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Article Cite This: ACS Omega 2019, 4, 2196−2205
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Eco-friendly Organic Nanotubes Encapsulating Alkaline Phosphatase and Ecotoxicology of Nanotubes to Natural Bacterial Assemblages in Coastal Estuarine Waters Michael T. Montgomery,*,† Greg E. Collins,‡ Thomas J. Boyd,† Christopher L. Osburn,§ Diana Oviedo Vargas,§,∥ and Qin Lu‡ Marine Biogeochemistry Section, Code 6114 and ‡Bio/Analytical Chemistry Section, Code 6112, Naval Research Laboratory, 4555 Overlook Avenue, Washington, District of Columbia 20375, United States § Department of Marine, Earth and Atmospheric Sciences, North Carolina State University, Jordan Hall, Box 8208, Raleigh, North Carolina 27695, United States ACS Omega 2019.4:2196-2205. Downloaded from pubs.acs.org by 185.89.100.155 on 01/30/19. For personal use only.
†
S Supporting Information *
ABSTRACT: Phosphatase-encapsulated nanotubes have potential in environmental remediation of organophosphate contaminants (e.g., pesticides, nerve agents). We investigated alkaline phosphatase (AP) activity when encapsulated in self-assembled lithocholic acid nanotubes (LCA-AP) in water samples along a transect from Cypress bog headwaters through estuarine waters and to Atlantic Ocean seawater. Apparent Vmax (appVmax) for both LCA-AP and unencapsulated AP (Free-AP) was most rapid at mid-estuary and most inhibited at the humic-rich bog. LCA-AP retained a higher-activity percentage, suggesting that encapsulation may afford some protection from denaturing effects of humics. Apparent Km (appKm) of Free-AP (1−2.3 μM) was largely unaffected by preincubation with transect water, whereas appKm of LCA-AP was higher with bog water (5.3 μM) relative to other stations. When comparing Free-AP and LCA-AP, increasing salinity generally decreased the catalytic efficiency of the LCA-AP, but had little effect on that of the Free-AP. In addition, both showed the same pattern of lowest efficiency in bog water, which increased with salinity to 21 practical salinity units before decreasing at full-strength salinity. With the exception of the similarly low values in the bog water (1.04 for LCA-AP, 1.34 for Free-AP), absolute values of catalytic efficiency for LCA-AP were about 17% (range: 14.5−19.3%) of that for Free-AP. Nanotube addition had little ecotoxicological effect on heterotrophic bacterial production in waters sampled along the transect. Microbially associated, intrinsic AP activity showed a similar pattern along the transect to LCA-AP, suggesting that AP environmental control and regulation in nature may inform study of nanomaterials.
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INTRODUCTION Enzyme-encapsulated carbon nanotubes have been proposed for use in surface decontamination,1 removal of endocrine disruptors,2 and as an environmental remediation strategy (e.g., wastewater treatment, contaminant degradation, sorbants3). Carbon nanotubes are also a vector for antibiotics4 and insulin5 although questions have been raised about the ecotoxicological effects and life cycle issues of nanomaterial release into natural ecosystems (e.g., refs 6, 7). One candidate for inclusion in nanotube remediation systems is phosphatase, which has broad substrate specificity8 and can degrade organophosphate contaminants, such as pesticides9−12 and nerve agents.13 These enzymes have been well studied in coastal aquatic ecosystems because of their potential importance in controlling phosphorus cycles.14 One benefit of nanotube encapsulation is to protect free enzymes from degradation and denaturation associated with release into coastal environments.15−17 Cypress bog water is a rich source of humics, which inhibit alkaline phosphatase (AP) © 2019 American Chemical Society
activity via noncompetitive inhibition, although this effect may only be temporary.18 Over short durations, bog water exposure may provide an extreme scenario for effects on enzymatic activity associated with environmental release. Depending on dissolved organic matter (DOM) characteristics in natural waters, there can be confounding effects on enzyme−substrate interactions. DOM can reduce enzyme activity by denaturing proteins or by altering local microenvironmental conditions (e.g., pH, metal chelation19) or, conversely, protect phosphatases from heat denaturation or protease cleavage.20 Physical trapping in humics can also preserve enzymatic activity in longterm incubations20 and enhance nanotube suspension in aqueous media, which would affect their transport in coastal waters.21,22 These model enzyme−nanotube systems are often tested using laboratory buffers so their activities are rarely Received: October 4, 2018 Accepted: December 11, 2018 Published: January 29, 2019 2196
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Figure 1. Alkaline phosphatase (AP; % of Tris buffer control) activity using 2 mM diphosphate fluorescein (DPF) substrate after preincubation in water from a salinity (PSU) transect from the Newport River Estuary, North Carolina (circles) or salinity control (squares) created from mixing rainwater with full-strength seawater relative to preincubation with Tris buffer with either encapsulated (LCA-AP; A) or dissolved enzyme (FreeAP; B). The error bars represent the standard deviation of triplicate samples.
