Article pubs.acs.org/est
Environmental Photoinactivation of Extracellular Phosphatases and the Effects of Dissolved Organic Matter Elisabeth M. L. Janssen* and Kristopher McNeill* Institute of Biogeochemistry and Pollutant Dynamics, ETH Zurich, CH-8092, Zurich, Switzerland S Supporting Information *
ABSTRACT: Alkaline phosphatases are ubiquitous extracellular enzymes in aquatic systems and play a central role in the biogeochemical cycling of phosphorus. Yet, the photochemical stability of phosphatase and effects of natural organic matter (DOM) are not completely understood. We demonstrate that phosphatase activity in natural biofilm samples decreased during sunlight exposure similar to well-defined bacterial phosphatase solutions. Direct photoinactivation was slowed by more than 50% in the presence of redox-active dissolved organic matter (DOM, 10 mgC L−1) or a model antioxidant (esculetin, 50 μM), even after light screening effects had been accounted for. Thus, DOM can not only inhibit enzymes (in the dark) or sensitize photodegradation by producing photochemically produced reactive intermediates but can also significantly quench direct photoinactivation of phosphatase. Our data further suggest that direct photooxidation of tryptophan residues within the protein structure are significantly involved in the photoinactivation of phosphatase because a loss of tryptophan-like fluorescence paralleled photoinactivation kinetics and because DOM acted as an antioxidant toward photoinactivation, a phenomenon recently established for the photooxidation of freely dissolved tryptophan. Thus, photoinactivation of phosphatase can be significantly slowed in the presence of naturally occurring antioxidants like DOM. The mechanistic link between tryptophan photooxidation and inactivation of phosphatase may have applicability to other extracellular enzymes but remains to be established.
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INTRODUCTION Life requires phosphorus as an essential nutrient, and its availability can be a limiting factor for growth and reproduction in environmental systems. Fast uptake rates of dissolved inorganic phosphorus can result in low ambient phosphorus concentrations and fast phosphorus fluxes from source to organism.1 Under phosphorus-limiting conditions, dissolved organic phosphorus becomes the major phosphorus source.1,2 However, dissolved organic phosphorus is not immediately bioavailable, as it needs to be converted to inorganic phosphorus, for example by enzymatic hydrolysis. Such transformation rates of dissolved organic phosphorus are of great interest to better understand the phosphorus turnover in aquatic systems. Phosphatases are a family of enzymes that catalyze the hydrolysis of phosphomonoesters to provide inorganic phosphorus. A diverse range of organisms including bacteria, phytoplankton, and zooplankton utilize phosphatase for their phosphorus supplies,3 and phosphatase activity can be detected in freshwater systems 3−5 as well as marine waters. 6 Phosphatases can be located within cells, but also on cell surfaces as ectoenzymes or extracellular as freely dissolved enzymes. These extracellular phosphatases can be present as a result of cell lysis or active release. Extracellular enzymes play a central role in the biogeochemical cycling in soils, sediments and aquatic systems. Research is © 2014 American Chemical Society
just beginning to investigate the numerous environmental factors that act on the stability and fate of extracellular enzymes. Extracellular enzymes are targets of microbial degradation and abiotic transformation, including photolysis reactions and sorption processes. Two effects of dissolved organic matter (DOM) on enzyme activity have been commonly observed: inhibition (in the dark)7−9 and increased inactivation by indirect photochemical processes.10 A third processes involves quenching of photoinactivation, which is of special interest in the presented work. It has been observed that photoinactivation rates of phosphatases during sunlight and UVB light exposure were slower in the presence of DOM and the authors hypothesized that DOM complexes with phosphatases and protects them from damage by light, but the underlying mechanism was not provided.9 In the present study, we were particularly interested in extracellular phosphatase within the photic zone of aquatic systems, released by algae or bacteria and how exposure to light and interactions with DOM affect the enzyme activity. Extracellular aquatic enzymes can be inactivated by direct photolysis due to damage of light-absorbing amino acids, Received: Revised: Accepted: Published: 889
August 27, 2014 December 10, 2014 December 11, 2014 December 11, 2014 DOI: 10.1021/es504211x Environ. Sci. Technol. 2015, 49, 889−896
Article
Environmental Science & Technology
