the role of nitrogen in chromophoric and fluorescent dissolved organic matter formation

3 (2007) 46–60www.elsevier.com/locate/marchem

Marine Chemistry 10

The role of nitrogen in chromophoric and fluorescent dissolvedorganic matter formation

Erin J. Biers a, Richard G. Zepp b, Mary Ann Moran a,⁎

a Department of Marine Sciences, University of Georgia, Athens, GA 30602-3636, USAb U.S. Environmental Protection Agency, 960 College Station Road, Athens, GA 30605-2700, USA

Received 14 October 2005; received in revised form 12 April 2006; accepted 12 June 2006Available online 25 July 2006

Abstract

Microbial and photochemical processes affect chromophoric dissolved organic matter (CDOM) dynamics in the ocean. Someevidence suggests that dissolved nitrogen plays a role in CDOM formation, although this has received little systematic attention inmarine ecosystems. Coastal seawater incubations were carried out in the presence of model dissolved organic nitrogen (DON: aminosugars and amino acids) and dissolved inorganic nitrogen (DIN) compounds to assess their role in biological and photochemicalproduction of CDOM. For several of the dissolved N compounds, microbial processing resulted in a pulse of CDOM that was mainlylabile, appearing and disappearing within 7 days. In contrast, a net loss of CDOM occurred when no N was added to the microbialincubations. The greatest net biological CDOM formation was found upon addition of amino sugars (formation of fluorescent,mostly labile CDOM) and tryptophan (formation of non-fluorescent, refractory CDOM). Photochemical formation of CDOM wasonly found with tryptophan, the one aromatic compound tested. This CDOM was highly fluorescent, with excitation–emissionmatrices (EEMs) resembling those of terrestrial, humic-like fluorophores. The heterogeneity in CDOM formation from thiscollection of labile N-containing compounds was surprising. These compounds are common components of biopolymers and humicsubstances in natural waters and likely to contribute to microbially- and photochemically-produced CDOM in coastal seawater.© 2006 Elsevier B.V. All rights reserved.

Keywords: CDOM; Nitrogen; Microbial activity; Photochemical reactions; EEM; Dissolved organic matter

1. Introduction

Chromophoric dissolved organic matter (CDOM) isan important component of the dissolved organic matter(DOM) pool in aquatic ecosystems. It gives color towater, absorbing sunlight from the ultraviolet (UV)region of the solar spectrum into the visible. Because itabsorbs UV, CDOM protects aquatic organisms from

⁎ Corresponding author. Tel.: +1 706 542 6481; fax: +1 706 5425888.

E-mail address: [email protected] (M.A. Moran).

0304-4203/$ - see front matter © 2006 Elsevier B.V. All rights reserved.doi:10.1016/j.marchem.2006.06.003

otherwise damaging radiation (Arrigo and Brown, 1996;Gao and Zepp, 1998). It also produces photochemicalintermediates upon UVabsorption (Bushaw et al., 1996;Uher and Andreae, 1997; Xie et al., 1998; Miller et al.,2002), some of which are biologically labile (Moran andZepp, 1997). Therefore, changes in the CDOM poolaffect many aspects of ocean photo-biogeochemistry, aswell as complicate remote sensing of biological proper-ties of the ocean (Carder et al., 1991; Blough and DelVecchio, 2002).

A major source of CDOM to marine environments isterrestrial input (Meyers-Schulte and Hedges, 1986;

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http://dx.doi.org/10.1016/j.marchem.2006.06.003

47E.J. Biers et al. / Marine Chemistry 103 (2007) 46–60

Blough and Del Vecchio, 2002). Delivery of terrestrialDOM to coastal oceans via riverine runoff amounts toapproximately 0.25×1015 g C year−1, 30–70% ofwhich can be highly colored humic substances (Aiken etal., 1985; Deuser, 1988; Hedges et al., 1997; Cauwet,2002). On a local scale, resuspension of bottomsediments can lead to a release of CDOM into aquaticsystems (Boss et al., 2001). In addition, photooxidativealteration of non-colored DOM (Tissot and Welte, 1978;Hedges, 1988; Kieber et al., 1997) or Schiff basereactions involving the combination of a carbonyl-groupwith a primary amine to form an imine (Hedges, 1988)can be sources of colored compounds.

In situ biological activities have also been shown tobe sources of marine CDOM. Phytoplankton wereinitially implicated as major biological producers ofCDOM, either by excreting colored compounds (Bri-caud et al., 1981; Carder et al., 1989) or releasing themupon cell lysis (Momzikoff et al., 1992). More recently,the role of heterotrophic microbes in the formation ofCDOM has been recognized (Coble et al., 1998; Nelsonet al., 2004). These studies indicate that phytoplanktonand bacteria can act in concert to form CDOM, withphytoplankton providing substrates that heterotrophicbacteria process into colored compounds (Rochelle-Newall and Fisher, 2002).

There is evidence that dissolved nitrogen may beinvolved in CDOM production, although this has not yetbeen studied systematically in marine systems. Studiesbased in soils report that nitrogenous compounds arereadily incorporated into humic substances (Maillard,1913; Flaig, 1988; Hedges, 1988; Stevenson, 1994;Hedges et al., 2000) and produce molecules with agreater potential to absorb light (Nissenbaum andKaplan, 1972) and to fluoresce (Klapper et al., 2002).In natural aquatic systems, dissolved organic nitrogen(DON) is susceptible to photooxidation, suggesting thatat least some components may be light-absorbingmolecules (Bushaw et al., 1996; Reitner et al., 2002;Vähätalo and Zepp, 2005).

In this study, we investigated the role of defineddissolved nitrogen compounds in CDOM formation bymicrobial and photochemical mechanisms. The nitrogencompounds surveyed included both dissolved inorganicnitrogen (DIN) in the form of ammonia and nitrite, anddissolved organic nitrogen (DON) in the form of aminoacids and amino sugars. The amino acids selected forstudy were aspartic acid and glutamic acid, two of themost abundant amino acids in humic substances andnatural waters (Thurman, 1985; McCarthy et al., 1998),and tryptophan, an aromatic amino acid that absorbssunlight (Plummer, 1987). The amino sugars selected

for study were D-glucosamine, D-galactosamine, mura-mic acid, and D-mannosamine, common components ofbiopolymers including cell wall material (Stevenson,1994; Benner and Kaiser, 2003). We found thatmicrobial processing and photooxidation of some, butnot all, dissolved N compounds resulted in CDOMformation.