Table 1. Dissolved Organic Carbon (DOC) Concentration (mg L−1), Salinity (PSU), and Stable Carbon Isotope Values (δ13DOC, ‰) for Stations in the Newport River Estuarine System and Bogue Sound, North Carolina (September 2016)a station analyses
NC-0A
NC-5A
NC-11A
NC-21A
NC-36A
NC-M1A
DOC (mg L−1) salinity (PSU) δ13-DOC (‰)
36.8 0 −28.1
17.5 5 −27.6
13.9 11 −27.2
6.82 21 −26.1
1.53 36 −23.1
2.68 36 −23.4
DOC concentration values have a coefficient of variation of 100%) in the salinity 2197
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Figure 2. Alkaline phosphatase appKm (μM) varied when preincubated with water from a salinity (PSU) transect across the Newport River Estuary, North Carolina (circles) or salinity control of rainwater mixed with seawater (squares) when the added enzyme was either encapsulated into LCA nanotubules (LCA-AP; A) or dissolved (Free-AP; B).
Figure 3. appVmax of alkaline phosphatase (μM min−1) varied when preincubated with water from a salinity (PSU) transect across the Newport River Estuary, North Carolina (circles) or salinity control of rainwater mixed with seawater (squares) when the added enzyme was either encapsulated into LCA nanotubules (LCA-AP; A) or dissolved (Free-AP; B).
LCA-AP and Free-AP, the difference was more dramatic with LCA-AP (2.39 vs 1.26 μM, respectively). The appKm values measured here are generally lower than 4.3−21.6 μM for most intrinsic phosphatase measurements in aqueous systems summarized by Sinsabaugh et al.32 but similar to 0.18−4.5 μM reported by Nedoma et al.33 for oligotrophic seawater and that of Free-AP in Tris buffer alone (Km = 1.77 (±0.06) μM; LCA-AP in Tris, Km = 0.92 (±0.06) μM). Here, the direct effect of decreasing salinity appears to be an increase in enzyme−substrate affinity, whereas high DOC concentration or other features that differentiate bog water from rainwater appear to preferentially counter this increase in affinity for LCA-AP relative to that seen with Free-AP. It is possible that substrate diffusion limitations resulting from humic-nanotube aggregation lead to higher appKm values34 at lower salinities, closer to the pocosin (palustrine wetland) source of humic-rich DOC. Humic sorption to nanotubes can render nanotubes less sensitive to pH, which, in this experiment, increased from freshwater to seawater.35 In addition, humic sorption increases with sodium cation concentration, which occurs when migrating from freshwater to seawater,35 although humic concentration decreases
control mixtures of seawater and rainwater (5−21 PSU; Figure 1). AP Apparent Km. LCA-AP appKm increased from 0.70 (±0.11) to 3.09 (±0.25) μM with salinity in the salinity control, suggesting some interaction between LCA and seawater ions (Figure 2A). Across the estuarine transect, however, LCA-AP appKm decreased dramatically from 5.31 (±0.96) μM at NC-0 to a low value of 2.16 (±0.46) μM at NC-11 (11 PSU) and then increased somewhat to 3.09 (±0.25) μM with full-strength seawater. This pattern of appKm for LCA-AP may suggest binding strength between enzyme and DPF substrate is highest at mid-salinity (11−21 PSU) and lowest in humic-rich bog water, whereas Free-AP showed relatively little change in appKm across the transect. Free-AP app Km ranged from 2.18 (±0.24) to 3.44 (±0.67) μM, but varied much less across the NRE transect with the highest value at 0 PSU (Figure 2B). For the salinity control with FreeAP, appKm gradually increased from 1.12 (±0.68) μM in the freshwater end member to 2.49 (±0.089) μM at full-strength seawater, possibly as increasing ionic strength decreased enzyme−substrate affinity (Figure 2B). Although appK m increased with increasing salinity in the control for both 2198
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Table 2. Kcat (s−1; AVG (SD)) and Catalytic Efficiency of Free-AP and LCA-AP When Preincubated with Water from a Salinity (PSU) Transect across the Newport River Estuary, North Carolina (NC), a Salinity Control of Rainwater Mixed with Seawater (Control), or Tris Buffer (TB) When the Added Enzyme Was either Encapsulated into LCA Nanotubules (LCA-AP) or Dissolved (Free-AP) Kcat (s−1; AVG(SD)) treatment
salinity (PSU)
Free-AP
0 5 11 21 36 TB (5) 0 5 11 21 36 TB (5)
LCA-AP
NC 4.61 117 209 284 266
(0.57) (4.18) (11.1) (3.31) (13.5)
5.53 26.7 37.1 55.2 47.S
(0.40) (1.38) (3.00) (2.13) (1.20)
control 139 244 262 267
(2.84) (9.10) (9.83) (4.40)
214 32.7 47.8 48.6 48.5
(7.53) (3.80) (2.46) (2.51) (4.92)
45.8 (3.38)
catalytic NC
efficiency control
1.34 51.3 95.8 120 107
