Cbc

BD1

h

•

•

•

•

•

a

ARR1AA

KUFPDD

0h

Journal of Hazardous Materials 264 (2014) 411– 419

Contents lists available at ScienceDirect

Journal of Hazardous Materials

jou rn al hom epage: www.elsev ier .com/ locate / jhazmat

hanges in dissolved organic matter fluorescence and disinfectionyproduct formation from UV and subsequenthlorination/chloramination

onnie A. Lyon1, Rose M. Cory2, Howard S. Weinberg ∗

epartment of Environmental Sciences and Engineering, Gillings School of Global Public Health, University of North Carolina at Chapel Hill,46 Rosenau Hall, Chapel Hill, NC 27599, United States

i g h l i g h t s

DBP formation from UV-chlorine/chloramine drinking watertreatment was measured.The effect of UV on DBP precursorswas evaluated by fluorescence andPARAFAC.UV alone decreasedprotein/tryptophan- and humic-likefluorescence.Loss of two components correlatedwith cyanogen chloride formation(R2 = 0.79–0.91).Loss of the components also corre-lated with chloral hydrate formation(R2 = 0.95–1.000).

g r a p h i c a l a b s t r a c t

r t i c l e i n f o

rticle history:eceived 4 July 2013eceived in revised form3 September 2013ccepted 28 October 2013vailable online 4 November 2013

eywords:ltraviolet treatment

a b s t r a c t

Ultraviolet (UV) irradiation is being increasingly used to help drinking water utilities meet finished waterquality regulations, but its influence on disinfection byproduct (DBP) precursors and DBP formation is notcompletely understood. This study investigated the effect of medium pressure (MP) UV combined withchlorination/chloramination on the fluorescent fraction of dissolved organic matter (DOM) isolated froma United States surface water with median total organic carbon content. Parallel factor analysis was usedto understand how UV may alter the capacity of DOM to form DBPs of potential human health concern.The production of chloral hydrate and cyanogen chloride from MP UV followed by chlorine or chloramine,respectively, correlated with a decrease in fluorescence intensity of a protein/tryptophan-like compo-

2 2

luorescence spectroscopyarallel factor analysisisinfection byproductsissolved organic matter characterization

nent (R = 0.79–0.99) and a humic-like component (R = 0.91–1.00). This suggests that the UV-inducedprecursors to these compounds originated from DOM with similar characteristics to these components.The fluorescent DOM components identified in this study are similar to reoccurring components thathave been previously identified in a range of raw and treated waters, and this work demonstrates the

value of using fluorescence anDBP formation under a range o

∗ Corresponding author. Tel.: +1 919 966 3859.E-mail address: howard weinberg@unc.edu (H.S. Weinberg).

1 Present address: Advanced Water Management Centre, University of Queensland, Lev2 Present address: Department of Earth and Environmental Sciences, University of Mic

304-3894/$ – see front matter © 2013 Elsevier B.V. All rights reserved.ttp://dx.doi.org/10.1016/j.jhazmat.2013.10.065

alysis of DOM to understand the relationships between DOM source andf treatment conditions.

© 2013 Elsevier B.V. All rights reserved.

el 4, Gehrmann Laboratories Building (60), Brisbane, Queensland 4072, Australia.higan, Ann Arbor, MI 48109, United States.

4 rdous

1

biittiarmutcn

t(oegndrttdticfacottuw

uttbripncao(mto

D[ntrnittD

12 B.A. Lyon et al. / Journal of Haza

. Introduction

Dissolved organic matter (DOM), formed from plant and micro-ial decay products, is present in all natural waters and plays an

mportant role in many natural and engineered processes [1–5]. Its the main precursor to disinfection byproducts (DBPs) of poten-ial human health concern that are produced during drinking waterreatment [6,7]. DBPs are produced when DOM, other chemicalsn water which can be of natural origin (e.g., bromide, iodide), ornthropogenic pollutants react with a disinfectant such as chlo-ine. Trihalomethanes (THMs) and haloacetic acids (HAAs) are twoajor DBP classes formed during chlorination, and a subset are reg-

lated in the United States [8]. As anthropogenic activity continueso stress source water quality and more stringent DBP regulationsome into effect, utilities are exploring treatment processes alter-ative to those conventionally practiced.

An occurrence study of drinking water treatment plants acrosshe United States found that the use of alternative disinfectantschloramine, chlorine dioxide, ozone) decreased the formationf regulated THMs and HAAs, while in many cases there wasnhanced formation of other DBPs that are thought to be moreeno- and cytotoxic than those currently regulated [9]. One alter-ative process stream uses ultraviolet (UV) irradiation as a primaryisinfectant in treatment plants employing chlorination or chlo-amination for secondary disinfection. The majority of past workhat has evaluated UV-chlorine/chloramine treatment found lit-le effect on regulated DBPs when UV was applied at disinfectionoses (40–186 mJ/cm2) [10–12], but less research has focused onhe formation of unregulated DBPs thought to be more toxicolog-cally potent [13]. In one study that did look at additional DBPlasses, increased trichloronitromethane formation was observedollowing medium pressure (MP) UV irradiation (40–140 mJ/cm2)nd post-chlorination compared to chlorination alone in watersontaining nitrate (1–10 mg N/L) [14]. The relative importancef source water characteristics (e.g., nitrate, DOM concentra-ion/quality) leading to increased trichloronitromethane after UVreatment is not well understood and requires further study to eval-ate the implications of integrating UV into the overall drinkingater production process.

Average characteristics of DOM are often studied to betternderstand DBP formation because DOM is a heterogeneous mix-ure comprised of thousands of compounds that vary spatially andemporally in source waters [15]. DOM character is increasinglyeing studied using fluorescence spectroscopy because with oneelatively quick and simple measurement, this technique providesnformation on at least three types of carbon within the DOMool. For example, three different fluorescent DOM (FDOM) sig-als have been correlated with low or high aromaticity carbon thatorresponds to microbial- vs. terrestrial-precursor material, aminocid-like compounds associated with free or combined amino acids,r polyphenols [16,17]. The application of parallel factor analysisPARAFAC), a statistical modeling technique, to excitation-emission

atrix (EEM) fluorescence data allows for quantitative identifica-ion of mathematically and chemically independent components,r types of carbon, within the FDOM [18].

Two recent studies used fluorescence and PARAFAC to trackOM fate through drinking and recycled water treatment plants

19,20]. Baghoth and colleagues observed changes in compo-ent fluorescence across drinking water treatment processeshat were consistent with previous work that used more labo-ious techniques (e.g., high performance liquid chromatography,uclear magnetic resonance spectroscopy) to follow DOM chem-

stry and its removal during treatment [19]. These results suggesthat while FDOM may only comprise a small fraction of theotal DOM, the reactivity of FDOM is representative of the bulkOM.

