Accepted Manuscript Evaluating fluorescence spectroscopy as a tool to characterize cyanobacteria intracellular organic matter upon simulated release and oxidation in natural water Julie A. Korak, Eric C. Wert, Fernando L. Rosario-Ortiz PII:

S0043-1354(14)00688-5

DOI:

10.1016/j.watres.2014.09.046

Reference:

WR 10909

To appear in:

Water Research

Received Date: 24 June 2014 Revised Date:

27 September 2014

Accepted Date: 30 September 2014

Please cite this article as: Korak, J.A., Wert, E.C., Rosario-Ortiz, F.L., Evaluating fluorescence spectroscopy as a tool to characterize cyanobacteria intracellular organic matter upon simulated release and oxidation in natural water, Water Research (2014), doi: 10.1016/j.watres.2014.09.046. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Unique  Fluorescence   250 250 Signatures?   300 400 400 500 500 300 400 400 500 500 300 300

550 550

0.09 0.09

00

00

2.6 2.6

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1.8 1.8

0.66 0.66

Lyngbya Lyngbya

1 0.8

0.6 0.6

0.18 0.18

0.4 0.4

0.09 0.09

0.2 0.2

00

300 400 400 500 500 300

0.75 0.75

0.2 0

b)

CRW CRW 0.27 0.27

0.6

300 0.4 400 500 500 300 400

P2 Fmax/Fmax Fraction Remaining 0

600 600

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0.3 0.3

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0.18) a) 0.18

P1 Fmax/Fmax0

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Readily  Oxidized?   0

0.5 0.50.5

1

1.5 1010

2

1

Species Oxidants 55 MA NH2Cl 500 500 OSC FCl 0.6 O3 LYG 00 00 00 00 450 450 ClO2 550 600 600 650 650 700 700 550 550 600 600 650 650 700 700 550 550 600 600 650 650 700 700 550 550 600 600 650 650 700 700 550 0.4 EmissionWavelength, Wavelength,nm nm Emission 0.9 0.9

Em (nm)

0.33 0.33

c)

0

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Humic Region Humic Region

Excitation Wavelength, nm

Excitation Wavelength, nm Ex (nm)

Pigment Region Pigment Region

Cyanobacteria  IOM  

400 400

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Microcystis Microcystis 450 450

Cyanobacteria   Intracellular     Organic  Ma3er   Oscillatoria Oscillatoria (IOM)   0.27 0.9 0.9 0.27

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0.25 0.25

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Evaluating fluorescence spectroscopy as a tool to characterize cyanobacteria

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intracellular organic matter upon simulated release and oxidation in natural

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water

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Julie A. Korak1, Eric C. Wert2 and Fernando L. Rosario-Ortiz1* 1

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Department of Civil, Environmental and Architectural Engineering, 428 UCB, University of Colorado, Boulder, CO 80309, USA

Southern Nevada Water Authority (SNWA), P.O. Box 99954, Las Vegas, NV 89193-9954 USA

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*Corresponding author: [email protected] Keywords: Cyanobacteria, intracellular organic matter, fluorescence, phycocyanin, oxidation, DOM, fluorescence index

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Abstract

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Intracellular organic matter (IOM) from cyanobacteria may be released into natural

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waters following cell death in aquatic ecosystems and during oxidation processes in drinking

14

water treatment plants. Fluorescence spectroscopy was evaluated to identify the presence of

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IOM from three cyanobacteria species during simulated release into natural water and following

16

oxidation processes (i.e. ozone, free chlorine, chloramine, chlorine dioxide). Peak picking and

17

the fluorescence index (FI) were explored to determine which IOM components (e.g., pigments)

18

provide unique and persistent fluorescence signatures with minimal interferences from the

19

background dissolved organic matter (DOM) found in Colorado River water (CRW). When IOM

20

was added to ultrapure water, the fluorescence signature of the three cyanobacteria species

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showed similarities to each other. Each IOM exhibited a strong protein-like fluorescence and

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fluorescence at Ex 370 nm and Em 460 nm (FDOM), where commercial fluorescence sensors

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monitor. All species also had strong phycobiliprotein fluorescence (i.e. phycocyanin or

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phycoerythrin) in the higher excitation range (500-650 nm). All three IOM isolates had FI values

25

greater than 2. When IOM was added to CRW, phycobiliprotein fluorescence was quenched

26

through interactions between IOM and CRW-DOM. Mixing IOM and CRW demonstrated that

27

protein-like and FDOM intensity responses were not a simple superposition of the starting

28

material intensities, indicating that interactions between IOM and CRW-DOM fluorescing

29

moieties were important. Fluorescence intensity in all regions decreased with exposure to ozone,

30

free chlorine, and chlorine dioxide, but the FI still indicated compositional differences compared

31

to CRW-DOM. The phycobiliproteins in IOM are not promising as a surrogate for IOM release,

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because their fluorescence intensity is quenched by interactions with DOM and decreased during

33

oxidation processes. Increases in both FDOM intensity and FI are viable qualitative indicators of

34

IOM release in natural waters and following oxidation and may provide a more robust real-time

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indication of the presence of IOM than conventional dissolved organic carbon or UV absorbance

36

measurements.

37 38

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Some cyanobacteria produce cyanotoxins, such as microcystin-LR, that may present a human

40

health risk (Hitzfeld et al., 2000; Latifi et al., 2008; Pearson et al., 2010; Sinclair and Hall, 2008;

41

WHO, 2003). Other species produce the metabolites geosmin and 2-methylisoborneol (MIB) that

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create taste and odor episodes (Juttner and Watson, 2007; Smith et al., 2008; Watson, 2004;

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2003). These metabolites may be released into natural waters along with intracellular organic

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matter following cell death in aquatic ecosystems and during oxidation processes in drinking

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water treatment plants.

Introduction

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Cyanobacteria blooms are an area of concern for protecting drinking water source quality.

Some drinking water treatment plants apply oxidants upstream of

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physical removal processes to meet various treatment objectives, which has the potential to

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damage cell integrity and release metabolite-containing intracellular organic matter (IOM) (Ding

48

et al., 2009; Latifi et al., 2008; Wert et al., 2013; 2014; Wert and Rosario-Ortiz, 2013; WHO,

49

2003; Zamyadi et al., 2012a). The release of cyanobacteria-derived organic matter is difficult to

50

quantify, because it is likely found at low concentrations compared to background dissolved

51

organic matter (DOM) and may be difficult to detect by dissolved organic carbon (DOC) or

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absorbance measurements (Wert et al., 2014).

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Monitoring the fluorescent components of IOM may be a valuable surrogate of interest

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for drinking water utilities. By targeting specific excitation and emission wavelengths,

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fluorescence can selectively measure different optically-active moieties. Analysis can be

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performed rapidly on bench-top fluorometers with samples collected along a treatment train or

57

measured in real-time using fluorescence probes (Carpenter et al., 2012; Downing et al., 2012;

58

Zamyadi et al., 2012b). A monitoring strategy that can detect the release of IOM within a

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drinking water treatment plant could be implemented during cyanobacteria blooms to determine

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if preoxidation processes are compromising cell integrity and potentially releasing cyanotoxins,

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taste and odor metabolites and/or disinfection byproduct precursors (Wert and Rosario-Ortiz,

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2013).