2. Materials and methods

2.1. Study site and sample collection

Seawater was collected in January, July, andNovember 2004 at high tide from Dean Creek, a tidalcreek on Sapelo Island, GA that drains a Spartinaalterniflora dominated marshland. Salinity of the watervaried little between collection times (20–25) as didfluorescence excitation emission spectra and sampleabsorption coefficients at 350 nm (7.69–8.89 m−1; seeSection 2.4 for analysis methods). Water was filtered on-site (1 μm pore-size, Poretics membrane filter),transported to the lab, and stored in the dark at 4 °Cfor <24 h before use. The 0.1–1.0 μm fraction wasconcentrated (20-fold) using a peristaltic tangential flowfiltration unit (Millipore) and used as a microbialinoculum. The remaining water was filtered twicethrough a 0.2 μm pore-size membrane filter (Poretics)and used as seawater medium (SW).

2.2. Microbial incubation experiments

To conduct microbial experiments, either artificial ornatural seawater medium was amended with a singlenitrogen source to either high (100 μM) or low (1 μM)final concentration. Artificial seawater medium (con-sisting of 20 g L−1 Sigma sea salts dissolved in distilledwater) was amended with a vitamin solution andphosphorus (KH2PO4; 62.5 nM in low N treatments,6.25 μM in high N treatments) to prevent nutrientlimitation during incubations. Natural seawater medium(consisting only of 0.1-μm-filtered coastal seawater)was not amended further. Nitrogen sources includedthree amino acids (L-glutamic acid, L-aspartic acid, andL-tryptophan), four amino sugars (D-galactosamine, D-glucosamine, muramic acid, and D-mannosamine), andtwo inorganic nitrogen sources (ammonium and nitrite).Control treatments were established as follows: nocarbon or nitrogen addition; carbon but no nitrogenaddition (added as succinate, the amine-free analog ofaspartic acid); and carbon and nitrogen addition withouta microbial inoculum (added as aspartic acid orglucosamine).

48 E.J. Biers et al. / Marine Chemistry 103 (2007) 46–60

Cultures (200 mL in 250 mL Erlenmeyer flasks) wereinoculated with the Dean Creek microbial concentrate toa final concentration equal to that in the original watersample (1×). Triplicate cultures for each nitrogen sourcewere gently shaken to keep samples aerated and wereincubated in the dark at 22.5 °C for 56 days. At regulartime points, 30 mL subsamples from each replicate werevacuum filtered (0.2 μm pore-size, Poretics) and storedin the dark at 4 °C until analysis.

2.3. Photochemical experiments

To conduct photochemical experiments, 0.2-μm-filtered tidal creek water was amended with a singlenitrogen source in either high (100 μM) or low (1 μM)final concentration. ASW was not used as a medium forthe photochemical experiments because of the likelyimportance of background DOM in photochemicalprocesses. Nitrogen sources were the same as thoseused for the microbial incubation experiments exceptthat muramic acid was not included. Control treatmentswere established as follows: no carbon or nitrogenaddition; and carbon but no nitrogen addition (added assuccinate). For each treatment, six 125 mL quartz flaskswere established, three of which were covered withaluminum foil. The flasks were put into a circulatingwater bath and exposed to 8 h of artificial sunlight at∼70% of the natural UV-A intensity of midday summersun at 30°N latitude. Artificial sunlight was generatedwith a xenon arc lamp (Atlas RM-65-3 Xenon Arc LightSystem, Atlas Electric Devices, Chicago, IL, USA). Thewater baths maintained a temperature between 4 and10 °C throughout the irradiation. Samples were storedin the dark at 4 °C until analysis.

2.4. Sample analysis

Sample absorbance was measured from 200 to700 nm (DU-640 Beckman spectrophotometer with aDI water reference) using either a 1 cm or a 5 cm pathlength quartz cell and converted to absorption coeffi-cients (αλ) using the following equation

ak ¼ 2:303Dk=L ð1Þ

where Dλ is the absorbance at wavelength λ and L is thepath length of the quartz cuvette in meters. In this study,we used absorption coefficients at 350 nm (α350) as theindex for CDOM concentration, similar to previousstudies (e.g. Moran et al., 2000). Differences in α350between treatments were assessed with ANOVAs andpaired t-tests (SPSS Inc., Chicago, IL) using a p-value

of 0.05 to determine significance. The spectral slopecoefficient (S) was calculated by using Sigma Plot(SPSS Inc.) to fit the absorption coefficients from 290 to500 nm to a nonlinear equation

ak ¼ ak0e�Sðk�k0Þ ð2Þ

where αλ is the absorption coefficient at wavelength λ,αλ0 at reference wavelength λ0 (412 nm), and S is thespectral slope coefficient (Zepp and Schlotzhauer, 1981;Blough and Green, 1995). The spectral slope coefficientwas calculated for each replicate sample and averaged.

Excitation–emission matrix spectra (EEMs) werecompiled for select samples using an ISA SPEXFluorolog 3–12 scanning fluorometer with R928Pdetector. Scans were run at 5 nm excitation intervals(230–500 nm) and 2 nm emission intervals (280–650 nm) with both excitation and emission slits set at5 nm band widths. Scans were corrected for instrumentconfiguration as well as Rayleigh and Raman scatterpeaks and converted into quinine sulfate equivalents(QSE) using a MATLAB® program developed by WadeSheldon (University of Georgia, USA; Zepp et al.,2004). Previously described excitation/emission peak-regions corresponding to fluorophore types (Coble,1996) were integrated in MATLAB® and displayed asQSE (Moran et al., 2000). Samples with high absor-bance were diluted to minimize inner filtering effects bydecreasing the absorption coefficient to below0.02 cm−1 from wavelengths 230–500 nm. Initialexperiments showed that replicate samples had lowvariability (<5% difference), so only one replicate wasanalyzed.

Apparent fluorescence quantum yields (φ) at 350 nmwere calculated from the absorbance and fluorescencedata by applying the following equation:

ðuf ÞFDOM¼ ðIf ÞFDOMaqs;k=ðIf ÞqsaFDOM;k

h i� ðuf Þqsh i

ð3Þ

where (φf)FDOM is the apparent fluorescence quantumyield of the sample at 350 nm, (If)FDOM and (If)qs are theintegrated fluorescence intensities of the sample andquinine sulfate reference solution, respectively, betweenthe excitation wavelength and 650 nm, and αFDOM andαqs,λ are the absorption coefficients of the sample andquinine sulfate solution at wavelength λ. A solution of40 ppt quinine sulfate in 0.1 N H2SO4 was used as areference solution where (φf)qs is the fluorescencequantum yield of quinine sulfate (0.51) (Velapoldi andMielenz, 1980).