Materials 264 (2014) 411– 419

There is potential to use fluorescence/PARAFAC to understandlinkages between DOM and DBP formation, particularly for eval-uating the effects of different treatment processes and theiroptimization to minimize DBP formation. Beggs and Summersfound that an aromatic humic-like precursor (associated with morehydrophobic DOM) correlated with regulated THM and HAA forma-tion following chlorination (R2 = 0.89 and 0.69, respectively), whilemore hydrophilic/protein-like components showed little correla-tion (R2 ≤ 0.40) [21]. The finding that different sources and typesof DOM are associated with DBP formation is not new, but theresults of this study and others [22–25] suggest that fluorescencespectroscopy could be used to monitor or predict DBP formationpotential across treatment plants in real-time. While this approachhas promise, little work has investigated the relationship betweenDOM fluorescence and unregulated byproducts that are potentiallymore geno- and cytotoxic than the regulated THMs and HAAs [13].Similarly, few studies have explored changes in DBP precursorsfrom engineered UV processes, especially with methods that areable to detect subtle changes in DOM character.

The objectives of this study were to evaluate and relate changesin DOM fluorescence to DBP formation during sequenced UV pro-cesses (i.e., UV followed by chlorine or chloramine). Settled watercollected from a drinking water utility treating surface water wasexposed to UV, chlorine, and chloramine and analyzed by EEMfluorescence spectroscopy and for a suite of DBPs. Fluorescencewas also measured on samples treated with UV alone (prior tochlorination/chloramination), and all fluorescence data were ana-lyzed using PARAFAC. Results were compared to samples withoutUV treatment but with a chlorine/chloramine dose adjusted toobtain a similar target residual after 24 h. A subset of sampleswere spiked with additional bromide and nitrate to investigate therole of inorganic precursors. UV doses ranging from disinfection(40–186 mJ/cm2) to higher doses (1000 mJ/cm2) were applied. Arecent utility survey showed that many full-scale UV disinfectionplants were delivering higher doses than originally designed for asa result of improved upstream processes [26], so the investigationof higher UV doses is relevant.

2. Materials and methods

2.1. Sample collection

Primary disinfection with UV during drinking water treat-ment is usually applied to settled water following coagula-tion/flocculation/sedimentation. Moreover, reverse osmosis (RO)concentration has been demonstrated as a method through whichhigh organic carbon recoveries (80–99%) can be obtained and theoriginal source water reactivity can be preserved [27,28]. There-fore, in this study, settled water from the Orange Water and SewerAuthority drinking water treatment plant (Carrboro, NC, USA) wasconcentrated by a factor of 15 (by volume) using a custom-builtportable RO system so that a large amount of DOM could be col-lected to provide the same matrix for a series of experiments (seeSupporting Information (SI) Text S1). The concentrated DOM char-acter was monitored by EEM fluorescence, dissolved organic carbon(DOC), and UV/visible absorbance and remained relatively constantbetween experiments, which took place over three months (changein fluorescence index [16,29] less than 0.5%; change in specificUV absorbance at 254 nm (SUVA254) less than 7%). SUVA254 wascalculated by dividing the UV absorbance at 254 nm (in units ofm−1) by the dissolved organic carbon concentration (mg C/L). Prior

to an experiment, the RO concentrate was diluted in laboratorygrade water (LGW) to obtain a DOC concentration of 2.7 mg C/L.LGW was prepared from a Dracor system (Durham, NC, USA)which pre-filters inlet 7 M� deionized water to 1 �m, removes

rdous

r0w(mrU8snmtccrl

2

fHdatpgd

2

7a(btmwatd

2

scgaDtdhh(ePdcA

2

Y

B.A. Lyon et al. / Journal of Haza

esidual disinfectants, reduces total organic carbon to less than.2 mg C/L with activated carbon resin, and removes ions to 18 M�ith mixed bed ion-exchange resins. Characteristics of ambient

unspiked) untreated water are provided in SI Table S1. DOC waseasured with a Shimadzu TOC-VCPH Analyzer (Shimadzu Corpo-

ation, Atlanta, GA, USA) following Standard Method 5310 [30].V/visible absorbance was measured using a Hewlett Packard452A spectrophotometer (Agilent, Santa Clara, CA, USA). A sub-et of samples were spiked with additional bromide (1 mg/L) oritrate (10 mg N/L) in the sodium salt form (Fisher ACS grade, Ther-oFisher Scientific, Waltham, MA, USA). These levels were chosen

o investigate trends, although it is recognized that these are higheroncentrations than those typically observed in surface waters. Foromparison, the United States Environmental Protection Agencyegulates nitrate in drinking water at a maximum contaminantevel of 10 mg N/L [31].

.2. UV treatment

UV treatment was performed using quasi-collimated irradiationrom a custom-built unit containing a 550 W MP lamp (Ace-anovia, Vineland, NJ, USA) with a 4-in. aperture similar to systemsescribed by Bolton and Linden [32]. Samples were retained in

250-mL Pyrex crystallization dish and stirred during irradia-ion. Constant sample temperature was maintained at 20–25 ◦C bylacing the dish in a copper coil which was connected to a pro-rammable refrigerated recirculating water unit. UV doses wereetermined as previously described [33].

.3. Chlorination/chloramination

Immediately following irradiation, samples were buffered to pH.5 with 5 mM phosphate buffer and dosed with chlorine from

dilution of a concentrated sodium hypochlorite stock solutionFisher laboratory grade, 5.6–6%) or pre-formed monochloramine,ased on a target residual of 1.0 ± 0.4 mg Cl2/L after 24 h of contactime, calculated from demand tests performed prior to treat-

ent. For the full experiment, duplicate samples were treatedith UV-chlorine/chloramine and held in chlorine demand-free

nd headspace-free 250-mL amber glass bottles with caps and poly-etrafluoroethylene (PTFE)-lined septa for 24 h at 20 ◦C. Additionaletails are provided in SI Text S2.

.4. Disinfection byproduct analysis

After 24 h, chlorine/chloramine residuals were measured andamples for DBP analysis were transferred to 60-mL glass vialsapped with PTFE-lined septa and containing l-ascorbic acid (ACSrade Sigma Chemical Co., St. Louis, MO, USA) as a quenchinggent. Quenched samples were stored headspace-free at 4 ◦C untilBP analysis, which was carried out as two separate extrac-

ions: (1) THMs (four regulated species: trichloro-, bromodichloro-,ibromochloro- and trichloro-methane), 4 haloacetonitriles, twoaloketones, two halonitromethanes, chloral hydrate, and 11aloacetamides were co-extracted within 24 h of quenching; and2) cyanogen chloride was extracted within 48 h. DBPs werextracted with methyl tert-butyl ether and analyzed on a Hewlett-ackard 5890 gas chromatograph with 63Ni electron captureetector following the procedures described by Sclimenti et al. foryanogen chloride [34] and Weinberg et al. for all other DBPs [35].dditional details of DBP analysis are described in SI Text S3.