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A few studies have explored the fluorescence characteristics of isolated cyanobacteria

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IOM in ultrapure water matrices (Fang et al., 2010; Henderson et al., 2008; Li et al., 2011;

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Ziegmann et al., 2010). While recognizing that IOM has a fluorescent signature distinct from

66

DOM, no studies have investigated whether these fluorescence fingerprints have utility for

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monitoring IOM release during preoxidation processes. Taking this next step requires

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determining if the fluorescence signatures change through interactions with DOM present in the

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water before cell lysis and determining the effect of oxidation on IOM fluorescence

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characteristics. Of these studies, only one study investigated the fluorescent properties of

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phycobiliproteins common to all cyanobacteria (Ziegmann et al., 2010).

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Phycobilisomes are light harvesting complexes attached to the thylakoid membrane that

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expand the range of wavelengths the organism can utilize for photosynthesis. They absorb light

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in the visible range predominately from 550 to 650 nm, where chlorophyll a cannot absorb light

75

(Bryant, 1994). Phycocyanin and phycoerythrin are two fluorescent phycobiliproteins in

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phycobilisomes that are commonly used to characterize cyanobacteria populations in natural

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systems (Brient et al., 2008; Izydorczyk et al., 2009; 2005; McQuaid et al., 2011; Watras and

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Baker, 1988). On-line fluorescence monitoring of cyanobacteria has mainly focused on probes

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that measure in vivo fluorescence of chlorophyll a and phycocyanin and correlate responses to

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cell concentrations or biomass (Bastien et al., 2011; Brient et al., 2008; Gregor et al., 2007;

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Heaney, 1976; Stewart and Farmer, 1984; Watras and Baker, 1988; Zamyadi et al., 2012c). The

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ratio of in vivo chlorophyll a to phycocyanin fluorescence has been related to the relative

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abundance of cyanobacteria compared to other algal phyla (Watras and Baker, 1988). In vivo

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measurements have also been correlated to total extractable chlorophyll a and phycocyanin

85

(Zamyadi et al., 2012c). For a thorough discussion, refer to the Supplemental Information Text 1.

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Since phycobiliproteins are limited to a few phyla (i.e., cyanobacteria, rhodophyta and

87

cryptophyta), they are important for distinguishing cyanobacteria from other algal phyla, such as

88

green algae (Bryant, 1994). These unique fluorescent signatures occur in a wavelength region

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where DOM does not fluoresce strongly and may be good candidates to detect the release of

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cyanobacteria-derived organic matter under oxidizing conditions. No investigations have been

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performed to evaluate if fluorescence probes designed to measure in vivo phycocyanin

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fluorescence are also suitable to detect IOM released into natural waters during water treatment

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operations, such as oxidation processes. The study objectives were to (1) identify the fluorescence signature of IOM, phycocyanin,

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and phycoerythrin in ultrapure water, (2) evaluate matrix interferences between the IOM

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fluorescence signature and background DOM, and (3) determine if the fluorescence signature

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persists after oxidation processes in drinking water treatment by evaluating IOM oxidation in

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ultrapure water using ozone, free chlorine, chlorine dioxide and monochloramine. After

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accomplishing these objectives, the feasibility of fluorescence as a surrogate for IOM in a

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heterogeneous DOM matrix can be determined.

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2

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2.1

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Texas, Austin, TX), Oscillatoria sp. (OSC) (LM 603d, Metropolitan Water District of Southern

105

California (MWDSC), La Verne, CA), and Lyngbya sp. (LYG) (SDC 202d, MWDSC, La Verne,

106

CA) were selected for the study based upon their occurrence in source water supplies,

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availability of an axenic culture, ability to produce odorous or toxic metabolites, and different

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cell morphology. MA and LYG are both planktonic species, and OSC is a benthic species. MA is

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a unicellular species and confirmed producer of microcystin-LR; OSC and LYG are filamentous

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species and confirmed producers of MIB and geosmin, respectively (Wert and Rosario-Ortiz,

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2013). Each cyanobacteria was cultured in batches. Details regarding the culturing methods,

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growth medias and growth curves can be found elsewhere (Wert et al., 2013).

Methods

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Cell Cultures and IOM Extraction Microcystis aeruginosa (MA) (LB 2385, Culture Collection of Algae at the University of

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MA, OSC, and LYG cells were harvested during the late exponential growth phase (28

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days) when intracellular metabolites are expected to be the greatest (Pietsch et al., 2002;

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Watanabe et al., 1989; Watanabe and Oishi, 1985). The cells were separated from the growth

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media by centrifugation (1900 rpm, 14 min). The supernatant containing the growth media and

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the extracellular organic matter (EOM) was discarded. The remaining cell pellet was rinsed with

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10 mM phosphate buffered lab-grade water (pH = 7.5), centrifuged and the supernatant discarded

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with any residual growth media and/or EOM. For OSC and LYG, a mortar and pestle was used

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to grind the cell pellet to improve IOM extraction from these filamentous cyanobacteria. Three

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freeze-thaw sequences (-77 °C freezer, 35 °C water bath) and sonication (60 min in an ice bath)

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were used to release the IOM from the cells (Wert and Rosario-Ortiz, 2013). Filtration separated

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the cell debris from dissolved IOM (combusted and rinsed 0.7 µm GF/F, Whatman). The filtrate

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was used as the extracted IOM standard for each species.

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Phycocyanin (#P2172, Sigma Alrich, St. Louis, MO) and phycoerythrin (#52412, Sigma

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Aldrich, St. Louis, CO) standards were acquired to characterize the fluorescence interactions and

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oxidation behaviors for model compounds. The phycocyanin standard was a 40% w/w

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lyophilized powder from Spirulina sp. preserved with sucrose, dithilerythritol and sodium azide.

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The phycoerythrin standard was dissolved in 150 mM sodium phosphate with 60% ammonium

130

sulfate, 1 mM EDTA and 1 mM sodium azide.

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2.2

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concentrations of 2 mgC/L for MA and 1 mgC/L for OSC and LYG. IOM was also spiked into 0.7

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µm GF/F filtered Colorado River Water (CRW) at varying concentrations. The water quality

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characteristics of CRW were as follows: DOC of 2.65 mgC/L, pH of 8.0, alkalinity of 138 mg/L

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Experimental Methods IOM standards were diluted in lab grade water with 10 mM phosphate buffer (pH=7.5) to

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as CaCO3 and absorbance at 254 nm (UV254) of 0.052 cm-1. Similarly, phycocyanin and

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phycoerythrin standards were prepared in phosphate buffer for oxidation studies and spiked to

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CRW at varying concentrations. All experiments and optical measurements were conducted at

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room temperature (~20°C).