In order to determine whether there was a drift inabsorption spectra or EEMs due to sample storage,


selected triplicate samples were analyzed 1 day and1 week after collection. There was less than 1%difference in the absorption at 350 nm and 2–5%difference in the EEM integrations after 1 week of darkstorage at 4 °C (data not shown). All samples wereanalyzed within 1 week of collection.

3. Results

3.1. Microbially-mediated CDOM changes

Starting values for absorption coefficients at 350 nm(α350) in inoculated treatments were 0.34±0.02 m−1 inASW and 8.22±0.16 m−1 in SW (Table 1). In the100 μM treatments, initial α350 was elevated only whennitrite or tryptophan was added to ASW. During theexperiments, treatments without microbes presentshowed little to no change in α350 (CDOM; Fig. 1).Treatments with microbes but without added N showeda decrease in CDOM absorbance (Fig. 1a,b). However,when microbes and dissolved nitrogen compounds wereincubated together, significant increases in CDOM weremeasured (Fig. 1c–f ).

When CDOM absorption is expressed as net changesabove no-N controls (i.e., after subtraction of CDOMabsorbance in control incubations with microbes butwithout added N), a net increase was found for allnitrogen compounds tested in this study (Fig. 2),although not all time points were significantly different

Table 1Initial absorption coefficients at 350 nm and average spectral slope coefficie

Class Treatment α350 (m−

ASW

Controls Control (no microbes) –Aspartic acid (no microbes) 0.17±0.0Glucosamine (no microbes) 0.23±0.0Control (microbes) 0.34±0.0

Inorganic Ammonia 0.26±0.0Nitrite 0.83±0.0

Amino Acids Aspartic Acid 0.29±0.0Glutamic acid 0.31±0.0Tryptophan 0.49±0.0

Amino sugars Galactosamine 0.27±0.0Glucosamine 0.24±0.0Mannosamine 0.28±0.0Muramic acid 0.36±0.0

No nitrogen Succinate 0.31±0.0

Succinate is the no-nitrogen analog of aspartic acid. Absorption coefficients ((SW) experiments. The spectral slope coefficient (S), averaged from all reabsorption coefficients from 290 to 500 nm to a non-linear equation. All daa Initial S above detection due to absorptivity of added substrate.⁎ Significantly different from the Control (microbes; p≤0.05).⁎⁎ Significantly different from the Control (microbes; p≤0.005).

from the control. For inorganic N additions, CDOMwasproduced by day 3 and was generally gone by day 7(Fig. 2a,b). A larger, rapid increase in CDOMwas foundwhen amino sugars were added to ASW (Fig. 2g–j),with much of the CDOM in these amino sugartreatments removed by day 14. The initial large burstof CDOM observed when amino sugars were added toASW did not occur in SW, but change in CDOMabsorption was equivalent in SW and ASW by day 14.CDOM formation was also found when amino acids(aspartic acid and glutamic acid) were added to bothtypes of seawater media (Fig. 2d,e). The addition oftryptophan caused rapid generation of CDOM, themajority of which remained throughout the experiments(Fig. 2f ).

Starting values for spectral slope coefficients (S) forCDOM in inoculated treatments ranged from 0.0107 to0.0281 in ASWand from 0.0166 to 0.0172 in SW (Table1). The spectral slope coefficients did not change in the100 μM treatments after 56 days of microbial processingexcept for the mannosamine treatment (in which Ssteadily increased over time) and the tryptophantreatment (in which a rapid decrease in S occurred thatwas maintained throughout incubation; Fig. 3).

In the 1 μM treatments, changes in the CDOM poolwere generally undetectable or were consistent withresults from the 100 μM treatments but at a significantlyreduced level. For example, 1 μM tryptophan additionspromoted CDOM formation (α350) in a pattern similar to

nts for microbial experiments1) S

SW ASW SW

7.69±0.01 – 0.01691 ⁎ 7.75±0.16 0.0144 0.01682 ⁎ 7.87±0.07 0.0143 0.01692 8.22±0.16 0.0107 0.01680 ⁎ 7.74±0.04 ⁎ 0.0120 0.01671 ⁎⁎ 8.26±0.07 0.0107 0.01661 7.67±0.02 ⁎ 0.0112 0.01690 8.00±0.16 0.0124 0.01684 ⁎⁎ 7.87±0.03 a a

0 7.90±0.01 0.0125 0.01681 ⁎ 7.78±0.02 ⁎ 0.0123 0.01681 8.31±0.01 0.0281 0.01721 – 0.0134 –2 7.71±0.03 ⁎ 0.0119 0.0168

α) are given in m−1 for artificial seawater (ASW) and natural seawaterplicates over the course of the experiments, is calculated by fittingta are averages±S.E.M.

Fig. 1. Change in CDOM (Δα350) for treatments with and without microbes (filled and open symbols, respectively) in artificial seawater (ASW) andnatural seawater (SW) incubations. No substrate was added to the controls (a, b); aspartic acid and glucosamine were added to 100 μM finalconcentration (c–f ). The change in CDOM (Δα350) is expressed relative to initial α350 and reported as the average±S.E.M. (n=3). Asterisks (⁎)above data points indicate a significant difference between cultures containing microbes and those not containing microbes (t-test, p<0.05).


that from 100 μM additions, but this increase was atleast one order of magnitude lower than that generatedwith 100 μM additions (data not shown).

3.2. Microbially-mediated FDOM change

Excitation–emission matrices (EEMs) of initialsamples (Fig. 4) showed little background FDOM inthe ASW medium compared to the natural SW medium.However, the trace vitamins that were added only toASW treatments had an excitation/emission peaksimilar to that of tryptophan protein-like fluorophores(T peak). The natural seawater EEM is similar to otherstaken from estuarine environments influenced byterrestrial input (Kowalczuk et al., 2003; Zepp et al.,2004) as seen by the dominant UV humic-likefluorophores (A) along with marine humic-like (M)and terrestrial humic-like (C) fluorophores. Of thenitrogen sources we examined, only tryptophan pro-duced initial EEMs different from controls, withmaximum excitation/emission occurring in the trypto-phan protein-like (T) region.

The only measurable increase in FDOM observedduring the 100 μM addition experiments was withamino sugars (Fig. 5); other N classes did not producemeasurable changes in FDOM. Increased FDOM in theamino sugar treatments was more transient in ASWcompared to SW, but in all of these treatments, the A-fluorophore region was dominant. In SW, FDOM build-up was most pronounced in the mannosamine treatment,as emission from each fluorophore region increasedlinearly over 28 days (Fig. 6). The FDOM pool did notchange measurably in the 1 μM addition treatments(data not shown). Statistical analysis of EEM differ-ences was not possible because only one replicate wasanalyzed per treatment.