.5. Fluorescence

A Fluorolog-321 fluorescence spectrophotometer (Horiba Jobinvon, Edison, NJ, USA) with a charge-coupled device detector was

Materials 264 (2014) 411– 419 413

used to measure fluorescence and generate EEM data. EEMs werecollected for treated samples at excitation wavelengths from 250to 450 nm at 5 nm intervals and emission wavelengths from 320to 550 nm at 1 nm intervals. Chlorine/chloramine residuals did notinterfere with sample fluorescence and were not quenched priorto fluorescence analysis. Samples for fluorescence measurementwere transferred to chlorine demand-free and headspace-free 25-mL amber glass vials and stored at 4 ◦C until analysis (within2 days for chlorinated samples, 3 days for chloraminated sam-ples, and 2–7 days for samples treated with UV only). Changesin fluorescence over the holding times were minimal, shown inSI Figs. S1–S3. Prior to analysis, samples were allowed to cometo room temperature. Samples and LGW blanks (collected daily)were corrected for instrument response by multiplying by themanufacturer-supplied excitation/emission correction factors, andthen blank EEMs were subtracted from sample EEMs. EEM inten-sities were normalized to the area under the water Raman peak(�ex = 350 nm, �em = 368–450 nm) of the LGW blank to obtain datain Raman Units. Sample absorbance was measured with an OceanOptics USB4000 spectrometer (Ocean Optics, Dunedin, FL, USA).A total of 269 EEMs were collected, consisting of 28 control sam-ples with no treatment, 84 UV-only treated, 20 chlorinated, 19chloraminated, 60 UV + chlorine, and 58 UV + chloramine-treated.A four-component PARAFAC model was developed using Matlab(Version 7.12.0, Mathworks, Natick, MA, USA) and the DOMFluortoolbox following the tutorial by Stedmon and Bro [36] to resolvethe components that explained variability across the entire dataset.The proper number of components was chosen using four-waysplit-half analysis, which compares the excitation and emissionloadings (spectra) between the model and four separate splits of thedataset using Tucker Congruence Coefficients [36,37] and by visual-ization of residual EEMs (difference between model and measuredexcitation/emission loadings) and sum-of-squared error (SSE).Two-, three- and four-component models could be split-half val-idated but five- and six-component models were not validated andwere, therefore, discarded. Residual EEMs of the four-componentmodel contained mostly background noise and explained 99.9% ofthe variance, while two- and three-component models had consid-erable residual EEMs and higher SSE values (SSE = 95.7, 24.4, and13.6 for random initialization of two-, three-, and four-componentmodels, respectively), so the four-component model was selected.The excitation and emission loadings of the four components in themodel compared to individual data validation splits are shown inSI Fig. S4. Fluorescence intensities are reported as the maximumfluorescence (Fmax) in Raman units, which is a unique value foreach component in every sample that correlates with the relativeamount of that fluorescing component [36,38]. A subset of sampleswere analyzed in triplicate (same sample measured three times)to determine the analytical reproducibility of component quan-tification. The relative standard deviation of Fmax values betweentriplicates ranged from 0.1 to 5.8%, with an average of 1.6%.

3. Results and discussion

3.1. DOC and spectral absorption characterization

Any changes in DOC values from UV treatment were withinexperimental error (relative percent difference, RPD, betweenexperimental duplicates ≤13%), shown in SI Table S2. The lowUV/visible absorbance of the settled water (A254 = 0.04 cm−1) priorto UV treatment and the low sensitivity of the spectrophotome-

ter made it difficult to evaluate the shifts in absorbance spectra ofthe water after UV and chlorine treatments. The slope ratio val-ues (S275–295:S350–400 [39]) ranged from 0.15 to 2.45, but the RPDsbetween experimental duplicates were greater than 20%. The low

4 rdous

sUeo(

3

ooawoewpct[t[adhtf[pflct(

3

o(tshtsrlFic

TDs

14 B.A. Lyon et al. / Journal of Haza

ensitivity of both measurements combined with non-specificity ofV/visible absorbance to detect changes in DOM character furthermphasizes the need for alternative sensitive measures such as flu-rescence spectroscopy to evaluate the effect of water treatmentse.g., UV, chlorine, and chloramine) on DOM.

.2. PARAFAC analysis

A four-component PARAFAC model was validated for a rangef sample treatment conditions. Similar components have beenbserved in previous studies [18,19,40–42] and have exhibited

range of removal/transformation efficiencies through drinkingater and wastewater treatment but ultimately persist to vari-

us degrees in finished water [19,20]. The presence of multiplemitting fluorophores within the complex DOM solution in naturalaters makes it difficult to definitively associate FDOM com-onents to something other than broad humic- or protein-likelasses. For example, components two and three (C2 and C3) con-ribute to the ubiquitously identified humic-like peaks A and C41], and are enriched in the hydrophobic humic substances frac-ion of DOM relative to unfractionated DOM in filtered water43]. Further, humic FDOM represented by C2 and C3 have beenssociated with terrestrially-derived material such as lignin break-own products [44], aromatic carbon content [16,43], and DOMaving the optical properties of degraded lignin [45]. The excita-ion and emission maxima of C1, also considered as a humic-likeraction of DOM, correspond to recent autochthonous production42] or wastewater organic matter [40]. The protein-like com-onent C4 has excitation and emission maxima similar to theuorescence of aromatic amino acids [16,40]. On the basis of theseomparisons, this paper follows the conventions for the descrip-ions of FDOM as represented by PARAFAC components C1 to C4Table 1).

.3. Effect of UV treatment on PARAFAC components

The Fmax values of C2 and C4 decreased following UV treatmentf ambient samples, while those for C1 and C3 were unaffectedFig. 1). C2 and C4 Fmax values were reduced 27% and 66%, respec-ively, at the highest UV dose (1000 mJ/cm2). Tryptophan absorbstrongly between 200 and 300 nm [46], and because MP UV lampsave a polychromatic output in this range (SI Fig. S5), the reac-ivity of tryptophan-like C4 to MP UV would be expected. FDOM,imilar to the humic-like component “A”, absorbs light in the UVCegion of 100–280 nm [18,41], which overlaps with the MP UV

amp emission and is consistent with the observed decrease in C2max from 1000 mJ/cm2 MP UV. In contrast to ambient samples,rradiation of nitrate-containing samples decreased the Fmax of allomponents. This suggests that all components were amenable to

able 1escription of four components (C1–C4) identified in the PARAFAC model, their excitatio

imilar peaks identified in previous work.

