII-135 ability to metabolize PAH molecules was minimally impacted by low levels of AgNP (0·05 mg l−1), but this ability is severely restricted when exposed to higher levels of AgNP (Fig. 2). Previous work has shown that carbon nanotubes can change the bioavailability of organic pollutants (Zhou et al. 2013), whereas the continued loss of various PAH at the higher concentrations of AgNP suggests reduced bioavailability did not inhibit degradation. However, a change in the lipid bilayer could impact the up-take of these hydrophobic compounds. Nanoparticles have been shown to adsorb to cell membrane, but do not penetrate into the cell (Sondi and Salopek-Sondi 2004; Morones et al. 2005; Pelletier et al. 2010). These various NP-cell membrane interactions have been shown to change fatty acid composition (Anas et al. 2013), up-regulate generalized stress responses (Pelletier et al. 2010; Arnaout and Gunsch 2012) and even decrease dechlorination activity (Xiu et al. 2010). It is difficult to speculate on the specific mechanism controlling reduced PAH degradation, however, changes to membrane fluidity or proteins that carry out dihydrodiol formation may be induced by the AgNP. Table 2. Concentration of parent polycyclic aromatic hydrocarbons (PAH; average μg l−1 and standard deviation) and per cent degraded after 48-h incubation. Data from 0·25 and 0·5 mg l−1 not shown due to only 1-MeNAPT and 2-MeNAPT concentrations being reduced. Abbreviations for PAH are defined in Table 1 PAH Control 0·05 mg l−1 AgNP μg l−1 Per cent loss μg l−1 Per cent loss NAPT 9·5 ± 1·8 62 11 ± 4·6 60 1-MeNAPT 0 ± 0 100 0·07 ± 0·21 99 2-MeNAPT 1·9 ± 0·34 89 7·3 ± 3·6 52 ACNY 5·4 ± 1·2 70·9 10·6 ± 2·7 38 ACEN 6·8 ± 1·7 56·7 9·7 ± 3·5 32 FLUR 0·15 ± 0·55 99·6 0·02 ± 0·056 99·9 PHEN 0·69 ± 0·96 95·4 0·093 ± 0·26 99·3 ANTH 6·4 ± 4·7 70·7 6·5 ± 2·8 68·2 FLTH 2·8 ± 1·98 72·2 3·4 ± 2·8 67·3 PYR 2·4 ± 2·3 76·2 3·9 ± 3·4 62·4 Figure 2Open in figure viewerPowerPoint Total polycyclic aromatic hydrocarbons (PAH) loss for two-ring (NAPT, 1-MeNAPT, 2-MeNAPT), three-ring (ACNY, ACEN, FLUR, PHEN, ANTH) and four-ring (FLTH, PYR) molecules. Error bars represent the sum of standard deviations for each PAH. □2-ring; 3-ring; 4-ring. Increasing use of nanoparticles in consumer products may result in the presence of these unique materials in the natural environment. The interaction of bacteria with nanoparticles has the potential to alter the metabolic processes of bacteria. In the case of the PAH-degrading bacterium, RJGII-135, we have demonstrated that silver nanoparticles greatly impact cell growth in the presence of both simple and recalcitrant carbon compounds and changed PAH degradation at sublethal levels. These results indicated that AgNP exposure alters microbial driven processes with possible impacts on remediation efforts, wastewater treatment and carbon cycling in soil and aquatic environments signifying the threat of nanoparticle pollution to these environments. Materials and methods Silver Nanoparticle Synthesis and Characterization Silver nanoparticles were synthesized according to the procedure of Choi et al. (2008). A UV–Vis spectrum was used to confirm the formation of nanoparticles based upon the presence of an absorbance peak at 400 nm. Transmission electron microscopy (Zeiss EM10C) was used to determine the size of the AgNP by comparing to 90 nm polystyrene latex spheres (Ted Pella, Inc, Redding, CA). The spherical AgNP were c. 20 nm in diameter. Toxicity Studies Toxicity of AgNP and silver ion (AgNO3) were determined for RJGII-135 using the 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) cell viability assay (Malekinejad et al. 2011). Overnight cultures were grown at 28°C in R2B medium (EMD Millipore, Billerica, MA) for all experiments. Both forms