Apparent fluorescent quantum yields at 350 nm(φ350) ranged from 0.92×10−2 to 1.46×10−2 in the SWtreatments. These values are in the same range as thosepreviously reported for FDOM in the Satilla Estuary(Zepp et al., 2004). A change in φ350 followingmicrobial processing of added nitrogen compounds inthe 100 μM treatments was evident only for mannosa-mine treatments, in which φ350 increased steadily over

Fig. 2. Net change in CDOM (Δα350) during microbial growth on various nitrogen sources in artificial seawater (ASW; open symbols) and naturalseawater (SW; filled symbols) incubations. Each nitrogen compound was added in 100 μM final concentration. The net change in CDOM (Δα350) isexpressed relative to the control (with microbes, without added substrate) and reported as the average±S.E.M. (n=3). Asterisks (⁎) above data pointsindicate a change in CDOM that is significantly different from the control at that time point (ANOVA, p<0.05). Note the different scale used forgalactosamine (j).


time from 1.19×10−2 to 1.46×10−2, and tryptophantreatments, in which φ350 decreased from 1.19×10−2 to0.92×10−2. Apparent fluorescent quantum yields in theASW treatments were highly variable, most likely dueto low absorbance values that were near the limit ofdetection (data not shown). No change in φ350 wasevident in 1 μM treatments.

3.3. Photochemical CDOM change

In the controls for the photochemical experiments,the starting absorption coefficient for SW was 8.89±

0.15 m−1 at 350 nm (α350; Table 2), comparable withthose in the microbial SW experiments (Table 1). Allother treatments had similar starting absorptioncoefficients.

Irradiation significantly decreased α350 for almostevery 100 μM addition treatment (Table 2). In thecontrol treatments without added nitrogen, CDOMdecreased during irradiation with an 8% loss of α350after 8 h (Table 2). Relative to the control, only onetreatment had increased photobleaching at α350 (nitrite).Only tryptophan caused significantly less photobleach-ing at α350 relative to the control (p<0.05). In fact, when

Fig. 3. Spectral slope coefficients during microbial incubation with 100 μM substrates in SW medium. S is reported as the average±S.E.M. (n=3).The control contained microbes but no added substrate. Asterisks (⁎) above data points indicate a significant difference between the treatment and thecontrol at that time point (ANOVA, p<0.05).


tryptophan was added to SW, CDOM at α350 wascreated following the 8-h irradiation and was clearlyvisible as a browning of the seawater (Table 2).

Changes in spectral slope coefficients followingirradiation were minor and followed no apparent patternfor the 100 μM treatments (Table 2). While mosttreatments did not cause a change in spectral slopecoefficient relative to the control, one treatment

Fig. 4. EEMs of initial FDOM for artificial seawater cultures (ASW) and natuwere generated by the MATLAB program described in Zepp et al. (2004). A r320–360/420–460 nm; M region maxima 290–310/370–410 nm; T region mequivalents (QSE). EEMs are from one replicate sample.

(tryptophan) caused the spectral slope to decreaserelative to the control.

Changes in the CDOM pool of 1 μM treatments weresimilar to those in controls and most 100 μM treatments.Tryptophan, the one treatment with photochemically-mediated CDOM generation, showed similar patterns ofCDOM generation with 1 μM and 100 μM addition.However, the magnitude of CDOM generation (α350)

ral seawater cultures (SW) after inoculation with microbes. EEM plotsegion excitation/emission maxima 260/400–460 nm; C region maximaaxima 230, 275/340 nm. Fluorescence was measured as quinine sulfate

Fig. 5. Increase in FDOM during microbial growth on amino sugars in artificial seawater (ASW; open symbols) and natural seawater (SW; filledsymbols) incubations. No substrate was added to the control; amino sugars were added to 100 μM final concentration. Other treatments andconcentrations that did not produce FDOM are not shown. Increase in FDOM is expressed relative to initial integrated EEM peak areas. EEMs arefrom one replicate sample. (Peak descriptions are given in the legend of Fig. 4.)

Fig. 6. EEMs showing FDOM production after 3, 17, and 28 days of microbial processing of natural seawater alone (control) and with 100 μMmannosamine amendment. EEMs are subtraction plots calculated by subtracting the initial sample fluorescence from the fluorescence at the indicatedsample day. EEMs are from one replicate sample.


Table 2Effect of irradiation on natural seawater CDOM amended with various nitrogen compounds

Class Treatment α350 (m−1) S

Initial (m−1) Change (m−1) Initial Change

Control Control a 8.89±0.15 −0.69±0.10 (−8%) b 0.0170 0.0001Inorganic Ammonia 8.65±0.01 −0.72±0.05 (−8%) b 0.0170 0.0001

Nitrite 9.44±0.02 ⁎⁎ −0.92±0.05 (−10%)b ⁎⁎ 0.0167 −0.0001Amino acids Aspartic acid 8.90±0.01 −0.76±0.05 (−9%) b 0.0171 0.0001

Glutamic acid 8.87±0.02 −0.69±0.05 (−8%) b 0.0171 0.0000Tryptophan 9.03±0.05 16.79±0.71 (190%)b ⁎⁎ 0.0567 ⁎⁎ −0.0385b ⁎⁎

Amino sugars Galactosamine 8.98±0.01 −0.71±0.04 (−8%) b 0.0170 0.0000Glucosamine 9.01±0.02 −0.62±0.07 (−7%) b 0.0170 0.0000Mannosamine 8.76±0.02 −0.63±0.05 (−7%) b 0.0171 0.0000

No nitrogen Succinate 8.57±0.01 ⁎ −0.74±0.04 (−9%) b 0.0172 0.0001

Samples were irradiated for 8.0 h under simulated solar radiation. Initial absorption coefficients (m−1) and the change in absorption coefficients due tophotobleaching (m−1 and %) are listed. S is the spectral slope coefficient; initial S and the change in S due to photobleaching are listed. Values aregiven as averages±S.E.M.a n=9; all other treatments n=3.b Significant change due to photobleaching (p<0.05).⁎ Change significantly different relative to control (p<0.05).⁎⁎ Change significantly different relative to control (p<0.001).


with 1 μM addition was at least two orders lower (datanot shown).