124 122 135 122
1.04 8.38 17.2 23.1 15.5
121 46.9 49.3 32.3 19.1 49.6
contact between humic acids (and other higher-molecularweight inhibitors) and the enzyme itself. Reducing such interaction could reduce competition for the enzyme active site39 and also decrease tertiary and quaternary structural changes to the enzyme that may result from contact with natural organic matter. Humic acids and proteins can form bonds40 and encapsulation in nanotubes may protect the enzyme in situ from naturally inhibitory humic acids. Scully et al.41 reported that acidic iron-catalyzed reactions deactivated commercial phosphatase in humic lake water. Steen and Arnosti42 reported up to 42% loss of activity using a commercial AP product (Free-AP) within 5 h of exposure to full-strength seawater (although incubations were at 7 °C rather than 25 °C). Differential effects on activity along this transect could be due to competing changes in association with humics, pH, and/or ionic strength.43 Kcat and Catalytic Efficiency. Two related measures for comparing the function of a single enzyme under different conditions are Kcat, which measures the turnover of substrate molecules per enzyme molecule per second, and catalytic efficiency (Kcat/Km). In general, Kcat and catalytic efficiency of LCA-AP were about 20% of that for Free-AP for most of the treatments and controls (Table 2). Kcat of LCA-AP across the salinity control was lowest in rainwater (0 PSU) at 32.7 ± 3.80 s−1 and ranged narrowly from 47.8 (±2.46) to 48.6 (±2.51) s−1 across the rest of the control salinities. However, Kcat of LCA-AP ranged much more broadly in the estuarine waters from 5.53 (±0.40) s−1 in the 0 salinity bog water to 55.2 (±2.13) s−1 at 21 PSU. The rate in the NC-21 PSU station water was even higher than that measured for the Tris buffer control (55.2 (±2.13) vs 45.8 (±3.38) s−1). Catalytic efficiency of Free-AP in the salinity control was highest at mid-salinity (135; 11 PSU) and lowest in fullstrength seawater (107; 36 PSU) but varied within a relatively narrow range of 107−135. For the estuarine transect water, catalytic efficiency ranged from 1.34 in the bog water end member (NC-0) up to 120 at 21 PSU, which is nearly within the range of the salinity control. The highest catalytic efficiency at 21 PSU was also very similar to that of the Tris buffer control (121, TB). That for LCA-AP was similar for the freshwater and brackish water samples in the control (46.9 at 0 PSU, 49.3 at 5 PSU) but then decreased with increased salinity to 15.5 in full-strength seawater. Catalytic efficiency was lowest
dramatically with migration to seawater confounding this potential effect. It is also possible that changes in appKm with salinity could be due in part to changes in dissolved organic and inorganic phosphate pools along the transect. Changes in phosphate levels would compete with DPF substrate, although values of 0−1.5 μg L−1 reported for the Newport River Estuary36 are over 2 orders of magnitude lower than the substrate we add, and this is before we dilute the sample water (1:4) with Tris buffer for the assay. Competitive inhibition at these levels would not be detectible with the assay as performed. AP Apparent Vmax. For NRE transect preincubations, enzyme activities of both LCA-AP and Free-AP were inhibited by over 90% with humic-rich water (NC-0) relative to their highest value in NC-21 mid-estuarine water (Figure 3), with LCA-AP ranging from 0.044 (±0.003) to 0.45 (±0.017) μM min−1 and Free-AP ranging from 0.037 (±0.005) to 2.27 (±0.026) μM min−1. Across the NRE transect, appVmax values were lower for LCA-AP, although changes with increasing salinity (Figure 3) were very similar. Higher Vmax values for Free-AP have been reported comparing other dissolved verses immobilized enzymes (see ref 37 and references therein) and more specifically to encapsulation by LCA.27 In the salinity control, appVmax increased to 11 PSU and then leveled off to full-strength seawater for both LCA-AP (0.26 ± 0.03−0.38 ± 0.009 μM min−1) and Free-AP (1.11 ± 0.023−2.13 ± 0.11 μM min−1), with both having their lowest rates in full-strength rainwater (0 PSU; Figure 3). For the Tris buffer control, app Vmax values for Free-AP and LCA-AP were 1.71 (±0.06) and 0.37 (±0.03) μM min−1, respectively, which were similar to the 5 PSU salinity control (note the salinity of the Tris buffer was about 5 PSU). In general, DOC concentration (including humics) was very high at the freshwater end member, 36.8 mg L−1 at NC-0A, but then decreased throughout the estuary to 1.53 mg L−1 at NC36A due to dilution with seawater and bacterial metabolism38 (Table 1). This general trend of less enzymatic inhibition with decreasing DOC was not unexpected; however, the nanotubeencapsulated enzymes recovered their appKm more rapidly in incubations transiting from freshwater toward the marine end member (down estuary). One possible mechanism for protection of enzymes from the denaturing effect of humic acids is that the nanotubular structure sterically reduces direct 2199