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The oxidation methods have been published in more detail elsewhere (Wert et al., 2014;

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2013). Briefly, the three IOM isolates in phosphate buffer and the CRW sample were exposed to

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four oxidants: ozone, free chorine, chlorine dioxide and chloramines as batch processes at room

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temperature (20-25°C). Ozone stock solutions were prepared by dissolving gaseous ozone at a

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concentration of 80 mg/L (Standard Methods 4500-O3) (APHA et al., 1998) into deionized water

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at 2°C and adding an aliquot to the sample (Wert et al., 2009). Chlorination experiments used a

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5.6% liquid sodium hypochlorite solution (NaOCl, Fisher Scientific, USA). Chloramination

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experiments used a preformed chloramine solution by first adding ammonia (29% ammonium

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hydroxide solution, J.T. Baker, USA) followed by sodium hypochlorite (5.6% sodium

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hypochlorite, NaOCl, Fisher Scientific, USA) using a chlorine:ammonia ratio of 3:1. The pH of

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the preformed chloramine solution was above 10 favoring monochloramine at the time of

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addition. Chlorine dioxide experiments were performed using a 3,000 mg/L solution (CDG

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Environmental, Bethlehem, PA, USA). For the IOM oxidation studies, ozone and chlorine

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dioxide were dosed at oxidant to DOC mass ratios between 0 and 1. Free chlorine and

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chloramine were dosed at ratios between 0 and 2. Residual ozone decayed after 30 minutes and

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no quenching agent was required. Samples with chlorine-based oxidants and their accompanying

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controls were quenched with 80 mg/L of sodium thiosulfate after 120 min. The effects of

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quenching agents on fluorescence are discussed in SI-Text 2.

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2.3

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Technologies, CA) in a quartz cell with a 1 cm path length. Spectra were baseline corrected to

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lab grade water. The average relative percent difference between duplicate absorbance

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measurements at 254 nm was 2%, and all values were below 5%. Dissolved organic carbon

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(DOC) was measured using a TOC analyzer (Shimadzu, Columbia, MD) following SM5310B.

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The coefficient of variance (CV) for DOC concentration for all 2 mgC/L standards throughout the

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study was 2.4% (n=58). excitation-emission

matrices

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(EEMs)

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Analytical Methods UV-VIS absorbance spectra were measured from 200 to 800 nm (Cary 100, Agilent

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were

collected

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spectrofluorometer (Fluoromax 4, John Yvon Horiba, NJ). EEMs were collected as a series of

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emission scans (300 nm to 800 nm in 2 nm increments) at set excitation wavelengths (250 nm to

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700 nm in 10 nm increments). A 5 nm bandpass was used for both excitation and emission

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monochromators, and the integration time was 0.25 s. All EEMs were collected in ratio mode

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(signal divided by reference) and incorporated instrument-specific correction factors. All EEMs

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were corrected and normalized following published methods (Murphy et al., 2010). Briefly,

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EEMs were blank subtracted to minimize Raman scattering, inner filter corrected following

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Lakowicz (2006) and normalized to the Raman area of 18.2MΩ-cm ultrapure water at an

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excitation of 350 nm yielding Raman Units (RU) (Lawaetz and Stedmon, 2009). All samples had

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an optical density less than 0.2 at 254 nm. All samples were stored filtered at 4°C in the dark and

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analyzed within 10 days.

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Several fluorescence metrics were used to analyze the EEMs. Fluorescent dissolved

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organic matter (FDOM) intensity is defined as a fixed wavelength intensity located at excitation

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370 nm and emission 460 nm to follow the wavelength pairs commonly used in FDOM sensors

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for online monitoring applications (Downing et al., 2012; Fleck et al., 2013; Kraus et al., 2010;

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Saraceno et al., 2009). Peak P intensity is the maximum intensity in the region extending from

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260 to 290 nm excitation wavelengths and 300 to 350 nm emission wavelengths. This region has

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conventionally been analyzed by splitting the regions into Peaks B and T, which are both related

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to amino acid functional groups (Coble, 1996). This region was analyzed as a whole designated

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Peak P, because the EEM contours exhibited single peaks that extended continuously through

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both regions. Additionally, potential online sensors would be unlikely to have the spectral

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resolution to differentiate between regions (Carpenter et al., 2012). Phycocyanin fluorescence is

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represented by the intensity measured at excitation 610 nm and emission 640 nm (Bryant, 1994;

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Debreczeny et al., 1993). Phycoerythrin fluorescence is defined as the intensity at 560 nm

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excitation and 578 nm emission (Bryant, 1982). Specific peak intensity is the intensity

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normalized to the DOC concentration presented in the units RU L mgC-1. The fluorescence index

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(FI) is defined as the ratio of intensities at 470 nm and 520 nm at an excitation of 370 nm and has

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been correlated to sample aromaticity (Cory et al., 2010; McKnight et al., 2001). These metrics

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are depicted in Figure 1.

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EEMs of each isolated IOM in phosphate buffer showed two main regions with strong

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fluorescence signals (Figure S-1). Figure 1 breaks the full EEM apart displaying each region

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separately. The region displayed in the top row (Excitation 250-450 nm, Emission 300-560 nm)

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will be referred to as the Low Wavelength Range. The region presented in the bottom row (Ex

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450-700 nm, Em 550-700 nm) will be referred to as the High Wavelength Range.

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3

Results and Discussion

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3.1

Isolated IOM Fluorescence Characterization in Ultrapure Water

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3.1.1

Qualitative Fluorescence Comparisons Comparing IOM EEMs with those of the background DOM from CRW (CRW-DOM)

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reveals that several fluorescence metrics could be used as an IOM surrogate (Figure 1). The

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presence of distinct peaks suggest that the fluorescing moieties in IOM are more homogeneous

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than CRW-DOM and EOM, which both show a continuum of fluorescence in the Low

208

Wavelength Range (see methods for nomenclature) similar to humified material (Qu et al., 2012).

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This section identifies fluorescence indicators that are common to all three cyanobacteria

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examined and should be evaluated further as IOM surrogates.

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All three cyanobacteria species examined exhibited strong fluorescence signals in the Peak

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P region, as outlined in Figure 1. Fluorescence in this region has been commonly associated with

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protein-like functional groups, because the phenolic functional group in tyrosine and the indolic

214

functional group in tryptophan both fluoresce in this region (Coble, 1996; Lakowicz, 2006).

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Humic-associated polyphenolic compounds have also been shown to fluoresce in this region

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(Hernes et al., 2009; Maie et al., 2007). Given that IOM is of microbial origin and has been

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shown to contain fewer phenolic functional groups compared to allochthonous organic matter

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(Nguyen et al., 2005), it is likely that the fluorescence in this region is mainly associated with

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proteinaceous material. Figure 1 and Table 1 highlight some compositional differences between

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cyanobacteria species. The Peak P emission for MA IOM occurs at 330 nm, which corresponds

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closely with the indolic functional group tryptophan (Lakowicz, 2006; Reynolds, 2003) and has

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been assigned the conventional name Peak T (Coble, 1996; Fellman et al., 2010). The peak

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emission intensity for OSC and LYG IOM is blue-shifted to lower wavelengths with a shoulder

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that extends to the longer emission wavelengths. The peak location in OSC and LYG is

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consistent with the presence of the phenolic functional group in tyrosine (Lakowicz, 2006) and

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falls in the conventionally assigned Peak B region. Even though tyrosine is less commonly

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measured by fluorescence due to its efficient energy transfer to tryptophan (Lakowicz, 2006),

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distinct contributions from the phenolic functional group is evident in IOM. Free tyrosine amino

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acids have been measured in MA IOM (Fang et al., 2010) and several phycobiliproteins have

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more tyrosine residues relative to tryptophan (Glazer and Bryant, 1975; Glazer and Fang, 1973;

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Ong and Glazer, 1991). Although fluorescence peaks shifted between the conventionally-

232

assigned Peak B and T regions, only the maximum intensity in the larger, encompassing region

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(Peak P) will be analyzed in subsequent sections for simplicity. Further discrimination between

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the B and T regions is not necessary to meet the study objectives, and online probes currently do

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not offer this level of resolution (Carpenter et al., 2012).