3.4. Photochemical FDOM change

EEMs of initial samples for the irradiation experi-ments were similar to those for the microbial experi-ments (see Fig. 4) except that the overall fluorescencewas heightened by ∼20% due to the difference inbackground FDOM in the water sample used for thisexperiment. Of the dissolved nitrogen compoundstested, only tryptophan produced initial EEMs differentfrom other added N-compounds and the control, withmaximum excitation/emission predictably falling withinthe T region. After irradiation, controls lost 18% of the

Table 3Change in FDOM after 8.0 h of simulated solar irradiation on natural seawa

Class Treatment Change in peak areas (QSE×104 (%

A C

Control Control −8.1±0.3(−18±1%)

−2.1±0.1(−24±3%)

Inorganic Ammonia −8.1 (−19%) −2.1 (−25%)Nitrite −7.1 (−16%) −1.9 (−22%)

Amino acid Aspartic Acid −9.4 (−21%) −2.3 (−26%)Glutamic Acid −7.9 (−17%) −1.9 (−22%)Tryptophan o/r (o/r) 6.8 (75%)

Amino sugars Galactosamine −8.1 (−18%) −2.0 (−23%)Glucosamine −7.0 (−16%) −1.8 (−21%)Mannosamine −7.6 (−18%) −1.9 (−23%)

No nitrogen Succinate −5.6 (−13%) −1.4 (−17%)

Peak descriptions are given in the legend of Fig. 3. Data were calculated froo/r=out of range.

fluorescence in the A region, 24% in the C region, 22%in the M region, and 4% in the T region due tophotobleaching (Table 3). In the presence of addednitrogen compounds, photobleaching generallydecreased FDOM to the same degree as the control.However, photooxidation in the presence of tryptophanincreased FDOM, particularly in the C-fluorophoreregion.

Irradiation decreased apparent fluorescent quantumyields at 350 nm (φ350) for all treatments (Table 3).Starting values for φ350 in all treatments ranged from1.05×10−2 to 1.15×10−2 and final values ranged from0.74×10−2 to 0.99×10−2 after irradiation. The largestdecrease in φ350 relative to the control occurred in thetryptophan treatments.

ter amended with nitrogen compounds

)) φ350 (10−2)

M T Initial Irradiated

−2.0±0.2(−22±4%)

−0.1±0.1(−4±5%)

1.13±0.02 0.94±0.02

−2.0 (−23%) −0.1 (−5%) 1.11 0.92−1.7 (−19%) −0.8 (−33%) 1.05 0.92−2.2 (−23%) −0.1 (−5%) 1.15 0.94−1.8 (−20%) −0.4 (−17%) 1.12 0.95o/r (o/r) o/r (o/r) 1.15 0.74

−1.8 (−20%) −0.1 (−5%) 1.14 0.96−1.6 (−18%) 0.0 (−1%) 1.10 0.95−1.8 (−20%) −0.1 (−5%) 1.10 0.92−1.3 (−15%) 0.0 (−1%) 1.10 0.99

m one replicate sample. φ350=apparent quantum yield at 350 nm.


Changes in the FDOM pool of 1 μM treatments weresimilar to those in most 100 μM treatments. Tryptophanwas the one exception, for which irradiation of 1 μMtryptophan produced results similar to the controls (lossof FDOM) while irradiation of 100 μM tryptophan infact produced FDOM.

4. Discussion

4.1. Microbes and CDOM formation

The presence of microbes was required to producenet increases in the CDOM pool during dark incubationswith added N compounds. Microbes alone (no Naddition) decreased CDOM absorption over the 2-month experiment, while N additions alone resulted inno net change in CDOM. Generally, when CDOM wasproduced, it was generated within 3 days and thenrapidly reduced. Similar trends of microbial formationand utilization of labile CDOMwere found by Nelson etal., (1998, 2004). In those studies, CDOM productionwas related to bacterial activity, with the highest CDOMformation corresponding to a shift in bacterial growthstage from log-phase to stationary-phase growth.

4.2. Comparisons of N compounds for CDOMformation

As a class, amino sugars supported the largest pulseof net CDOM production (Table 4). The new CDOMwas quite labile, mostly disappearing <7 days afterformation. Addition of non-aromatic amino acids tomicrobial cultures also stimulated CDOM production,but generally to a lesser extent than amino sugars.

Table 4Summary of CDOM and FDOM production by microbes and photooxidatio

Class Treatment Microbial

CDOM (α35

Amino acids Tryptophan Yes (r)Aspartic acid, glutamic acid Yes (l)

Amino sugars Glucosamine, galactosamine Yes (l)Mannosamine Yes (l)

Inorganic Ammonia, nitrite Trace (l)

CDOM production is defined as increase in α350. For microbial production oCDOM remained after 56 days of incubation) or labile (l, “new” CDOM waeither occurred or not. FDOM production is further classified as increase in pafluorophores were produced, they are listed in order of quantity, the first beia See Fig. 2.b See Fig. 5.c See Table 2.d See Table 3.e When 100 μM was added, the T peak was out of range, so increase co

Addition of the aromatic amino acid tryptophan resultedin the rapid formation of CDOM at wavelengths higherthan where this compound naturally absorbs (350 nmversus natural absorbance ∼280 nm). Furthermore, therapidly-produced tryptophan-derived CDOM persistedthroughout the experiment, suggesting that unlike aminosugar-derived CDOM, the majority of this new CDOMwas refractory. This is consistent with other studies thathave found biologically-produced CDOM with recalci-trant properties (Carlson, 2002; Hedges, 2002). Aroma-ticity has been positively correlated with bothhumification (Stevenson, 1994) and resistance tobiological degradation (Sun et al., 1997) in previousstudies, although it is not yet clear if tryptophan isdirectly incorporated into the CDOM. Relative totryptophan and the amino sugars, the two non-aromaticamino acids (aspartic acid and glutamic acid) producedless CDOM. Similarly, when processing DIN, microbesproduced a small amount of CDOM that was highlylabile and disappeared by the end of the experiments.

Since absorbance of DOM at a single wavelength oflight (e.g. 350 nm) only describes one fixed character-istic of CDOM, we used spectral slope coefficients (S) todescribe the full spectral dynamics of CDOM changes.Most nitrogen compounds did not affect S on the timescale of this experiment (∼2 months), indicating thatCDOM absorbance at all wavelengths between 290 and500 nm were affected (or not affected) equally bymicrobial processing. In natural seawater, S is relativelyconstant over time (Bricaud et al., 1981; Zepp andSchlotzhauer, 1981; Blough and Del Vecchio, 2002;Twardowski et al., 2004), and biological degradation ofCDOM has not been found to alter S (Moran et al.,2000), which is consistent with our results. The reason

n using natural seawater amended with various nitrogen compounds

Photochemical

0)a FDOMb CDOM (α350)

c FDOMd

Trace Yes C, T e

Trace No NoA, C–M, T No NoM, A, C, T No NoNo No No

f CDOM, production was classified as refractory (r, significant “new”s remineralized during incubation). Photochemical CDOM productionrticular fluorophores (described in the legend of Fig. 3). When multipleng the highest.

uld not be quantitatively assessed.


that two compounds (mannosamine and tryptophan)were exceptions and showed a net change in S is notclear, but may have been due to shifts in molecularweight or aromaticity of dissolved compounds broughtabout by microbial activity (Carder et al., 1989; Pagesand Gadel, 1990; Blough and Del Vecchio, 2002).