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Table 3. LCA Nanotubules (μM C Final Concn.) Were Added to Unfiltered Water Samples from Newport River Estuary, North Carolina (September 2016) To Determine the Effect of This Carbon Source on Heterotrophic Production (AVG (SD) μg C L−1 day−1) of the Natural Bacterial Assemblagea bacterial production (AVG (SD) μg C L−1 day−1) LCA addition (μM C) 0 0.32 3.2 32 320
NC-0A 24 22 23 23 24
(0.49) (0.61) (0.55) (1.8) (2.9)
NC-5A 95 102 105 100 96
(6.1) (11) (5.7) (3.0) (13)
NC-11A
NC-21A
NC-36A
NC-M1A
99 100 97 98 98
52 54 58 56 57
12 11 11 11 12
27 30 28 26 29
(14) (3.3) (3.3) (5.8) (2.8)
(7.3) (5.4) (0.87) (3.0) (2.9)
(1.0) (2.5) (0.76) (0.91) (1.0)
(3.3) (2.0) (1.7) (3.8) (2.1)
a
All experiments with the transect were performed using LCA. An addition experiment using LCA-AP was used with Bogue Sound seawater (NCM1A) and also showed no effect of the nanomaterial.
LCA-AP) by LCA or LCA-AP addition during the 1 h exposure (Table 3). Intrinsic Phosphatase Activity by Natural Microbial Assemblages. Intrinsic phosphatase activity of unfiltered (whole) water was measured along the NRE transect (September 2016) to determine if there was commonality between the salinity gradient activity change and changes measured for LCA-AP and Free-AP additions to filtered transect water samples. Such intrinsic activity is typically cellassociated from both bacteria and phytoplankton among the natural microbial assemblage and considered important to understanding phosphorus cycling in coastal waters.51 Although it is expected that changes in phosphatase activity would be most closely related to in situ (autochthonous) enzyme production by the assemblage, rates can also be a function of environmental conditions52 that could reflect on the potential usefulness of added LCA-AP. Similar to that seen with laboratory and commercial AP preparations, intrinsic phosphatase activity was most rapid at mid-salinity (11, 21 PSU) and least rapid in the humic-rich bog water (0 PSU; Figure 4A). Intrinsic phosphatase rates measured in the estuarine and marine portion of the NRE of 0.50−1.0 μmol L−1 day−1 were bounded by the appVmax for phosphatase summarized for ecosystems by Sinsabaugh et al.32 of 0.22−160 μmole L−1 day−1. For comparison, appVmax for commercial and laboratory preparations of Free-AP and LCA-AP presented here (at presumably much higher AP concentrations) was about 1−2 orders of magnitude more rapid than most of the reported natural microbial assemblage rates.32 Intrinsic phosphatase activity change with salinity appeared to be similar to that of bacterial production, although phosphatase activity was least rapid at 0 PSU (as was seen with LCA-AP and Free-AP incubated in cell-free sample) and bacterial production was least rapid at 36 PSU (Figure 4). When phosphatase activity is normalized to bacterial production, the percentage increased with salinity (Figure 5). If phosphatase activity was a simple function of enzyme production by the natural assemblage, this would suggest that a larger component of bacterial metabolism was devoted to scavenging phosphorus with migration down the estuary. This may be counterintuitive given that pocosin bog water is typically much more phosphorus-limiting to bacterial metabolism than estuarine or marine waters.53 However, based on the findings that phosphatase activity of both Free-AP and LCA-AP are inhibited by an order of magnitude from 21 to 0 PSU (Figure 3), this relationship may be more of a function of inhibition of microbially expressed enzyme rather than devotion of a greater amount of cellular resources to phosphate cycling down estuary. It is also possible that a shift in higher