Component �ex/�em Description

C1 <250(320)/414 Microbial humic-like

C2 <250(360)/434 Humic-like

C3 <250(360)/489 Terrestrial humic-lik

C4 <250(280)/357 Protein,tryptophan-like

Materials 264 (2014) 411– 419

indirect photolysis (i.e., reaction with hydroxyl radicals producedfrom photolysis of nitrate). Nitrate absorbs strongly below 240 nm,which overlaps with the MP UV emission (SI Fig. S5). UV photolysisof nitrate generates nitrogen dioxide (•NO2) and hydroxyl radicals(•OH) with decreasing quantum yield (˚�) as wavelength increasesbetween 200 and 300 nm (˚205 nm = 0.129 and ˚253.7 nm = 0.037)[47]. Hydroxyl radicals have been shown to react relatively uns-electively with DOM [48], and their formation in the UV-treatednitrate-spiked settled water explains the impact on the Fmax valuesof all components in this current study (SI Table S3).

The presence of bromide resulted in greater Fmax decreasesin samples treated with UV doses of 186 mJ/cm2 (C2 and C4) or1000 mJ/cm2 (all components) compared to ambient samples (SITable S3). One explanation could be the formation of reactive halo-gen species from direct or indirect photolysis of bromide (e.g.,Br2

•−, BrOH•−), which have been shown to contribute to DOMdegradation and react more selectively with electron-rich chro-mophores than hydroxyl radicals [49]. However, this increased Fmax

reduction was not observed when comparing nitrate-spiked withbromide- and nitrate-spiked samples (SI Table S3), suggesting thata maximum possible Fmax decrease may have already been reachedfrom the reactive species formed from nitrate photolysis. Nitratewas spiked at 10 mg N/L (0.71 mM) compared to bromide at 1 mg/L(0.013 mM), and based on the overlap of their respective absorp-tion spectra with the MP UV output, nitrate would be expectedto produce more reactive species than bromide following MP UVirradiation.

The order of component reactivity to UV treatment of ambi-ent samples was: C4 > C2 > C1∼C3, which is opposite to the trendthat is typically observed for sunlight photolysis in natural systems[40], suggesting that findings on DOM reactivity under sunlightphotolysis conditions may not be applicable to predict behaviorin engineered UV systems. Hofbauer and Andrews observed a 28%decrease in total fluorescence intensity when 3000 mJ/cm2 MP UVwas applied to a Suwannee River DOM solution but they did notinvestigate the effect of UV on the underlying fluorescing compo-nents [50]. A study by Murphy and colleagues that used PARAFACto evaluate the effects of UV and other advanced wastewater treat-ment processes on FDOM showed inconsistent patterns for theeffect of UV on the Fmax of individual components across six dif-ferent treatment plants but, in general, component fluorescenceintensity decreased following UV treatment and to a lesser extentwhen chlorine was applied prior to UV [20]. In a few cases, the Fmax

of several components appeared to increase following UV, althoughit was not clear whether these changes were within the associated

analytical/experimental error. In addition, the UV dose or lamp type(i.e., medium vs. low pressure) used in each treatment plant was notreported, which may have contributed to the inconsistent observedeffect of UV on Fmax.

n and emission peaks (with secondary peaks in parentheses), and a comparison to

Comparison to past work Reference

C6: <250(320)/400 [40]C2: 250(320)/410 [19]Peaks C or A [41]C4: <250(360)/440 [40]P8: <260(355)/434 [42]C5: 250(340)/440 [19]

e C3: 270(360)/478 [18]C1: 260(360)/480 [19]Peak T [41]C7: 280/344 [40]C4: <250(290)/360 [19]

B.A. Lyon et al. / Journal of Hazardous Materials 264 (2014) 411– 419 415

F mum

w expe

3c

widoscrhsom3elcpowtiasr

bnrwv

ig. 1. Effect of UV-dosed settled water (ambient and spiked) on component maxiere 1 mg/L bromide and 10 mg N/L nitrate. Bar height represents average between

.4. Effect of UV-chlorine/chloramine treatment on PARAFAComponents

Chlorination decreased the Fmax of all components (Fig. 2),hich is in agreement with past work [20–22]. The reduction

n fluorescence intensity following chlorination is thought to beue to oxidation of FDOM components, such as the breakdownf carbon double bonds in aromatic molecules and halogen sub-titution reactions [38,51]. The order of component reactivity tohlorination in ambient samples was: C3 > C4 > C2 > C1, with Fmax

eductions of 67%, 53%, 46%, and 37%, respectively. While theumic-like C3 was found to be most affected by chlorine in thistudy, Beggs and Summers found that the fluorescence intensityf protein-like components was decreased more than that of aro-atic humic-like upon chlorination (average of 67% compared to

0% reduction, respectively) [21]. One explanation for this differ-nce is that the DOM source material in that study was pine needleeachate, which had a higher protein-like fluorescence contributionompared to that of humic-like components that were the moreredominant fraction of the DOM used in this current work. Twof the four identified components in the Beggs and Summers studyere polyphenolic/protein-like, and they accounted for 78–84% of

he fluorescence in coagulated fresh leachate samples. In compar-son, the protein/tryptophan-like component in the present studyccounted for 24% of the fluorescence in an ambient sample, theource of which was coagulated/settled surface water and is moreepresentative of a typical drinking water source.

Treatment of ambient samples with 1000 mJ/cm2 UV followedy chlorine resulted in Fmax decreases for C1 and C3 that were

ot observed for those components when treated with either chlo-ine or UV alone. This suggests that UV produced precursors thatere more amenable to reaction with chlorine, which was pre-

iously shown [33] and is consistent with the observed increase

fluorescence intensity (Fmax, in Raman units). Spiking amounts, where applicable,rimental duplicate samples, with the duplicate values shown by inset lines.

in chlorine demand of ambient samples that were pretreated with1000 mJ/cm2 UV (SI Table S4). Samples were compared on an equiv-alent targeted residual basis, so higher chlorine doses were appliedto the 1000 mJ/cm2 UV-treated samples, which could have also con-tributed to the greater Fmax decrease that was observed for C1 andC3.

The addition of bromide prior to chlorination reduced Fmax forall components compared to ambient samples (47%, 30%, 30%, and9% Fmax reduction for C1, C2, C3, and C4, respectively), shown inSI Table S5. This effect of bromide spiking prior to chlorinationon DOM fluorescence has not been previously demonstrated, tothe best of our knowledge. Bromide can be oxidized by free chlo-rine (HOCl) to form hypobromous acid (HOBr), which reacts withDOM faster than aqueous chlorine, and when both are present,bromine tends to act more as a substituting agent while chlorinereacts preferentially as an oxidant [52,53]. These results suggestthat the presence of HOBr results in more breakdown of fluorescingDOM constituents than HOCl alone and/or that fluorescence is beingquenched by bromine substitution reactions [38]. Bromide spikingdecreased component Fmax during chlorination with or without UV,while nitrate required UV treatment to affect Fmax.