To explicitly separate the effect of the nitrogencomponent of the added compounds from the effect ofthe organic carbon component in the microbial forma-tion of CDOM, we included the nitrogen-free analog

Fig. 7. Comparison of fluorophores created during microbial processing and testuary, GA. End member EEMs were taken from Zepp et al. (2004). Mannosinitial EEM) that show fluorophore generation in 100 μM addition treatmeexcitation/emission maximum for the C–M fluorophore region.

(succinate) of one of the added compounds (asparticacid). However, little CDOM formation (α350) occurredin either treatment, as was typical for the amino acids(except tryptophan; Fig. 2).

4.3. Marine versus terrestrial fluorophore signatures

Coble et al. (1993) described two visible fluoro-phores that were designated “M” for marine humic-likefluorophores and “C” for terrestrial humic-like

hose present in marine and freshwater end members of the Satilla Riveramine and galactosamine EEMs are subtraction plots (final EEMminusnts after 28 days of microbial processing. Asterisks (⁎) indicate the


fluorophores. As of yet, the source materials for thesedifferent fluorophores are unknown, as are specificmechanisms of their formation. In our study, themicrobial community processed nearly identical sub-strates (mannosamine, galactosamine, and glucosamine)into CDOM and FDOM with very different opticalproperties and biological lability (Fig. 7). Mannosaminein SW produced fluorophores more reminiscent ofcoastal seawater (similar AQY and major excitation/emission peak in the M region; Zepp et al., 2004), whileglucosamine and galactosamine produced fluorophoresresembling riverine-derived organic matter (Zepp et al.,2004).

While low concentrations of free mannosamine havebeen found in various ocean waters, elevated levels arecharacteristic of the cytoplasm of some plankton,including Synechococcus sp. (Benner and Kaiser,2003). Planktonic sources of mannosamine in coastalwaters would have the potential to provide highconcentration pulses to surrounding waters that wouldbe available for microbial processing. Fluorescentcompounds formed during the microbial processing ofmannosamine or microbial enzymes released inresponse to mannosamine availability might be a sourceof M fluorophores. Both glucosamine and galactosa-mine are typically present in higher concentrations thanmannosamine in ocean water and the cytoplasm ofvarious plankton groups (Benner and Kaiser, 2003).However, neither of these compounds caused absor-bance at λ=280 nm to increase to the same degree as formannosamine, and fluorescence in the C region, ratherthan in the M region, dominated the FDOM for theseother amino sugars. These different CDOM and FDOMsignatures were also found when these amino sugarswere processed in ASW. The fact that compounds withidentical chemical composition but differing stereo-chemistry (i.e. variable hydroxyl group orientation)result in microbially-formed CDOM and FDOM withsuch distinctly different properties suggests that theremay be pronounced steric effects on the enzymaticprocesses that lead to CDOM formation.

4.4. Importance of background DOM during micro-bially-mediated CDOM formation

Parallel experiments in the presence and absence of anatural DOM background were conducted to examinehow existing DOM may interact with added Ncompounds during CDOM formation. In these experi-ments, α350 for natural SW initially was ∼24 timesgreater than α350 for the ASW (Table 1). With twoexceptions, equivalent net CDOM formation (α350) was

found in these experiments, regardless of backgroundCDOM (i.e., in ASW versus SW). For one group ofexceptions, amino sugar addition to ASW stimulated agreater initial burst of CDOM than that occurring withnatural background CDOM (SW). However, netincrease in CDOM was similar between ASW and SWafter 14 days. In the second exception, tryptophanadditions resulted in greater amounts of more persistentCDOM in SW than in ASW (Fig. 2). Other studies havehypothesized that complex parent molecules present inthe DOM pool can lead to refractory CDOM formation(Carlson and Ducklow, 1996; Nelson et al., 2004), butour results do not appear to support this. However, it ispossible that the very low background CDOM in theASW medium allowed small changes in absorbance tobe more easily observed relative to when significantbackground color was present. In contrast to CDOMproduction, however, FDOM production was consis-tently greater and the material produced was morebiologically resistant when background DOM waspresent (SW treatments) than when it was absent(ASW treatments; Fig. 4). We also found productionof the T-fluorophore only when a natural DOMbackground was present (Fig. 5).

4.5. Photochemical production of CDOM

Among all the nitrogen compounds we tested, onlytryptophan contributed to photochemical production ofCDOM in a coastal seawater sample (Table 4). Reitner etal. (2002) also found that tryptophan was the onlycompound out of alanine, tryptophan and bovine serumalbumin to photoreact and produce CDOM. Sincebovine serum albumin contains 3 residues of tryptophanand 21 of tyrosine per ∼600 residues (Brown, 1975;Hirayama et al., 1990), photooxidation of protein-boundaromatic amino acids may not produce the same effect asfor their free analogs. In addition, Reitner et al. (2002)found that tryptophan only produced color whenirradiated in natural water with background DOM, asdid we. This finding may be related to the fact thattryptophan is susceptible to photosensitized oxidation(by singlet molecular oxygen) in seawater (Momzikoff etal., 1983). All other treatments showed CDOM loss byphotobleaching, but this loss was enhanced significantlyby nitrite addition. Since nitrite is photoreactive(Jankowski et al., 1999), our results are not surprising.

4.6. CDOM formation in natural seawater

Of the two concentrations of added nitrogen used inour experiments, the lower concentration (1 μM)


produced results near the lower limit of detection forchange in CDOM and FDOM but were generallyconsistent with those from the higher concentration(100 μM). The lower concentration additions are morecomparable to those found in natural seawater, sinceDON typically remains below ∼10 μM (Lara et al.,1993; McCarthy et al., 1998; Vidal et al., 1999; Bronk,2002; Aminot and Kerouel, 2004) with higher concen-trations found in estuaries (22.5±17.3 μM) and coastal/continental shelf waters (9.9±8.1 μM) than in the openocean (surface: 5.8±2.0 μM; deep: 4.3±2.1; see Bronk,2002). Of the identifiable compounds within the DONpool, dissolved amino acids comprise the largestfraction (14–28% of ultrafiltered DON; McCarthy etal., 1996), with amino sugars (2–7%; Benner andKaiser, 2003) and urea (5.2±3.4%; Bronk, 2002)making smaller contributions. Dissolved free aminoacids (DFAA) can reach 30–400 nM in natural seawater(Lee and Bada, 1977; Mopper and Lindroth, 1982)compared to 14.6–30.1 nM for galactosamine andglucosamine individually (Benner and Kaiser, 2003).These concentrations increase during algal boom events,but elevated DON concentrations are typically transient(Meon and Kirchman, 2001). Higher concentrations ofnitrogen are also found inside algal and bacterial cells(Benner and Kaiser, 2003) which could provide localpulses of DON upon cell lysis.