Switzerland (lat 47.368°, long 8.497°) in June 2014. The biofilm was removed from the vertical face of stones in the creek using a nylon brush, suspended with creek water, and incubated at 4 °C in open tubes for 24 h. The suspensions were centrifuged (5 min, 5000 rpm) and filter sterilized (0.2 μm, sterile cellulose acetate membrane), and the pH was adjusted from originally pH 7.7−8.3 to pH 8.5. The total organic carbon content was determined as 25 mgC L−1 (Shimadzu Corporation, TOC-L analyzer). Filtered samples were stored at 4 °C for less than 3 h before use. Enzyme Activity Measurements. Phosphatase activity was determined by hydrolysis of the phosphomonoester 4methylumbelliferone phosphate (MUP). Two enzyme activity assays were used, (1) based on bulk fluorescence measurements and (2) based on HPLC separation and fluorescence detection of the assay’s product. Bulk Fluorescence Activity Assay. Sample aliquots (120 μL) were transferred to a 96-well plate (Nunc-Immuno MicroWell, clear, Sigma) in triplicate, and MUP substrate solution (20 μL, buffered at pH 8.5) was added. For photolysis experiments, 140 μM and 70 μM MUP final concentrations were sufficient to reach the maximum substrate conversion rate (Vmax) for biofilm samples and ECAP solutions (0.05 nmol L−1), respectively. The fluorescence of the product, 4-methylumbelliferone (MUF), was monitored with an excitation wavelength of 360 nm and emission wavelength of 460 nm and was read every minute for 15 min (Synergy HT Microplate reader, Bio-TEK). The fluorescence was converted to MUF concentration by external calibration with MUF solutions to yield maximum enzyme activity, Vmax (μM MUF s−1). The first-order inactivation rate constant of phosphatase, k (s−1), was assessed by linear regression of ln(Vmax/Vmax,0) versus irradiation time (s). The bulk fluorescence method allowed higher sample throughput and high data resolution based on following the kinetics of product formation. However, the fluorescence of the MUF product can be affected by phenolic compounds (DOM, model antioxidants) in solution.12 HPLC-based Activity Assay. The production of MUF by phosphatase was quantified following HPLC separation. Therefore, 80 μL of sample was added to 20 μL of MUP (0.5 mM, pH 7.5), and MUF product formation was stopped after exactly 20 min by inhibiting the enzyme with 1.4 mM phosphate and lowering the pH to 3.0 (addition of 20 μL of KH2PO4 solution). The concentration of MUF was measured by ultra performance liquid chromatography (UPLC) separation using 50% phosphate buffer (pH 7.5, 20 mM) and 50% methanol with 0.15 mL min−1 flow rate on a C18 column (Waters Aquity, BEH130 C18, 1.7 μm, 2.1 × 150 mm) and with fluorescence detection at an Excitation/Emission wavelength of 320/450 nm. Data on MUF stability and enzyme inhibition can be found in the Supporting Information (SI, Figure S1). For the natural biofilm samples, the bulk fluorescence activity assay was used. Even though natural phenolic constituents may interfere with the fluorescence measurement, this method allowed fast processing of the natural samples to guarantee that the samples did not deteriorate due to long storage times. On the other hand, for experiments with bacterial phosphatase solutions (ECAP), the HPLC-based assay was used because fresh sample solutions could be prepared daily. For the determination of the reaction rate constant of ECAP with singlet oxygen (1O2), the bulk fluorescence method was used
namely tryptophan, tyrosine, and the disulfide cystine. Among these, the indole side chain of tryptophan, if present, is the dominant amino acid based chromophore in proteins absorbing in the UVB range of the solar light spectrum. We previously demonstrated that DOM plays dual roles in the photodegradation of monomeric tryptophan acting as a sensitizer and quencher.11 Direct photoreaction of tryptophan as well as reaction of tryptophan with excited triplet-state DOM form an intermediate tryptophan radical cation, which can be converted back to tryptophan by suitable electron or hydrogen atom donors. With a better mechanistic understanding of tryptophan’s aquatic photochemistry, the question arises whether DOM can similarly affect the photodegradation of tryptophan residues within biomacromolecular structures of proteins. Here, we report an investigation into the effects of DOM on photoinactivation of phosphatase. UVB-light-mediated inactivation and decay of tryptophan-like fluorescence of phosphatase in natural biofilm samples was demonstrated, which led to the hypothesis that tryptophan photochemistry drives inactivation. We tested this hypothesis by adding known quenchers of tryptophan photooxidation (redox-active DOM and a model antioxidant) during UVB light exposure to assess whether those can slow photoinactivation of phosphatase from structurally well-characterized bacterial phosphatase under controlled solution conditions. We demonstrated that DOM quenches photoinactivation rates of phosphatase, and this effect correlates with reduced decay of tryptophan-like fluorescence of phosphatase, but the causative relationship may be quite complex.