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All three species exhibited fluorescence maxima at an excitation wavelength of 370 nm

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and emission wavelengths ranging from 456 to 462 nm, which coincides with the wavelengths

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used in some FDOM sensors (Downing et al., 2012). In addition, there is a second peak at a

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lower excitation wavelength (250 nm) with the same emission wavelength suggesting that the

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fluorescence may be related to each other by Kasha’s rule (Kasha, 1950). This signal is red-

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shifted compared to the Peak A region for humic-like material (Coble, 1996). This signal is

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likely related to the major intrinsic fluorophore in all cells: nicotinamide adenine dinucleotide

243

(NADH or NAD(P)H) (Lakowicz, 2006). NAD(P)H is a coenzyme in all cell metabolism and

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has a characteristic fluorescence emission at 460 nm (Lakowicz, 2006). Pure NAD(P)H in water

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has absorption maxima at 260 and 340 nm (Wolfbeis, 1985), but the differences observed here

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may be due to interactions with other constituents that can affect optical signatures (Lakowicz,

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2006; Wolfbeis, 1985). Fluorescence in the same region has been attributed to NAD(P)H in

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fluorescence EEM analysis of other cyanobacteria but is not cyanobacteria specific (Dartnell et

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al., 2010). This fluorescence signature occurs in the same region where FDOM intensity and FI

250

are quantified and are viable surrogates for monitoring IOM release. These fluorescence metrics

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are also of particular interest, because they have already been incorporated into online, in-situ

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sensors (Carpenter et al., 2012; Downing et al., 2012).

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Finally, cyanobacteria IOM contain phycobiliproteins that are highly fluorescent in the

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High Wavelength Range (Figure 1). All three species exhibited fluorescence emission at

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wavelengths characteristic of phycocyanin (PC) in the monomer or trimer forms (Bryant, 1994;

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Debreczeny et al., 1993). For OSC, a benthic species, its greatest fluorescence intensity occurred

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at lower wavelengths associated with phycoerythrin (Bryant, 1982). Phycoerythrin (PE) is

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beneficial to benthic cyanobacteria, because the longer wavelengths of sunlight are attenuated

259

with increasing depth resulting in predominately blue-green light exposure (MacColl, 1998).

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Phycoerythrin increases the ability of benthic cyanobacteria to utilize these wavelengths. Not all

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species produce phycoerythrin but some strains of both OSC and LYG are known producers

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(Bryant, 1982; Suda et al., 2002). For the purpose of identifying a universal cyanobacteria IOM

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surrogate, phycocyanin is the only viable fluorescent surrogate in the High Wavelength Range,

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because all naturally occurring cyanobacteria species produce phycocyanin whereas not all

265

produce phycoerythrin (Bryant, 1994; 1982).

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3.1.2

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intensities demonstrate compositional differences between species. Table 1 tabulates the specific

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fluorescence intensities for Peak P, FDOM, PC and PE. Preservation with sodium thiosulfate led

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to systematic changes in FDOM intensity; only unpreserved data are reported for this measure

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Quantitative Fluorescence Comparisons While each cyanobacteria exhibited similar spectral features, specific fluorescence

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(see SI-Text 2). Across all fluorescence regions, MA is more fluorescent than OSC and LYG by

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emitting greater fluorescence intensities per unit carbon. MA IOM also has a higher SUVA (1.59

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L mgC-1 m-1) compared to OSC and LYG (0.66 and 0.26 L mgC-1 m-1, respectively) and is

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enriched in optically active moieties (Wert et al., 2014). These cultures were harvested in the late

275

exponential phase where metabolite concentrations have been shown to be the greatest (Pietsch

276

et al., 2002; Watanabe et al., 1989), but it is unknown how IOM fluorescence properties change

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across growth phases. From a practical monitoring standpoint, measuring fluorescence intensities

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to detect IOM release would be more sensitive towards more optically active species (e.g., MA)

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where the fluorescence response per unit mass of IOM released is greater.

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Comparing the cyanobacteria fluorescence to CRW-DOM demonstrates some of the

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advantages and disadvantages related to monitoring regional fluorescence. The cyanobacteria-

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derived fluorescence overlaps with regions of strong fluorescence in CRW-DOM and other

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natural waters in the Low Wavelength Range. For all species, Peak P specific intensity is greater

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in cyanobacteria compared to CRW-DOM, suggesting that this region would have greater

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fluorescence sensitivity for capturing increases in IOM in the dissolved phase. The specific

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FDOM intensity for CRW falls in between LYG and OSC and is about half as high as MA. This

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result suggests that FDOM may be more responsive to detect highly fluorescent IOM from

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species such as MA and more difficult to detect IOM contributions over background DOM from

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less fluorescent species like LYG. Finally, CRW-DOM is weakly fluorescent in the High

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Wavelength Region demonstrating that phycocyanin fluorescence intensity could be a universal

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fluorescence indicator across all cyanobacteria species in ultrapure water.

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FI values for all three species are much greater than both CRW-DOM and the range of

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values typically reported in literature, including wastewater effluent organic matter (Dong and

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Rosario-Ortiz, 2012; McKnight et al., 2001). All IOM FI values are greater than 2.2 (Table 1).

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Values above 2 are high compared to microbially derived fulvic acids and have been attributed to

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nonhumic organic matter released directly from algae (McKnight et al., 2001). The large

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difference in FI between cyanobacteria IOM and CRW-DOM suggests that FI could be a viable

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surrogate to monitor IOM contributions by measuring compositional changes in the dissolved

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phase. It is important to note that during a cyanobacteria bloom, the background DOM will also

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contain EOM, which can increase FI and make the differences between IOM and background

301

DOM smaller.

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3.2

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fluorescence behavior in a heterogeneous background matrix of DOM. An ideal fluorescence

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surrogate for IOM would exhibit minimal interactions with background DOM. This section

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evaluates if IOM fluorescence indicators behave independently of CRW-DOM or if they interact

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to either enhance or quench IOM fluorescence. Isolated IOM was spiked into CRW at increasing

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concentrations to determine the effect of a background matrix on the IOM fluorescence signature.