CDOM formation is likely to be a gradual processthat results in the accumulation of measurable poolswhen formation rates exceed biological and physicalsinks. Nelson et al., (1998, 2004) have clearly shown aseasonal trend of CDOM formation and loss at theBermuda Atlantic Time series Study (BATS) station.Formation of CDOM at the sea surface coincides withthe spring bloom while accumulation at depth (60–100 m) occurs during the summer when phytoplanktonproductivity is low (Nelson et al., 1998). In the currentstudy, we found that the microbial community couldproduce CDOM from added DON compounds, includ-ing both labile components and recalcitrant componentswhich remain even after ∼2 months of incubation. Thissuggests that the balance of CDOM formation and lossin the ocean is likely to be strongly influenced by thechemical structure of the precursors that are acted uponby microbial and photochemical processes.

5. Conclusions

Overall, amino sugars and the aromatic amino acidtryptophan were most likely to enhance CDOM andFDOM formation over that in unamended seawater.Microbial formation of CDOM in the presence of N-rich

dissolved compounds was greater than photochemicalformation of CDOM in their presence, except fortryptophan which rapidly formed significant CDOMduring irradiation with simulated sunlight. The highlyvariable optical properties and biological lability ofCDOM formed from the suite of DON compounds wassurprising. Even for isomers within the same compoundclass (i.e., the chemically identical amino sugarsmannosamine, galactosamine, and glucosamine),CDOM and FDOM with distinctly different opticalcharacteristics were formed. Dissolved free amino acidsand dissolved amino sugars are typically found at lowconcentrations in coastal marine systems; however,consistent dilute pools as well as transient pulses ofhigher concentrations of these dissolved N compoundscould provide suitable conditions for the naturalformation of CDOM.

Acknowledgements

We thank Rosalynn Lee for help with sampleprocessing and Wade Sheldon for the use of hisMATLAB Fluorescence Toolbox program. Thisresearch was supported by grants from the Office ofNaval Research to MAM (N00014-03-1-0659) andRGZ (N0001403IP20075), and from the Gordon andBetty Moore Foundation to MAM. This paper has beenreviewed in accordance with the U.S. EnvironmentalProtection Agency’s peer and administrative reviewpolicies and approved for publication. Mention of tradenames or commercial products does not constitute anendorsement or recommendation for use by the U.S.EPA. This is contribution 941 to the University ofGeorgia Marine Institute.

References

Aiken, G.R., McKnight, D.M., Wershaw, R.L., MacCarthy, P. (Eds.),1985. Humic Substances in Soil, Sediment, and Water –Geochemistry, Isolation, and Characterization. Wiley, New York.

Aminot, A., Kerouel, R., 2004. Dissolved organic carbon, nitrogen andphosphorus in the N-E Atlantic and the N-W Mediterranean withparticular reference to non-refractory fractions and degradation.Deep-Sea Research: Part 1. Oceanographic Research Papers 51(12), 1975–1999.

Arrigo, K.R., Brown, C.W., 1996. Impact of chromophoric dissolvedorganic matter on UV inhibition of primary productivity in the sea.Marine Ecology. Progress Series 140 (1–3), 207–216.

Benner, R., Kaiser, K., 2003. Abundance of amino sugars andpeptidoglycan in marine particulate and dissolved organic matter.Limnology and Oceanography 48 (1), 118–128.

Blough, N.V., Del Vecchio, R., 2002. Chromophoric DOM in thecoastal environment. In: Hansell, D.A., Carlson, C.A. (Eds.),Biogeochemistry of Marine Dissolved Organic Matter. AcademicPress, Amsterdam, pp. 509–545.


Blough, N.V., Green, S.A., 1995. Spectroscopic characterization andremote sensing of nonliving organic matter. In: Zepp, R.G.,Sonntag, C. (Eds.), The Role of Nonliving Organic Matter in theEarth's Carbon Cycle. Wiley, New York, pp. 23–45.

Boss, E., Pegau, W.S., Zaneveld, J.R.V., Barnard, A.H., 2001. Spatialand temporal variability of absorption by dissolved material at acontinental shelf. Journal of Geophysical Research – Oceans 106(C5), 9499–9507.

Bricaud, A., Morel, A., Prieur, L., 1981. Absorption by dissolvedorganic matter of the sea (yellow substance) in the UV and visibledomains. Limnology and Oceanography 26 (1), 43–53.

Bronk, D.A., 2002. Dynamics of DON. In: Hansell, D.A., Carlson, C.A. (Eds.), Biogeochemistry of Marine Dissolved Organic Matter.Academic Press, Amsterdam, pp. 153–249.

Brown, J.R., 1975. Structure of bovine serum albumin. FederationProceedings 34 (3), 591–591.

Bushaw, K.L., Zepp, R.G., Tarr, M.A., Schulz-Jander, D., Bourbon-niere, R.A., Hodson, R.E.,Miller,W.L., Bronk, D.A.,Moran,M.A.,1996. Photochemical release of biologically available nitrogen fromaquatic dissolved organic matter. Nature 381 (6581), 404–407.

Carder, K.L., Steward, R.G., Harvey, G.R., Ortner, P.B., 1989. Marinehumic and fulvic-acids – their effects on remote-sensing of oceanchlorophyll. Limnology and Oceanography 34 (1), 68–81.

Carder, K.L., Hawes, S.K., Baker, K.A., Smith, R.C., Steward, R.G.,Mitchell, B.G., 1991. Reflectance model for quantifying chlor-ophyll-A in the presence of productivity degradation products.Journal of Geophysical Research – Oceans 96 (C11),20599–20611.

Carlson, C.A., 2002. Production and removal processes. In: Hansell,D.A., Carlson, C.A. (Eds.), Biogeochemistry of Marine DissolvedOrganic Matter. Academic Press, Amsterdam, pp. 91–151.