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MATERIAL AND METHODS Reagents and Solutions. The following reagents were used as received: Escherichia coli alkaline phosphatase (ECAP; Sigma Life Science, 62.77 units mg−1 protein, 3.25 mg mL−1 in glycine buffer); 4-methylumbelliferyl phosphate, 4-methylumbelliferone, tryptophan (>98% purity), cystine (cysteine disulfide), and tyrosine (Sigma); Rose Bengal, Aldrich Humic Acid (AHA), 6,7-dihydroxycoumarin (esculetin, ≥ 98% purity), histidine (purity >99%), phenylalanine (purity >99%), ammonium acetate (>99.99% purity, Sigma-Aldrich), furfuryl alcohol (FFA, distilled prior to use), potassium dihydrogen phosphate (puriss.), and acetic acid (Fluka); acetonitrile, methanol (≥99.9% purity), and disodium tetraborate decahydrate (Merck KGaA,); tris(hydroxylmethyl)aminomethane (Tris, Aldrich, ≥ 99% purity); and potassium phosphate dibasic (Riedel-de-Haën, puriss). All solutions were prepared with nanopure water (resistivity >18 MΩ cm, Barnstead NANOpure Diamond). Solutions of ECAP were prepared daily, and experiments were carried out in buffered nanopure water (Tris buffer, 10 mM, ionic strength 30 mM with NaCl, pH 7.5). Waskish Peat Organic Matter (WKP) and Suwannee River Fulvic Acid II (SRFA II) were obtained from the International Humic Substances Society (IHSS; St. Paul, MN, USA). Organic matter solutions were prepared by dissolving approximately 250 mgOM L−1 in nanopure water at pH 10.0 by adding sodium hydroxide and sonicating the solutions intermittently until the pH remained stable. The DOM solutions were then adjusted to approximately pH 7 with hydrochloric acid, filter sterilized (0.2 μm, sterile cellulose acetate membrane), and either used the day of preparation or frozen until use. Biofilm Sampling and Preparation. Biofilm samples were collected at the Döltschibach, a shallow creek in Zurich, 890
DOI: 10.1021/es504211x Environ. Sci. Technol. 2015, 49, 889−896
Article
Environmental Science & Technology
Figure 1. (A) Phosphatase activity change of natural biofilm samples during irradiation with a solar simulator (blue triangles), a solar simulator with a 455 nm cutoff filter (black circles), a UVA light source (green squares), and a UVB light source (orange diamonds). Inset shows the pseudo-firstorder inactivation rate constants. (B) Degradation rates of freely dissolved tryptophan (diamonds), cystine (cysteine disulfide: circles), and tyrosine (triangles) under the same UVB light conditions and in borosilica tubes as used for the biofilm samples. Error bars represent one standard deviation, and activity was assessed by the bulk-fluorescence method.
1.7 μm, 2.1 × 150 mm) with an injection volume of 5 μL. For furfuryl alcohol, a 75% sodium acetate buffer (pH 5.9, 15.6 mM) and 25% acetonitrile with a 0.15 mL min−1 flow rate and absorbance detection at 219 nm absorbance was used. Tryptophan and tyrosine were analyzed under the same chromatographic conditions and detected by fluorescence with excitation/emission wavelengths of 280/350 nm and 270/350 nm, respectively. Cystine and histidine were analyzed upon AQC-derivatization as described elsewhere.14 Briefly, the same UPLC with a C18 column was used with acetate butter (A) and acetonitrile (B) at a 0.35 mL min−1 flow rate and the following gradient: 0−0.5 min, 90%A−10%B; 0.5−2 min, 80% A−20%B; 2−11 min, 65%A−35%B; 11−12 min, 90%A−10%B. The AQC-derivatized amino acids were detected by fluorescence with an excitation/emission wavelength of 245/395 nm. Phenylalanine was used as an internal standard. All first-order degradation rate constants, kobs (s−1), were assessed by linear regression of log-transformed signals normalized by initial values (ln(A/A0)) versus irradiation time. All data reported herein were corrected for light screening by DOM or esculetin, and details are provided in the Supporting Information (SI, Table S1).
because no interference with MUF fluorescence measurements was apparent. Photolysis Experiments. Solutions were exposed to light generated by (1) a Xe lamp (Newport, at 300 W) simulating the solar light spectrum and with and without a low-pass filter (