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3.2.1

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initial addition of IOM leads to the greatest increase in FI with subsequent additions leading to

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incrementally smaller FI increases. Past work has demonstrated that FI does not follow

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conservative property balance principles, because there are competing effects of peak emission

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wavelength and spectrum curvature (Korak et al., 2014). In this study, the peak emission

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wavelength at excitation 370 nm for OSC and LYG was blue-shifted compared to CRW, but MA

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was not. As OSC and LYG IOM are spiked into CRW, the observed increase in FI was due to

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primarily a shift in peak emission wavelength. For MA, there was no systematic shift in peak

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Low Wavelength Range (Ex 250-450 nm, Em 300-560 nm) FI increases with increasing IOM concentration but the trend is not linear (Figure 2a). An

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emission, and the observed increase was due to a change in local curvature. For monitoring

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applications, an increase in FI may be an indicator for released cyanobacteria organic matter in a

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water treatment plant, but the magnitude of the increase is not directly proportional to the mass

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released.

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Figure 2b depicts the Peak P fluorescence response to increasing IOM concentrations in

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CRW. Figure 2b shows that peak intensity increases proportionally to the IOM spike

324

concentration. OSC and LYG both had similar specific Peak P intensities and showed similar

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Peak P increases when spiked in CRW. MA IOM had a higher specific Peak P intensity, and as a

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result it showed a larger increase in Peak P fluorescence per unit increase in IOM DOC. By

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comparing the observed increases for IOM spiked into CRW to the specific peak intensities of

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the isolated IOM, an assessment of the interactions can be made. If there are no interactions, then

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the measured intensities should increase proportional to the specific fluorescence intensities in

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Table 1. Using OSC as an example, adding 1 mgC/L of OSC IOM to CRW-DOM should increase

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the fluorescence intensity by 0.29 RU based on the specific Peak P intensity in Table 1. The

332

measured increase was only 0.17 RU, indicating that the mixture fluoresces with lower

333

efficiency and is apparently quenched. Regressions were fit to each data series in Figure 2b, and

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the slopes represent the average increase in Peak P intensity per unit DOC added. Comparing

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these slopes to the specific peak intensities in Table 1, the average observed increase of the

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mixture was 20-35% less for all three species than predicted by the specific intensity of the

337

isolated IOM. When the IOM contribution in the dissolved phase increases, the fluorescence

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signal measured is not simply a superposition of IOM and CRW-DOM intensities. Therefore, it

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would be difficult to quantify the amount of IOM released in a treatment plant from changes in

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intensity alone.

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Lastly, the FDOM fluorescence intensity also increases with increasing IOM

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concentration (Figure 2c). Similar to Peak P intensity, there is evidence of non-ideal interactions

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such that the measured fluorescence of the mixture is not a superposition of signals from CRW-

344

DOM and IOM. The observed increase in intensity in Figure 2c is 20% less for OSC than the

345

amount predicted by the specific peak intensities of the isolated IOM (Table 1). These results

346

suggest that the interactions between the IOM and CRW-DOM lead to a quenching of intensity

347

in this region. For LYG, the observed fluorescence increase when spiked into CRW was about

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40% greater than predicted suggesting a fluorescence enhancement. Intensity increases were

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greatest for MA IOM and were about 80% greater than predicted by specific fluorescence

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intensities of the isolated IOM.

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These results highlight the importance of selecting appropriate metrics for monitoring

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dynamic systems or compositional changes in DOM. FI has potential to be an easily monitored

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fluorescence indicator, because it is operationally easy to measure and does not require inner

354

filter corrections for most applications (Korak et al., 2014; McKnight et al., 2001). Since it is

355

largely concentration-independent, it may be more robust towards fluctuations in background

356

DOM quantity or quality and be less dependent on differences in specific fluorescence intensities.

357

The downside is that an FI increase is not directly proportional to the quantity of IOM introduced.

358

Single point fluorescence intensities can provide better direct correlations between fluorescence

359

response and changes in IOM mass. Disadvantages are that any quantitative calibrations must

360

include interaction effects and cannot be based solely on isolated standards. Quantitative

361

relationships that attempt to relate changes in fluorescence intensity to a quantity of IOM

362

released into the bulk phase are further complicated by mixed species cultures, as found in the

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environment. For this reason, fluorescence is better suited as a qualitative indicator of IOM

364

release in a water treatment plant to prompt further action.

365 366

3.2.2

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strongly affected by interactions between CRW-DOM and IOM in both intensity and spectra.

368

The interactions were assessed by spiking both IOM and pigment standards into CRW. Adding

369

either phycocyanin or phycoerythrin standards to CRW-DOM found no indication of interaction

370

effects. There was no change in peak location and the fluorescence intensity increase was

371

representative of the standard fluorescence in phosphate buffer (Figure 3, Figures S-2 and S-3).

372

When IOM is spiked into CRW, however, the pigment fluorescence intensity decreased by more

373

than an order of magnitude across the entire region (Figure 4). In addition to decreased intensity,

374

new fluorescence behavior was observed from the interacting species (Figure 4c, 4i). For OSC,

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there was no indication of phycobiliproteins when 1 mgC/L is added to CRW (Figure 4f), but

376

fluorescence signatures similar to MA and LYG appear at higher concentrations (Figure S-6)

377

The emission at 650 nm showed two distinct excitation maxima around 580 nm and 640 nm.

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There was a new fluorescence peak that appears at 580 nm excitation and 600 nm emission that

379

was not present in the isolated IOM. This behavior was observed for all three species. It is likely

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due to the interactions of phycocyanin rather than the other phycobiliproteins, because

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phycoerythrin is not common to all species.

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High Wavelength Range (Ex 450-700 nm, Em 550-700 nm) Analysis of the phycobiliprotein region indicates that the fluorescence in this region is

The specific mechanism causing the fluorescence quenching and wavelength shifts

383

observed between CRW-DOM and IOM are unknown. One study examined the interactions

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between phycobiliproteins and both glutardialdyhyde and benzoquinone and found that two

385

different mechanisms can lead to a decrease in phycobiliprotein fluorescence intensity (Köst et

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al., 1981). Phycobiliproteins fluoresce, because the tertiary polypeptide backbone provides

387

molecular rigidity and prevents non-radiative decay (Köst et al., 1981). For glutardialdehyde, it

388

was suspected that the intramolecular interactions of the tertiary structure are interrupted, which

389

increases molecular flexibility and decreases fluorescence. Benzoquinone, on the other hand, is

390

thought to quench fluorescence through energy transfer interactions (Köst et al., 1981). Both

391

quenching mechanisms have kinetic half lives ranging from less than 10 minutes to several hours

392

(Köst et al., 1981). In the current study, samples from both the IOM and phycobiliproteins

393

standard experiments were subjected to the similar experimental conditions, storage and holding

394

times, which suggests that there is something compositionally different about the IOM samples

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affecting pigment fluorescence. The purified phycobiliproteins appear to behave differently and

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may be due to the absence of other components, such as polypeptide linkers (MacColl, 1998).