in bog water (1.04, NC-0) and highest at 21 PSU (23.1). When comparing Free-AP and LCA-AP, increasing salinity generally decreased the catalytic efficiency of the LCA-AP, but had little effect on that of the Free-AP. In addition, both showed the same pattern of lowest efficiency in bog water, which increased with salinity to 21 PSU before decreasing at full-strength salinity. With the exception of the similarly low values in the bog water (1.04 for LCA-AP, 1.34 for Free-AP), absolute values of catalytic efficiency for LCA-AP were about 17% (range: 14.5−19.3%) of that for Free-AP. Effect of LCA on Bacterial Production. LCA may be both cytotoxic and metabolized as a carbon source by different components of the natural bacterial assemblage (see ref 44 and references therein), so release of such nanomaterial into natural waters may have unpredictable biological consequences. Also, direct binding of bacteria to nanotubes can reduce nanotube binding to inorganic surfaces and enhance their physical transport through natural systems.45 The effect of LCA addition on heterotrophic production (i.e., Leucine method) of natural bacterial assemblages was used as metric for assessing the short-term environmental impact of LCA release.46 LCA (final concentrations: 0.319, 3.19, 31.9, and 319 μM LCA-C) was added to unfiltered NRE transect water from the five salinities used in the previous experiment, although from a separate sampling (September 2016) designated as NC0A, -5A, -11A, 21A, and -36A. The bioassay was performed using LCA-AP at a single Bogue Sound station (NC-M1A; 36 PSU). Although it is not known what levels of LCA would be added to estuarine waters as part of a future application, this range was chosen to bracket that used in the enzymatic activity assay. Maximum LCA concentration used here (in terms of carbon) would increase the full-strength seawater DOC concentration (NC-36A) by about 20% and Cypress bog water concentration by about 1% (NC-0A; Table 3) and may likely exceed the practical use concentration of nanomaterials in aquatic systems.47 To date, there have only been limited studies of such bile acids in aquatic systems and those are mostly as an ecological indicator for agricultural input to aquatic ecosystems rather than as an anthropogenic contaminant.25,48 14C-radiolabeled nanotubes have been used to measure mineralization of nanotubes, although this work involved metabolism by cultured bacterial isolates rather than natural assemblages.49 Bile salts as a class of organic matter are reported to be widely metabolized by environmental bacteria.50 In this short-term bioassay, bacterial production ranged from 12 (±1.0) to 99 (±14) μg C L−1 day−1 over the unamended transect waters and there was no measurable (200% of bacterial production) and decreased with salinity, although this system did not have humic-rich bog water in the freshwater end member. Although there may be structural differences between commercially and estuarine assemblage-produced enzymes,39,65 coupling experiments using laboratory preparations of enzymes with intrinsic activity measurements may inform both ecological and bioremediation studies.
Figure 4. Intrinsic phosphatase activity (A, μmol L−1 day−1) of the natural bacterial assemblage from the Newport River Estuary, North Carolina transect (September 2016) showing highest activity at midsalinity (PSU) similar to the results seen with added alkaline phosphatase laboratory preparations. It also showed a similar pattern to heterotrophic bacterial production (B, μg C L−1 day−1).
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CONCLUSIONS We investigated the utility of organic nanotubes for enzyme encapsulation and delivery of potentially bioremediating agents to coastal waters. The use of natural waters in such investigations is relatively rare in this discipline but may be an important strategy for understanding the potential of these vectors in more chemically diverse and relevant media than laboratory buffers. Although catalytic efficiency of LCA-AP was about 17% of that for Free-AP (as one might expect given the barriers to diffusion with LCA), the encapsulated enzyme maintained much of the same pattern of functionality when preincubated with complex estuarine transect waters. Highest activities of both forms of AP, as well as naturally occurring AP, were often highest at mid-salinity, which was unexpected. In addition, preliminarily, the LCA did not appear to either negatively or positively impact the growth of natural bacterial assemblages, which has been an ecological concern with use of other nanomaterials. Although these are just the first steps in this investigation, LCA may hold some potential as one of the few delivery agents for in situ environmental cleanup in coastal waters or reservoirs.