Changes in Fmax were smaller for chloramine treatment (Fig. 3)compared to chlorination, which is consistent with the lower chlo-ramine vs. chlorine demand of samples (SI Table S4). Fmax valuesof C3 and C4, which were the most affected by chlorine, weredecreased 15–16% for ambient samples treated with chloramine,compared to no treatment, while C1 and C2 Fmax values did notchange following chloramination. The presence of bromide dur-ing chloramination had no effect on Fmax compared to the same

treatment in ambient samples. Bromide can be oxidized by chlo-ramine to produce active bromine species, including NH2Br, NHBr2,and NHBrCl [54], but their formation in a surface water containingbromide and treated with preformed monochloramine at pH 7.5

416 B.A. Lyon et al. / Journal of Hazardous Materials 264 (2014) 411– 419

Fig. 2. Effect of UV-dosed settled water (ambient and spiked) followed by chlorination on component Fmax. Spiking amounts, where applicable, were 1 mg/L bromide and10 mg N/L nitrate. Bar height represents average between experimental duplicate samples, with the duplicate values shown by inset lines.

Fig. 3. Effect of UV-dosed settled water (ambient and spiked) followed by chloramination on component Fmax. Spiking amounts, where applicable, were 1 mg/L bromide and10 mg N/L nitrate. Bar height represents average between experimental duplicate samples, with the duplicate values shown by inset lines.

B.A. Lyon et al. / Journal of Hazardous Materials 264 (2014) 411– 419 417

Table 2Effect of MP UV followed by chlorination (HOCl) or chloramination (NH2Cl) on the formation of DBPs in ambient and nitrate-spiked samples (10 mg N/L). Values representthe average between experimental duplicate samples (RPD ≤10%).

Spiking condition UV dose (mJ/cm2) Post-disinfectant Concentration (�g/L)

THM4a Trichloronitromethane Tribromonitromethane Chloral hydrate Cyanogen chloride

Ambient

0

HOCl

82 0.2 <0.1 7.8 <0.140 84 0.2 <0.1 9.8 <0.1

186 89 0.3 <0.1 15 <0.11000 111 0.5 <0.1 32 <0.1

Nitrate

0

HOCl

81 0.2 <0.1 8.1 <0.140 85 1.3 0.2 11 <0.1

186 95 4.0 0.3 18 <0.11000 109 8.0 0.6 32 <0.1

Ambient

0

NH2Cl

1.0 <0.1 0.1 0.2 0.440 1.1 <0.1 0.2 0.2 0.4

186 1.0 <0.1 0.2 0.2 0.41000 0.8 <0.1 0.1 0.2 0.8

Nitrate

0

NH2Cl

1.1 <0.1 0.1 0.2 0.540 1.0 0.2 0.1 0.2 0.6

ane co

(bi

3r

cdrcmbUpgwoct1

tcwwdnsaDoSfndwnnipc

Chloral hydrate formation increased with increasing UV dosewhen followed by chlorination, compared to its formation in sam-ples treated with chlorine alone. Nitrate spiking had no effecton chloral hydrate formation, but samples spiked with bromide

R² = 0.96

R² = 0.96

0.0

0.2

0.4

0.6

0.8

1.0

0 5 10 15 20 25 30 35

C2

Fm

ax

(C/C

0)

R² = 0.99

R² = 0.97

0.0

0.2

0.4

0.6

0.8

1.0

0 5 10 15 20 25 30 35

C4

Fm

ax

(C/C

0)

[Chloral hydrate] (µg/L)

(a)

(b)

186 0.9 0.51000 0.6 0.8

a THM4 = sum of trichloro-, bromodichloro-, dibromochloro- and tribromo-meth

i.e., the source waters used for this work) is slow and not likely toe significant [55,56]. This was also reflected in the lack of change

n chloramine demand with the addition of bromide (SI Table S4).

.5. Comparison of disinfection byproduct and fluorescenceesults

Only a subset of DBP results will be discussed here, focusing onompounds whose formation was affected by UV. The use of UV atisinfection doses (40–186 mJ/cm2) prior to chlorination or chlo-amination had little effect on the formation of the four regulatedhlorine- and bromine-containing THMs (THM4), which is in agree-ent with previous research [10,11]. However, when ambient or

romide-/nitrate-spiked samples were treated with 1000 mJ/cm2

V prior to chlorination, THM4 formation increased 30–35% com-ared to chlorination alone, suggesting that with higher doses, UVenerates THM4 precursors that are more amenable to reactionith chlorine. This is consistent with the greater Fmax decrease

bserved for C1 and C3 when UV treatment was used prior tohlorination (compared to samples treated with chlorine only) andhe increased chlorine demand observed for samples treated with000 mJ/cm2 UV (SI Table S4).

UV treatment increased the formation of trichloronitromethane,ribromonitromethane, and chloral hydrate when followed byhlorination or chloramination, compared to samples treatedith chlorine or chloramine alone. Cyanogen chloride formationas also increased by UV with subsequent chloramination. Theegree to which these species were affected was dependent onitrate/bromide spiking conditions and UV dose. Results for aubset of treatment conditions and DBPs are shown in Table 2;dditional results and discussion are published elsewhere [33].BP formation in bromide-spiked samples and the concentrationsf individual THM4 species are shown in SI Table S6 and SI Fig.6, respectively. The increased formation of halonitromethanesollowing UV irradiation and chlorination or chloramination initrate-spiked samples is hypothesized to result from the pro-uction of reactive nitrogen species from photolysis of nitrate,hich can act as nitrating agents toward aromatic DOM. Theseitrated organics then generate halonitromethanes after chlori-

ation/chloramination [14,57]. The presence of nitrate during UV

rradiation decreased Fmax values for all components when com-ared to ambient samples (SI Table S3), so limited conclusionsan be drawn about which type(s) of DOM were responsible for

0.1 0.2 0.90.2 0.3 1.9

ncentrations.

trichloro- and tribromonitromethane formation. When sampleswere treated with UV followed by chlorine, the presence of nitrateaffected C3 more than the other components, with respect toincreased reactivity toward chlorine following 40 or 186 mJ/cm2

UV (SI Table S5). However, it is difficult to correlate this trendwith the proposed halonitromethane formation mechanism, sincenitrated DOM is not necessarily more reactive with chlorine thanits un-nitrated counterpart [58].