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In any case, the interactions between the phycobiliproteins and CRW-DOM have some

398

practical implications for using phycobiliproteins as an IOM fluorescence surrogate. Calibration

399

to a standard solution would greatly underestimate the presence of IOM in a background DOM

400

matrix if interaction effects were not taken into account. There may also be kinetic aspects of the

401

quenching interactions to be considered. This study showed that sample hold times as short as 3

402

days exhibit severe quenching interactions. Monitoring for a decrease of in vivo fluorescence as

403

an indicator of IOM release in a water treatment plant may be a better strategy for commercially

404

available phycocyanin probes. A decrease in in vivo phycocyanin fluorescence may be inferred

405

as IOM release rather than directly measuring phycocyanin in the dissolved phase.

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3.3

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chloramine and chlorine dioxide to determine how their fluorescence signatures change in an

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IOM Oxidation Studies The three IOM isolates in phosphate buffer were exposed to ozone, free chlorine,

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oxidizing environment. Oxidation of IOM isolates is necessary to interpret the fluorescence

410

signal response for future oxidation studies of intact cells and to identify persistent fluorescence

411

characteristics. An ideal fluorescence surrogate would exhibit no change in its fluorescence

412

signature to act as a conservative indicator for the release of other metabolites of interest.

413

Oxidant dose is presented as the ratio of oxidant concentration to DOC concentration to

414

normalize for differences in IOM concentration.

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Analyzing changes in FI and FDOM in tandem reveals how both IOM quantity and

416

composition change during oxidation within the same EEM region. Figure 5 shows that ozone

417

oxidation led to the greatest loss of FDOM intensity and a simultaneous change in composition

418

as indicated by a decrease in FI. Chlorine dioxide oxidation decreased FDOM intensity but to a

419

lesser extent compared to ozone. The maximum decrease for chlorine dioxide was about 40% for

420

OSC whereas the decrease for MA and LYG was about 10%. Free chlorine oxidation did not

421

cause any appreciable decrease in FDOM intensity for MA but caused about a 40% decrease at

422

the highest dose ratio for LYG and OSC. FI decreased at the lowest chlorine dioxide and free

423

chlorine doses applied with little change at higher doses, indicating a compositional change due

424

to oxidation. Chloramine oxidation had little effect on FDOM intensity for OSC and LYG but

425

caused an enhancement for MA. This trend suggests that there are compositional differences

426

between the species that lead to different oxidation responses despite similarities in their

427

fluorescence signatures. There was little change in FI due to chloramine oxidation indicating that

428

this oxidant is too weak to cause significant compositional changes to the IOM.

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Fluorophores in the Peak P region are more susceptible to oxidation compared to the

430

region where FDOM and FI are measured. Peak P intensities decreased by upwards of 90% with

431

exposure to ozone, chlorine dioxide and free chlorine (Figure 5). Chloramines had little effect on

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Peak P intensity for OSC and caused about a 25% decrease in intensity for MA and LYG.

433

Fluorescence loss differed by species. In general, intensity loss was the greatest for MA and

434

lowest for OSC, indicating reactivity differences in IOM.

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Lastly, phycocyanin fluorescence is rapidly lost with exposure to all four oxidants

436

(Figure 5). Ozone, chlorine dioxide and free chlorine led to more than a 90% decrease in

437

intensity at the lowest oxidant doses. Even though chloramine oxidation had little effect on Low

438

Wavelength Region fluorescence indicators, phycocyanin fluorescence decreased 40-85% at the

439

lowest dose ratios. Phycoerythrin fluorescence in OSC also decreased with exposure to all four

440

oxidants (Figure S-7). The fluorescence responses of phycobiliproteins in IOM agree with

441

oxidation studies of phycocyanin and phycoerythrin standards in phosphate buffer (Figure S-8

442

and Table S-1).

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Comparing the different fluorescence responses demonstrates that the utility of

444

fluorescence as a monitoring tool is metric- and oxidant-specific. For the same oxidant, there was

445

a greater loss of intensity for Peak P and phycocyanin compared to FDOM, suggesting that the

446

fluorescing moieties in the FDOM region are less reactive. Therefore, it would be advantageous

447

to monitor fluorescence FDOM intensities rather than Peak P or phycocyanin wavelengths,

448

because IOM released in a water treatment plant would be more persistent. The use of

449

fluorescence as a surrogate for IOM is more promising for weaker oxidants (free chlorine and

450

chloramine) compared to stronger oxidants (ozone and chlorine dioxide), where the change in

451

IOM fluorescence due to oxidation is smaller. IOM released during ozonation will react with

452

continued exposure, lose its fluorescence signature, and become difficult to detect by intensity

453

changes alone. It is important to note that this study used preformed chloramines, but in

454

applications where chloramines are formed by sequential addition free chlorine and ammonia,

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observed fluorescence responses may be closer to that of free chlorine. FI may be more effective

456

than measuring regional intensity, because the lowest FI values observed are still higher than that

457

of bulk CRW and most other surface waters. The differences in FI between all three IOM and

458

CRW suggest that FI would be a better method than FDOM intensities for detecting IOM release

459

by tracking compositional changes.

460 461

4

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Conclusions

In the Low Wavelength range, MA, OSC and LYG all exhibited strong protein-like fluorescence, common emission at 460 nm and high FI values.

•

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In the High Wavelength Region, planktonic cyanobacteria species (MA and LYG) were

464

dominated by phycocyanin fluorescence whereas the benthic cyanobacteria specie (OSC)

465

had strong phycoerythrin fluorescence. •

467 468

Across all regions, MA was more optically active and exhibited a greater fluorescence response per unit carbon.

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Matrix effects are important when IOM was mixed with CRW-DOM. Fluorescence intensities of the mixture were not a linear superposition of the starting materials due to

470

interactions. Phycobiliprotein fluorescence intensity was severely quenched by

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interactions with CRW-DOM. •

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FDOM and FI fluorescence signatures were most persistent in oxidizing environments compared to protein-like fluorescence and phycobiliprotein fluorescence.

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The development of fluorescent probes to detect IOM release in a water treatment plant

should focus on the FDOM and FI region where an IOM signal is more persistent.

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476

•

This work would be directly applicable to the development of fluorescence monitoring strategies in a drinking water treatment plant where measurements could be taken directly

478

upstream and downstream of oxidation processes.

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Supporting Information

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Supporting information contains additional text, 1 table and 8 figures.

481

Acknowledgements

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The authors acknowledge the Water Research Foundation (project number 4406) for their

483

financial support of this project. J. Korak acknowledges the National Science Foundation for

484

their support through the Graduate Research Fellowship Program (DGE 1144083). The authors

485

thank Mei Mei Dong with the SNWA Water Quality Research and Development team for

486

preparing the intracellular organic matter standards. The authors also thank Kaelin Cawley and

487

Amanda Hohner for their insightful feedback to the manuscript.