Figure 5. Intrinsic phosphatase activity of the natural bacterial assemblage from the Newport River Estuary, North Carolina transect (September 2016) as a percentage of heterotrophic bacterial production vs salinity (PSU).
C/P ratio with low growth rate could occur in the 0 PSU bog water relative to mid-estuary.54,55 When further comparing the relationship between this normalized intrinsic phosphatase activity to a measure of aromatic, humic character of DOC in these natural waters, there is a strong relationship with UV absorptive parameters indicative of these properties,56 specifically a254 (R2 = 0.999) and specific UV absorption at 254 nm (SUVA254; R2 = 0.979; Figure 6). These results are supported by the δ13C values of DOC, which generally show a transition from humic-rich DOC in bog water to coastal marine DOC in Onslow Bay.
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MATERIAL AND METHODS LCA and LCA-AP Nanotubes. LCA nanotubes were prepared by the method of Terech et al.26 with modifications described by Lu et al.66 in 50 mM Tris buffer (TB). LCA-AP 2201
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Figure 6. Intrinsic phosphatase activity of the natural bacterial assemblage from the Newport River Estuary, North Carolina transect (September 2016) as a percentage of heterotrophic bacterial production vs a254 (A) and SUVA254 (L mg C−1 m−1) (B).
Figure 7. Newport River Estuary system water sample locations from the fresh headwaters at the Newport Cypress bog (NC-0, -0A; inset A), through the tidal creeks (NC-5) and estuarine mixing zone (NC-5A; NC-11, -11A; NC-21, -21A) to offshore of Beaufort Inlet (NC-36), Bogue Sound (NC-M1A) and Bogue inlet (NC-36A), NC in the mid-Atlantic (inset B). Station numbers represent the practical salinity units (PSU). Locations for August 2015 (green) and September 2016 (yellow; Google Maps).
was prepared by the method of Lu et al.66 except that alkaline phosphatase (Sigma# A2356, bovine intestinal mucosa, 2 mg mL−1 stock concentration) was used as the encapsulated enzyme. The amount of AP encapsulated in LCA nanotubes (stock concentration: 132 μg AP mg−1 LCA in 1 mL prior to dilution) was determined using a Pierce BCA Protein Assay Kit, also as described by Lu et al.27 Nanotube formation is further demonstrated by atomic force microscopy (AFM), confocal, and transmission electron microscopy (TEM) images
and optical spectra in the Supporting Information (Figures S1−S5). Environmental Sampling. Natural environmental water samples were collected along two transects of the NRE to offshore of the Lower Outer Banks and Bogue Sound, North Carolina, in August 2015 and September 2016. NRE is about 15 km long by 4−5 km wide with a mean depth of 1 m67 with about half its volume exchanging each tidal cycle36 and a mean residence time of ca. 4−10 days.64 Surface water was collected 2202
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Montgomery et al.46 by adding LCA alone or LCA-AP (5 μL) to the microfuge tube precharged with 3H-leucine just prior to incubation with whole water (unfiltered) transect sample (1 mL). Nanotube solutions (0.1% LCA in TB, pH 8.0) were diluted with TB prior to addition to the charged microfuge tubes giving final concentrations of 0.32, 3.2, 32, and 320 μM of LCA carbon (LCA-C), which bounded the maximum used in microtiter kinetics assays (38 μM) and is likely much higher than would be potentially used in an environmental application. Dissolved Organic Carbon Properties. Concentration, stable C isotope (δ13C-DOC) value, and UV absorptive properties of DOC were measured according to established methods.69,70 Calibration and normalization of DOC and δ13C-DOC values were achieved using prepared solutions of caffeine and sucrose, which bracket a range of δ13C-DOC values from −30 to −10‰.70 DOC light absorption at 254 nm was made on water blank-corrected absorbance values, which were converted to Napierian absorption coefficients (m−1). Specific UV absorption at 254 nm (SUVA254) was used to estimate the aromatic content of DOC and was computed as a sample’s decadic absorption at 254 nm divided by its DOC concentration, with units of L mg C−1 m−1.56 DOC concentration values have a coefficient of variation of