Fig. 4. Correlation between change in (a) C2 Fmax and (b) C4 Fmax with the formationof chloral hydrate in ambient (black shapes) and nitrate-spiked (white shapes) sam-ples treated with MP UV + chlorine. C0 refers to the Fmax value in the correspondingsample treated with chlorine only.

418 B.A. Lyon et al. / Journal of Hazardous

R² = 0.91

0.0

0.2

0.4

0.6

0.8

1.0

0.0 0.5 1. 0 1. 5 2.0

C2 F

max

(C/C

0)

R² = 0.79

0.0

0.2

0.4

0.6

0.8

1.0

0.0 0.5 1. 0 1. 5 2.0

C4 F

max

(C/C

0)

[Cyanog en chlor ide] (µg/L)

(a)

(b)

Fig. 5. Correlation between the change in (a) C2 Fmax and (b) C4 Fmax with the for-mat

hbmstctscbmcrpsfbshtn

bt[timTreid

Residue organic mixtures from drinking water show in vitro mutagenic and

ation of cyanogen chloride following MP UV + chloramine treatment of ambientnd nitrate-spiked samples. C0 refers to the Fmax value in the corresponding samplereated with chloramine only.

ad lower chloral hydrate formation, likely due to a shift to theromine-substituted species (not measured for this work). Theechanism for increased chloral hydrate formation with UV and

ubsequent chlorination is thought to be through the initial forma-ion of acetaldehyde, known to be produced from UV [11], whichan then generate chloral hydrate upon chlorination [59]. The reac-ivity of C2 and C4 to UV suggests that DOM with characteristicsimilar to these components was responsible for the production ofhloral hydrate precursors during UV irradiation. This is supportedy the correlation between the decrease in C2 and C4 Fmax and for-ation of chloral hydrate in samples treated with UV followed by

hlorination (R2 values of 0.96–1.00 and 0.95–0.99 for C2 and C4,espectively), shown in Fig. 4 for ambient and nitrate-spiked sam-les and SI Fig. S7 for bromide- and bromide- and nitrate-spikedamples. Although nitrate-spiking did not impact chloral hydrateormation, C2 and C4 Fmax values were decreased to a greater extenty UV in the presence of nitrate, likely due to indirect photoly-is. Thus, the linear regressions between C2 or C4 Fmax and chloralydrate formation exhibited different slopes for ambient comparedo nitrate-spiked samples, but the strength of the correlations wasot affected.

Cyanogen chloride is typically observed as a chloraminationyproduct, and previous work has identified amino acids includingryptophan and other organic nitrogen compounds as precursors60,61]. Therefore, constituents similar to C4 would likely be impor-ant precursors for cyanogen chloride during chloramination and,ndeed, the C4 Fmax value was decreased with chloramine treat-

ent alone when formation of cyanogen chloride was observed.he use of UV prior to chloramination increased cyanogen chlo-

ide formation compared to chloramination alone, and to a greaterxtent in nitrate-spiked samples. One proposed pathway for thisncrease is through the formation of formaldehyde, which can pro-uce cyanogen chloride upon chloramination and is known to be

Materials 264 (2014) 411– 419

generated during engineered UV processes [10,62]. The observeddecrease in C2 and C4 Fmax values following UV treatment suggeststhat UV-induced precursors responsible for increased cyanogenchloride formation originated from DOM similar to these compo-nents. Decreases in both C2 and C4 Fmax values correlated withcyanogen chloride formation in samples treated with UV followedby chloramine (R2 values of 0.91 and 0.79 for C2 and C4, respec-tively), shown in Fig. 5.

4. Conclusions

The use of fluorescence spectroscopy combined with DBP anal-ysis allowed for an investigation into the fluorescence character ofDOM in coagulated waters and the impact of MP UV treatment onthe reactivity of these FDOM components to chlorine/chloramine.Overall, the humic-like C2 and protein/tryptophan-like C4 werethe most susceptible to UV treatment and most strongly cor-related with UV-induced DBP formation. Correlations betweenFDOM and DBP type and concentration suggest that PARAFACanalysis of settled waters can assist in drinking water processdesign that optimizes for controlling DBP formation during UV-chlorine/chloramine drinking water treatment or other treatmentprocesses. For example, in waters containing high proportions ofC2 or C4-like DOM, additional pretreatment targeting these com-ponents could be used to minimize UV-induced DBP formation.These findings should be confirmed by evaluating a range of treatedsource waters.

Acknowledgements

This work was supported by Water Research Foundation Project4019 (project manager Alice Fulmer), a Royster Society of FellowsDissertation Completion Fellowship, and a Gillings DissertationAward. The authors thank Rachel Monschein and Orange Waterand Sewer Authority for providing laboratory space to collect watersamples. Thanks also to Ariel Atkinson, Rebecca Milsk, and Zoe Wol-szon for assistance with DBP extractions and Katie Harrold for helpwith the Fluorolog and PARAFAC modeling.

Appendix A. Supplementary data

Supplementary data associated with this article can be found,in the online version, at http://dx.doi.org/10.1016/j.jhazmat.2013.10.065.

References

[1] R.F. Christman, D.L. Norwood, D.S. Millington, J.D. Johnson, A.A. Stevens, Iden-tity and yields of major halogenated products of aquatic fulvic acid chlorination,Environ. Sci. Technol. 17 (1983) 625–628.

[2] D.R. Lovley, J.D. Coates, E.L. Blunt-Harris, E.J.P. Phillips, J.C. Woodward, Humicsubstances as electron acceptors for microbial respiration, Nature 382 (1996)445–448.

[3] S. Markager, W.F. Vincent, Spectral light attenuation and absorption of UV andblue light in natural waters, Limnol. Oceanogr. 45 (2000) 642–650.

[4] R. Maranger, M.J. Pullin, Elemental complexation by dissolved organic matter inlakes: implications for Fe speciation and the speciation and the bioavailabilityof Fe and P, in: S.E.G. Findlay, R.L. Sinsabaugh (Eds.), Aquatic Ecosystems –Interactivity of Dissolved Organic Matter, Academic Press, San Diego, CA, 2003,pp. 185–214.

[5] B. Kwon, S. Lee, J. Cho, H. Ahn, D. Lee, H.S. Shin, Biodegradability, DBP formation,and membrane fouling potential of natural organic matter: characterizationand controllability, Environ. Sci. Technol. 39 (2005) 732–739.

[6] J.C. Loper, D.R. Lang, R.S. Schoeny, B.B. Richmond, P.M. Gallagher, C.C. Smith,

transforming activity, J. Toxicol. Environ. Health A 4 (1978) 919–938.[7] K.P. Cantor, C.F. Lynch, M.E. Hildesheim, M. Dosemeci, J. Lubin, M. Alavanja, G.