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and Cylindrospermopsin. Marine Drugs 8, 1650–1680. doi:10.3390/md8051650 Pietsch, J., Bornmann, K., Schmidt, W., 2002. Relevance of Intra- and Extracellular Cyanotoxins for Drinking Water Treatment. Acta hydrochim. hydrobiol. 30, 7–15. Qu, F., Liang, H., Wang, Z., Wang, H., Yu, H., 2012. Ultrafiltration membrane fouling by extracellular organic matters (EOM) of Microcystis aeruginosa in stationary phase: Influences of interfacial characteristics of foulants and fouling mechanisms. Water Research 46, 1490–1500. Reynolds, D.M., 2003. Rapid and direct determination of tryptophan in water using synchronous fluorescence spectroscopy. Water Research 37, 3055–3060. Saraceno, J.F., Pellerin, B.A., Downing, B.D., Boss, E., Bachand, P.M., Bergamaschi, B.A., 2009. High-frequency in situ optical measurements during a storm event: Assessing relationships between dissolved organic matter, sediment concentrations, and hydrologic processes. J Geophys Res-Biogeo 114, G00F09. Sinclair, J.L., Hall, S., 2008. Chapter 3: Occurrence of Cyanobacterial Harmful Algal Blooms Workgroup Report, in: Hudnell, H.K. (Ed.), Cyanobacterial Harmful Algal Blooms: State of the Science and Research Needs. Springer New York, New York, pp. 45–103. Smith, J.L., Boyer, G.L., Zimba, P.V., 2008. A review of cyanobacterial odorous and bioactive metabolites: Impacts and management alternatives in aquaculture. Aquaculture 280, 5–20. Stewart, D., Farmer, F., 1984. Extraction, identification, and quantitation of phycobiliprotein pigments from phototrophic plankton. Limonology and Oceanography 29, 392–397. Suda, S., Watanabe, M.M., Otsuka, S., Mahakahant, A., Yongmanitchai, W., Nopartnaraporn, N., Liu, Y.D., Day, J.G., 2002. Taxonomic revision of water-bloom-forming species of oscillatorioid cyanobacteria. International Journal of Systematic and Evolutionary Microbiology 52, 1577–1595. Watanabe, M.F., Harada, K.-I., Matsuura, K., Watanabe, M., Suzuki, M., 1989. Heptapeptide toxin production during the batch culture of two Microcystis species (Cyanobacteria). J Appl Phycol 1, 161–165. Watanabe, M.F., Oishi, S., 1985. Effects of Environmental Factors on Toxicity of a Cyanobacterium (Microcystis aeruginosa) under Culture Conditions. Applied and Environmental Microbiology. Watras, C.J., Baker, A.L., 1988. Detection of planktonic cyanobacteria by tandem in vivo fluorometry. Hydrobiologia 169, 77–84. Watson, S.B., 2003. Cyanobacterial and eukaryotic algal odour compounds: signals or byproducts? A review of their biological activity. Phycologia 42, 332–350. Watson, S.B., 2004. Aquatic Taste and Odor: a Primary Signal of Drinking-Water Integrity. Journal of Toxicology and Environmental Health, Part A: Current Issues 67, 1779–1795. Wert, E.C., Dong, M.M., Rosario-Ortiz, F.L., 2013. Using digital flow cytometry to assess the degradation of three cyanobacteria species after oxidation processes. Water Research 47, 3752–3761. Wert, E.C., Korak, J.A., Trenholm, R.A., Rosario-Ortiz, F.L., 2014. Effect of oxidant exposure on the release of intracellular microcystin, MIB, and geosmin from three cyanobacteria species. Water Research 251–259. Wert, E.C., Rosario-Ortiz, F.L., 2013. Intracellular Organic Matter from Cyanobacteria as a Precursor for Carbonaceous and Nitrogenous Disinfection Byproducts. Environ Sci Technol 6332–6340. Wert, E.C., Rosario-Ortiz, F.L., Snyder, S.A., 2009. Effect of ozone exposure on the oxidation of

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trace organic contaminants in wastewater. Water Research 43, 1005–1014. WHO, 2003. Cyanobacterial toxins: Microcystin-LR in drinking water. Background document for preparation of WHO guidelines for drinking-water quality (No. WHO/SDS/WSH/03.04/57). World Health Organization, Geneva. Wolfbeis, O.S., 1985. Fluorescence of Organic Natural Products, in: Schulman, S.G. (Ed.), Molecular Luminescence Spectroscopy, Part 1. John Wiley & Sons, New York, pp. 167–370. Zamyadi, A., Ho, L., Newcombe, G., Bustamante, H., Prévost, M., 2012a. Fate of toxic cyanobacterial cells and disinfection by-products formation after chlorination. Water Research 46, 1524–1535. Zamyadi, A., MacLeod, S.L., Fan, Y., McQuaid, N., Dorner, S., Sauvé, S., Prévost, M., 2012b. Toxic cyanobacterial breakthrough and accumulation in a drinking water plant: a monitoring and treatment challenge. Water Research 46, 1511–1523. Zamyadi, A., McQuaid, N., Prévost, M., Dorner, S., 2012c. Monitoring of potentially toxic cyanobacteria using an online multi-probe in drinking water sources. J. Environ. Monit. 14, 579–588. Ziegmann, M., Abert, M., Müller, M., Frimmel, F.H., 2010. Use of fluorescence fingerprints for the estimation of bloom formation and toxin production of Microcystis aeruginosa. Water Research 44, 195–204.

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FI

FDOM Sp. Intensity (RU L mgC-1) 0.081 0.052 0.021 0.039 ± 0.002

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(--) 2.32 ± 0.02 2.65 ± 0.06 2.39 ± 0.05 1.50 ± 0.01

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MA IOM OSC IOM LYG IOM CRW DOM

(mgC L-1) 2.0 ± 0.15 1.0 ± 0.02 1.07 ± 0.09 2.44 ± 0.05

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Peak P (Ex, Em) Sp. Intensity (nm) (RU L mgC-1) 280, 330 0.49 ± 0.04 270, 302 0.29 ± 0.02 270, 301 0.27 ± 0.02 260, 350 0.16 ± 0.01

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Table 1. Summary of fluorescence metrics for isolated cyanobacteria IOM in phosphate buffer and CRW. The average and standard deviations are listed for the specific Peak P intensities and FI (n=4). Peak positions are rounded to the nearest integer. IOM specific FDOM intensities are for the unpreserved control sample. PC Sp. Intensity (RU L mgC-1) 1.29 ± 0.07 0.44 ± 0.11 0.57 ± 0.11 --

PE Sp. Intensity (RU L mgC-1) -1.35± 0.30 ---

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Figure 1. EEMs of three cyanobacteria IOM and CRW. Intensities are normalized to the DOC concentration and are presented in units of RU L mgC-1. Note the wavelengths and intensity scales are different between EEMs. The top row is the Low Wavelength Range and the bottom row in the High Wavelength Region. Each column represents a different IOM species or CRW. The different fluorescence metrics are annotated: Peak P (white box), FDOM intensity (unfilled square marker), FI (points connected by line), phycocyanin (PC marker) and phycoerythrin (PE marker).

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Figure 2. Response of Low Wavelength Range fluorescence indicators to IOM added into CRW in increasing concentrations: a) Fluorescence index, b) Peak P intensity and c) FDOM fluorescence intensity (Ex 370 nm, Em 460 nm)

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Figure 3. EEM series of phycobiliprotein standards added to CRW a-c) phycocyanin (PC) series and d-f) phycoerythrin (PE) series. Note- the CRW intensity is two orders of magnitude smaller than PC and PE.