of silver were examined between 0 and 1·5 mg l−1 (n = 6). To characterize toxicity at two different nutrient concentrations, the bacterium was exposed to both silver species in R2B and 0·1 × R2B medium. Cells (106) were exposed for 24, 48 and 72 h at 28°C under nonshaking conditions. After incubation, the optical density at OD 600 nm was measured using a plate reader (BioRad, Hercules, CA). MTT (5 mg l−1 in PBS) was added (15 μl) to each well and allowed to develop for 1 h. The converted dye was solubilized with 75 μl of 1% sodium dodecyl sulfate. Samples were read with a plate reader at 600 nm, and visual colour of the wells was recorded. Total viable cells counts were determined for 0, 0·05, 0·25 and 0·5 mg l−1 AgNP treatments to confirm toxicity measurements obtained with the MTT assay. Dilutions were plates in triplicate on R2A (BD, Franklin Lakes, NJ). Degradation of Polycyclic Aromatic Hydrocarbons To observe changes in Mycobacterium sp. strain RJGII-135 PAH metabolism, uninduced cultures were used to assess the ability of PAH to passively diffuse into the cell membrane. Three AgNP concentrations were chosen to represent a sublethal, moderately toxic and highly toxic level (0·05, 0·25 and 0·5 mg l−1). The AgNP and EPA Method 8310 PAH mixture (Restek, Bellefonte, PA) containing the 16 EPA priority PAH, and two different methylated naphthalenes were amended into 100 ml 0·1 × R2B media, so that the individual PAH molecules were present at the calculated final concentration of 25 μg l−1. An overnight RJGII-135 culture was added to each flask at a starting OD 600 nm of 0·039 (SD ± 0·0099). Each treatment was conducted in triplicate. Samples were removed from each treatment at 0, 24, 48 and 72 h to assess bacterial growth (OD 600 nm, n = 3) and loss of the parent PAH compounds (n = 3). A 0·5 ml sample of culture was combined with 0·5 ml of methylene chloride spiked with internal standards as follows: 50 μg l−1 of naphthalene-d8 (Fisher Scientific, Waltham, MA), 50 μg l−1 acenaphthene-d10 (Restek, Bellefonte, PA), 100 μg l−1 phenanthrene-d10 (Restek, Bellefonte, PA), 50 μg l−1 pyrene-d10 (Restek, Bellefonte, PA) and 50 μg l−1 perylene-d12 (Restek, Bellefonte, PA). Samples were mixed to allow all remaining PAHs to extract into the methylene chloride phase. After extraction, samples were stored at −80°C until analysis. GC-MS Analysis Samples were injected (1 μl) in splitless mode from the bottom of the vial (methylene chloride phase) into the inlet of a Thermo Focus GC and separated using a DB-5 MS column (Restek) equipped with a DSQ II mass spectrometer. The mass spectrometer was operated with EI ionization, and each PAH was detected in selected ion mode. Calibration curves for each PAH were prepared by analysis of standard mixtures with the internal standards and comparison of peak area ratios with the internal standards. The nearest (retention time) internal standard was used for quantitation of each PAH. Statistical Analysis The data were analysed using Minitab-16 (Minitab Inc, State College, PA). Analysis of variance was used to compare degradation of the individual PAH. Tukey's comparisons were determined to significant differences between the treatments. Paired t-tests were used to assess whether the PAH concentrations at 72 h were reduced as compared to the starting level. Acknowledgements This work was supported by University of Wisconsin Oshkosh Faculty development programme. We thank Dr Todd Kostman and Steve Rose for conducting the electron microscopy examination of the silver nanoparticles. We also thank Dr Brian K. Kinkle for supplying Mycobacterium sp. strain RJGII-135. Conflict of interest No conflict of interest declared. Supporting Information Filename Description lam12205-sup-0001-FigS1.tifimage/tif, 1.3 M