Craun, Drinking water source and chlorination byproducts I. Risk of bladdercancer, Epidemiology 9 (1998) 21–28.

rdous

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

[

B.A. Lyon et al. / Journal of Haza

[8] U.S. EPA. (Environmental Protection Agency), National Primary Drinking WaterRegulations: Stage 2 Disinfectants and Disinfection Byproducts Rule, Fed. Reg-ist. 71 (2006) 387–493.

[9] S.W. Krasner, H.S. Weinberg, S.D. Richardson, S.J. Pastor, R. Chinn, M.J. Sclimenti,G.D. Onstad, A.D. Thruston, Occurrence of a new generation of disinfectionbyproducts, Environ. Sci. Technol. 40 (2006) 7175–7185.

10] J.P. Malley, J.P. Shaw, J.R. Ropp, Evaluation of the Byproducts Produced fromthe Treatment of Groundwaters with Ultraviolet Irradiation, American WaterWorks Association and Water Research Foundation, Denver, CO, 1995.

11] W. Liu, S.A. Andrews, J.R. Bolton, K.G. Linden, C.M. Sharpless, M. Stefan, Compar-ison of disinfection byproduct (DBP) formation from different UV technologiesat bench scale, Water Sci. Technol. 2 (2002) 515–521.

12] A.D. Dotson, V.O.S. Keen, D. Metz, K.G. Linden, UV/H2O2 treatment of drink-ing water increases post-chlorination DBP formation, Water Res. 44 (2010)3703–3713.

13] M.J. Plewa, E.D. Wagner, M.G. Muellner, K.-M. Hsu, S.D. Richardson, Compara-tive mammalian cell toxicity of N-DBPs and C-DBPs, in: T. Karanfil, S.W. Krasner,P. Westerhoff, Y. Xie (Eds.), Disinfection By-Products in Drinking Water: Occur-rence, Formation, Health Effects, and Control, ACS Symposium Series 995,Oxford University Press, New York, 2008, pp. 36–49.

14] D.A. Reckhow, K.G. Linden, J. Kim, H. Shemer, G. Makdissy, Effect of UV treat-ment on DBP formation, J. Am. Water Works Assoc. 102 (2010) 100–113.

15] D.M. McKnight, G.R. Aiken, Sources and age of aquatic humus, in: D.O. Hesson,L.J. Tranvik (Eds.), Aquatic Humic Substances: Ecology and Biogeochemistry,Springer-Verlag, New York, 1998, pp. 9–39.

16] R.M. Cory, D.M. McKnight, Fluorescence spectroscopy reveals ubiquitous pres-ence of oxidized and reduced quinones in dissolved organic matter, Environ.Sci. Technol. 39 (2005) 8142–8149.

17] N. Maie, N.M. Scully, O. Pisani, R. Jaffé, Composition of a protein-like fluorophoreof dissolved organic matter in coastal wetland and estuarine ecosystems, WaterRes. 41 (2007) 563–570.

18] C.A. Stedmon, S. Markager, R. Bro, Tracing dissolved organic matter in aquaticenvironments using a new approach to fluorescence spectroscopy, Mar. Chem.82 (2003) 239–254.

19] S.A. Baghoth, S.K. Sharma, G.L. Amy, Tracking natural organic matter (NOM)in a drinking water treatment plant using fluorescence excitation-emissionmatrices and PARAFAC, Water Res. 45 (2011) 797–809.

20] K.R. Murphy, A. Hambly, S. Singh, R.K. Henderson, A. Baker, R. Stuetz, S.J. Khan,Organic matter fluorescence in municipal water recycling schemes: toward aunified PARAFAC model, Environ. Sci. Technol. 45 (2011) 2909–2916.

21] K.M.H. Beggs, R.S. Summers, Character and chlorine reactivity of dissolvedorganic matter from a mountain pine beetle impacted watershed, Environ. Sci.Technol. 45 (2011) 5717–5724.

22] P. Roccaro, F.G.A. Vagliasindi, G.V. Korshin, Changes in NOM fluorescencecaused by chlorination and their associations with disinfection by-productsformation, Environ. Sci. Technol. 43 (2009) 724–729.

23] B. Hua, K. Veum, J. Yang, J. Jones, B. Deng, Parallel factor analysis of fluorescenceEEM spectra to identify THM precursors in lake waters, Environ. Monit. Assess.161 (2010) 71–81.

24] C.W. Grandersen, A.D. Pifer, J.L. Fairey, An improved chloroform surrogate forchlorine dioxide- and alum-treated waters, J. Am. Water Works Assoc. 105(2013) E103–E114.

25] D.W. Johnstone, N.P. Sanchez, C.M. Miller, Parallel factor analysis of excitation-emission matrices to assess drinking water disinfection byproduct formationduring a peak formation period, Environ. Eng. Sci. 26 (2009) 1551–1559.

26] A.D. Dotson, C.E. Rodriguez, K.G. Linden, UV disinfection implementation statusin United States water treatment plants, J. Am. Water Works Assoc. 104 (2012)E318–E324.

27] M. Kitis, J.E. Kilduff, T. Karanfil, Isolation of dissolved organic matter (DOM)from surface waters using reverse osmosis and its impact on the reactivity ofDOM to formation and speciation of disinfection by-products, Water Res. 35(2001) 2225–2234.

28] H. Song, O. Orr, Y. Hong, T. Karanfil, Isolation and fractionation of natural organicmatter: evaluation of reverse osmosis performance and impact of fractionationparameters, Environ. Monit. Assess. 153 (2009) 307–321.

29] D.M. McKnight, E.W. Boyer, P. Westerhoff, P.T. Doran, T. Kulbe, D.T. Andersen,Spectrofluorometric characterization of dissolved organic matter for indicationof precursor organic material and aromaticity, Limnol. Oceanogr. 46 (2001)38–48.

30] APHA (American Public Health Association), American Water Works Asso-ciation, and Water Environment Federation, in: Standard Methods for theExamination of Water and Wastewater, 20th ed., American Public Health Asso-ciation, Washington, DC, 1999.

31] U.S. EPA. (Environmental Protection Agency), National Primary Drinking WaterRegulations: Synthetic Organic Chemicals and Inorganic Chemicals, Final Rule,Fed. Regist. 57 (1992) 31776–31849.

32] J.R. Bolton, K.G. Linden, Standardization of methods for fluence (UV dose) deter-

mination in bench-scale UV experiments, J. Environ. Eng.-ASCE 129 (2003)209–216.

33] B.A. Lyon, A.D. Dotson, K.G. Linden, H.S. Weinberg, The effect of inorganic pre-cursors on disinfection byproduct formation during UV-chlorine/chloraminedrinking water treatment, Water Res. 46 (2012) 4653–4664.