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Figure 4. EEM series of IOM added to CRW a-c) MA and d-f) OSC and g-i) LYG. Note- the CRW intensity is 1-2 orders of magnitude smaller than PC and PE.

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Ozone 2.80

Free Chlorine

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Figure 5. Response of fluorescence indicators to oxidation for IOM and CRW. Each row follows a different indicator: FI and the relative changes in FDOM, Peak P and phycocyanin (PC) intensities. Each column is a different oxidant. One free chlorine dose ratio (1) was run as experimental duplicates and the error bars represent the standard deviation, but most error bars are smaller than the marker size.

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-Differences in cyanobacteria IOM fluorescence signature are evaluated -Fluorescence Index (FI) and FDOM signals most persistent -Cyanobacteria pigment fluorescence is quenched by DOM

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-Cyanobacteria pigment fluorescence is readily oxidized

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Supplemental Information Evaluating fluorescence spectroscopy as a tool to characterize

oxidation in natural water

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cyanobacteria intracellular organic matter upon simulated release and

Julie A. Korak1, Eric C. Wert2 and Fernando L. Rosario-Ortiz1*

Department of Civil, Environmental and Architectural Engineering, 428 UCB,

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University of Colorado, Boulder, CO 80309, USA Southern Nevada Water Authority (SNWA), P.O. Box 99954, Las Vegas, NV 89193-

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9954 USA

*Corresponding author: [email protected]

Cyanobacteria Pigments (Phycobiliproteins) Background

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Phycobilisomes are light harvesting complexes attached to the thylakoid membrane in cyanobacteria that expand the range of wavelengths the organism can utilize for photosynthesis.

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They absorb light in the visible range predominately from 550 to 650 nm, where chlorophyll a

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cannot absorb light (Bryant, 1994). Phycobilisomes in most cyanobacteria are hemidiscoidal with a tricylindrical core (MacColl, 1998). The tricylindrical core closest to the thylakoid membrane contains the phycobiliprotein allophycocyanin. Six rods extending from the core contain other phycobiliproteins such as phycocyanin, phycoerythrin or phycoerythrocyanin. Phycobiliproteins are arranged such that the ones that absorb at the lowest wavelength (ie phycoerythrin) are located at the ends of the rods whereas allophycocyanin, which absorbs at the highest wavelength, is located in the core next to the membrane. This arrangement allows for 1

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efficient energy transfer of absorbed light through the phycobilisome to allophycocyanin, which is then transferred to chlorophyll a. Energy transfer is a very efficient, nonradiative process that occurs mainly due to Förster resonance energy transfer and some exciton state coupling

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(MacColl, 1998).

Bilins are open-chain tetrapyrroles and are the active chromo- and fluorophores. There are four bilins (phycocyanobilin, phycoerythrobilin, phycocourobilin and phycoviolobilin) with

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varying degrees of conjugated double bonds that determine the range in which they absorb light. Phycourobilin has 5 conjugated double bonds and absorbs near 495 nm whereas phycocyanobilin

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has 8 conjugated double bonds and absorbs at wavelengths greater than 600 nm (MacColl, 1998). Bilins are covalently bonded to a polypeptide backbone between 15,000 and 20,000 Da at cysteine residues to form the phycobiliproteins (MacColl, 1998). Each phycobiliprotein monomer consists of two subunits (denoted α and β) and is defined by which bilins are attached

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and the location at which they are attached. For example, phycocyanin has one phycocyanobilin attached to the α subunit at the 84th residue (α84) and two phycocyanobilins attached to the β subunit at residues 84 and 155 (denoted β84 and β155, respectively). Allophycocyanin only has

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one phycocyanobilin attached to each subunit (α84 and β84) and is a distinctly different phycobiliproteins (Debreczeny et al., 1993). The monomers have a distinct fluorescence

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signature and energy transfer between the bilins. Three monomers can aggregate to form a discshaped trimer, which can then stack together to form hexamers (Bryant, 1994; MacColl, 1998). The formation of larger aggregates changes the fluorescence signature. In addition to the phycobiliproteins, polypeptide linkers that connect the substructure of the phycobilisome also alter the optical properties (MacColl, 1998).

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Effect of preservation with sodium thiosulfate Preservation with sodium thiosulfate had no systematic effect on Peak P or

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phycobiliproteins fluorescence, so specific intensities presented in Table 1 include all control samples, with and without preservative. Adding preservative decreased the FDOM fluorescence specific intensity for MA and LYG by 0.02 and 0.14 RU L mgC-1, respectively, while slightly

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increasing the specific intensity for LYG by 0.005 RU L mgC-1. The reproducibility among the three preserved FDOM specific peak intensities had CV values between 6 and 10% (including

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DOC error) and suggest that the decrease is significant. Sample preservation did not lead to any systematic change in FI. Since FI did not change, it suggests that the fluorescence is quenched

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uniformly across those emission wavelengths.

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IOM and CRW Excitation-Emission Matrices (EEMs) MA IOM 2 mgc/L

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Figure S-1. Excitation-emission matrices of a) Colorado River Water, b) Oscillatoria sp., c) Microcystis aeruginosa and d) Lyngbya sp. The Low Wavelength and High Wavelength Regions are outlined in white boxes.

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IOM Phycobiliprotein Fluorescence Interactions with CRW-DOM CRW

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Figure S-2. EEMs of CRW, phycocyanin (PC) standards in phosphate buffer and phycocyanin spiked into CRW in increasing concentrations.

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Figure S-3. EEMs of CRW, phycoerythrin (PE) standards in phosphate buffer and phycoerythrin spiked into CRW in increasing concentrations.

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MA IOM 2.5 mgC/L

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Figure S-4. EEMs for a) Microcystis IOM at 2.5 mgc/L in LGW, b) Colorado River Water, and c-e) Microcystis IOM spiked into CRW at increasing concentrations.

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IOM Oxidation Studies OSC 1.0

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Figure S-8. Fraction of fluorescence intensity remaining of phycocyanin standard in phosphate buffer as a function of oxidant to DOC ratio for ozone, chlorine dioxide, free chlorine and chloramine.

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Table S-1. Phycoerythrin standard in phosphate buffer intensity after oxidation by ozone, free chlorine, chlorine dioxide and chloramine represented by the measured intensity and intensity relative to the control. Max Intensity Remaining (RU) Fluorescence (%) 9.05 100% 0.00 0.04% 2.14 24% 0.22 2.4% 8.97 99%

References

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Oxidant:DOC Ratio 0 1 1 1 1

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Oxidant Control Ozone Free Chlorine Chlorine Dioxide Chloramine

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Bryant, D.A., 1994. Molecular Biology of Cyanobacteria. Kluwer Academic Publishers. Debreczeny, M.P., Sauer, K., Zhou, J., Bryant, D.A., 1993. Monomeric C-phycocyanin at room temperature and 77 K: resolution of the absorption and fluorescence spectra of the individual chromophores and the energy-transfer rate constants. Journal of Physical Chemistry 97, 9852–9862. MacColl, R., 1998. Cyanobacterial phycobilisomes. J Struct Biol 124, 311–334.

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