[

Materials 264 (2014) 411– 419 419

34] M.J. Sclimenti, C.J. Hwang, G.E. Speitel, A.C. Diehl, The simultaneous determina-tion of cyanogen chloride and cyanogen bromide in chloraminated waters by asimplified microextraction GC/ECD technique, in: Proc. AWWA Water QualityTechnology Conference, Charlotte, NC, 1994, pp. 489–506.

35] H.S. Weinberg, S.W. Krasner, S.D. Richardson, A.D. Thruston Jr., The Occur-rence of Disinfection By-Products (DBPs) of Health Concern in Drinking Water:Results of a Nationwide DBP Occurrence Study, EPA/600/R/-02/068, US EPA,Athens, GA, 2002, pp. 365–373.

36] C.A. Stedmon, R. Bro, Characterizing dissolved organic matter fluorescence withparallel factor analysis: a tutorial, Limnol. Oceanogr. 6 (2008) 572–579.

37] U. Lorenzo-Seva, J.M.F.T. Berge, Tucker’s congruence coefficient as a meaningfulindex of factor similarity, Methodology 2 (2006) 57–64.

38] J.R. Lakowicz, Principles of Fluorescence Spectroscopy, 3rd ed., Springer, NewYork, 2006.

39] J.R. Helms, A. Stubbins, J.D. Ritchie, E.C. Minor, D.J. Kieber, K. Mopper, Absorptionspectral slopes and slope ratios as indicators of molecular weight, source, andphotobleaching of chromophoric dissolved organic matter, Limnol. Oceanogr.53 (2008) 955–969.

40] C.A. Stedmon, S. Markager, Resolving the variability in dissolved organic matterfluorescence in a temperate estuary and its catchment using PARAFAC analysis,Limnol. Oceanogr. 50 (2005) 686–697.

41] P.G. Coble, Characterization of marine and terrestrial DOM in seawater usingexcitation-emission matrix spectroscopy, Water Res. 51 (1996) 325–346.

42] K.R. Murphy, C.A. Stedmon, T.D. Waite, G.M. Ruiz, Distinguishing between ter-restrial and autochthonous organic matter sources in marine environmentsusing fluorescence spectroscopy, Mar. Chem. 108 (2008) 40–58.

43] R.M. Cory, D.M. McKnight, Y.-P. Chin, P. Miller, C.L. Jaros, Chemical character-istics of fulvic acids from Arctic surface waters: microbial contributions andphotochemical transformations, J. Geophys. Res. 112 (2007) G04S51.

44] P.J. Hernes, B.A. Bergamaschi, R.S. Eckard, R.G.M. Spencer, Fluorescence-basedproxies for lignin in freshwater dissolved organic matter, J. Geophys. Res. 114(2009) G00F03.

45] R. Del Vecchio, N.V. Blough, On the origin of the optical properties of humicsubstances, Environ. Sci. Technol. 38 (2004) 3885–3891.

46] G.D. Fasman (Ed.), Handbook of Biochemistry and Molecular Biology, Proteins,vol. I, 3rd ed., CRC Press, Cleveland, OH, 1976.

47] S. Goldstein, J. Rabani, Mechanism of nitrite formation by nitrate photolysisin aqueous solutions: the role of peroxynitrite, nitrogen dioxide, and hydroxylradical, J. Am. Chem. Soc. 129 (2007) 10597–10601.

48] P. Westerhoff, G. Aiken, G.L. Amy, J. Debroux, Relationships between the struc-ture of natural organic matter and its reactivity towards molecular ozone andhydroxyl radicals, Water Res. 33 (1999) 2265–2276.

49] J.E. Grebel, J.J. Pignatello, W. Song, W.J. Cooper, W.A. Mitch, Impact of halideson the photobleaching of dissolved organic matter, Mar. Chem. 115 (2009)134–144.

50] D.E.W. Hofbauer, S.A. Andrews, Influence of UV irradiation and UV/hydrogenperoxide oxidation process on natural organic matter fluorescence character-istics, Water Sci. Technol. 4 (2004) 41–46.

51] G.V. Korshin, M.U. Kumke, C.-W. Li, F.H. Frimmel, Influence of chlorination onchromophores and fluorophores in humic substances, Environ. Sci. Technol. 33(1999) 1207–1212.

52] J.J. Rook, A.A. Gras, B.G. van der Heijden, J. de Wee, Bromide oxidation andorganic substitution in water treatment, J. Environ. Sci. Health A 13 (1978)91–116.

53] P. Westerhoff, P. Chao, H. Mash, Reactivity of natural organic matter with aque-ous chlorine and bromine, Water Res. 38 (2004) 1502–1513.

54] T.W. Trofe, G.W. Inman, J.D. Johnson, Kinetics of monochloramine decomposi-tion in the presence of bromide, Environ. Sci. Technol. 14 (1980) 544–549.

55] A.C. Diehl, G.E. Speitel, J.M. Symons, S.W. Krasner, C.J. Hwang, S.E. Barrett,DBP formation during chloramination, J. Am. Water Works Assoc. 92 (2000)76–90.

56] M.J. Benotti, E.C. Wert, S.A. Snyder, C. Owen, R. Cheng, Role of Bromamines onDBP Formation and Impact on Chloramination and Ozonation, American WaterWorks Association and Water Research Foundation, Denver, CO, 2011.

57] A.D. Shah, A.D. Dotson, K.G. Linden, W.A. Mitch, Impact of UV disinfectioncombined with chlorination/chloramination on the formation of haloni-tromethanes and haloacetonitriles in drinking water, Environ. Sci. Technol. 45(2011) 3657–3664.

58] H. Gallard, U. von Gunten, Chlorination of phenols: kinetics and formation ofchloroform, Environ. Sci. Technol. 36 (2002) 884–890.

59] A. McKnight, D.A. Reckhow, Reactions of ozonation by-products with chlo-rine and chloramines, in: Proc. Annual AWWA Conference, Vancouver, BritishColumbia, 1992.

60] S.W. Krasner, M.J. McGuire, J.G. Jacangelo, N.L. Patania, K.M. Reagan, E.M. Aieta,The occurrence of disinfection by-products in US drinking water, J. Am. WaterWorks Assoc. 81 (1989) 41–53.

61] X. Yang, C. Fan, C. Shang, Q. Zhao, Nitrogenous disinfection byproducts for-

mation and nitrogen origin exploration during chloramination of nitrogenousorganic compounds, Water Res. 44 (2010) 2691–2702.

62] E.J. Pedersen, E.T. Urbansky, B.J. Marinas, D.W. Margerum, Formation ofcyanogen chloride from the reaction of monochloramine with formaldehyde,Environ. Sci. Technol. 33 (1999) 4239–4249.