Carlson, C.A., Ducklow, H.W., 1996. Growth of bacterioplankton andconsumption of dissolved organic carbon in the Sargasso Sea.Aquatic Microbial Ecology 10 (1), 69–85.

Cauwet, G., 2002. DOM in the coastal zone. In: Hansell, D.A.,Carlson, C.A. (Eds.), Biogeochemistry of Marine DissolvedOrganic Matter. Academic Press, Amsterdam, pp. 579–609.

Coble, P.G., 1996. Characterization of marine and terrestrial DOM inseawater using excitation–emission matrix spectroscopy. MarineChemistry 51, 325–346.

Coble, P.G., Mopper, K., Schultz, C.S., 1993. Fluorescence contouringanalysis of DOC intercalibration experiment samples: a compar-ison of techniques. Marine Chemistry 41, 173–178.

Coble, P.G., Del Castillo, C.E., Avril, B., 1998. Distribution andoptical properties of CDOM in the Arabian Sea during the 1995Southwest Monsoon. Deep-Sea Research: Part 2. Topical Studiesin Oceanography 45 (10–11), 2195–2223.

Deuser, W.G., 1988. Whither organic carbon? Nature 332, 396–397.Flaig, W.J.A., 1988. Generation of model chemical precursors. In:

Frimmel, F.H., Christman, R.F. (Eds.), Humic Substances andTheir Role in the Environment. Wiley, New York, pp. 75–92.

Gao, H.Z., Zepp, R.G., 1998. Factors influencing photoreactions ofdissolved organic matter in a coastal river of the southeasternUnited States. Environmental Science and Technology 32 (19),2940–2946.

Hedges, J.I., 1988. Polymerization of humic substances in naturalenvironments. In: Frimmel, F.H., Christman, R.F. (Eds.), HumicSubstances and their Role in the Environment. Wiley, New York,pp. 45–58.

Hedges, J.I., 2002. Why dissolved organics matter. In: Hansell, D.A.,Carlson, C.A. (Eds.), Biogeochemistry of Marine DissolvedOrganic Matter. Academic Press, Amsterdam, pp. 1–33.

Hedges, J.I., Keil, R.G., Benner, R., 1997. What happens to terrestrialorganic matter in the ocean? Organic Geochemistry 27 (5–6),195–212.

Hedges, J.I., Eglinton, G., Hatcher, P.G., Kirchman, D.L., Arnosti,C., Derenne, S., Evershed, R.P., Kögel-Knabner, I., de Leeuw,J.W., Littke, R., Michaelis, W., Rullkotter, J., 2000. Themolecularly-uncharacterized component of nonliving organicmatter in natural environments. Organic Geochemistry 31 (10),945–958.

Hirayama, K., Akashi, S., Furuya, M., Fukuhara, K., 1990. Rapidconfirmation and revision of the primary structure of bovine serumalbumin by ESIMS and Frit-Fab LC/MS. Biochemical andBiophysical Research Communications 173 (2), 639–646.

Jankowski, J.J., Kieber, D.J., Mopper, K., 1999. Nitrate and nitriteultraviolet actinometers. Photochemistry and Photobiology 70 (3),319–328.

Kieber, R.J., Hydro, L.H., Seaton, P.J., 1997. Photooxidation oftriglycerides and fatty acids in seawater: implication toward theformation of marine humic substances. Limnology and Oceano-graphy 42 (6), 1454–1462.

Klapper, L., McKnight, D.M., Fulton, J.R., Blunt-Harris, E.L., Nevin,K.P., Lovley, D.R., Hatcher, P.G., 2002. Fulvic acid oxidation statedetection using fluorescence spectroscopy. Environmental Scienceand Technology 36 (14), 3170–3175.

Kowalczuk, P., Cooper, W.J., Whitehead, R.F., Durako, M.J., Sheldon,W., 2003. Characterization of CDOM in an organic-rich river andsurrounding coastal ocean in the South Atlantic Bight. AquaticSciences 65 (4), 384–401.

Lara, R.J., Hubberten, U., Kattner, G., 1993. Contribution of humicsubstances to the dissolved nitrogen pool in the Greenland Sea.Marine Chemistry 41 (4), 327–336.

Lee, C., Bada, J.L., 1977. Dissolved amino acids in Equatorial Pacific,Sargasso Sea, and Biscayne Bay. Limnology and Oceanography 22(3), 502–510.

Maillard, L.C., 1913. Formation de matieres humiques par action depolypeptides sur sucres. Comptes Rendus de l'Académie desSciences 156, 148–149.

McCarthy, M., Hedges, J., Benner, R., 1996. Major biochemicalcomposition of dissolved high molecular weight organic matter inseawater. Marine Chemistry 55 (3–4), 281–297.

McCarthy, M.D., Hedges, J.I., Benner, R., 1998. Major bacterialcontribution to marine dissolved organic nitrogen. Science 281(5374), 231–234.

Meon, B., Kirchman, D.L., 2001. Dynamics and molecular composi-tion of dissolved organic material during experimental phyto-plankton blooms. Marine Chemistry 75 (3), 185–199.

Meyers-Schulte, K.J., Hedges, J.I., 1986. Molecular evidence for aterrestrial component of organic matter dissolved in ocean water.Nature 321 (6065), 61–63.

Miller, W.L., Moran, M.A., Sheldon, W.M., Zepp, R.G., Opsahl, S.,2002. Determination of apparent quantum yield spectra for theformation of biologically labile photoproducts. Limnology andOceanography 47 (2), 343–352.

Momzikoff, A., Santus, R., Giraud, M., 1983. A study of thephotosensitizing properties of seawater. Marine Chemistry 12,1–14.

Momzikoff, A., Dallot, S., Pizay, M.D., 1992. Blue and yellowfluorescence of filtered seawater in a frontal zone (Ligurian Sea,Northwest Mediterranean-Sea). Deep-Sea Research. Part A,Oceanographic Research Papers 39 (9A), 1481–1498.

Mopper, K., Lindroth, P., 1982. Diel and depth variations in dissolvedfree amino acids and ammonium in the Baltic Sea determined by


shipboard HPLC analysis. Limnology and Oceanography 27 (2),336–347.

Moran, M.A., Zepp, R.G., 1997. Role of photoreactions in theformation of biologically labile compounds from dissolved organicmatter. Limnology and Oceanography 42 (6), 1307–1316.

Moran, M.A., Sheldon, W.M., Zepp, R.G., 2000. Carbon loss andoptical property changes during long-term photochemical andbiological degradation of estuarine dissolved organic matter.Limnology and Oceanography 45 (6), 1254–1264.

Nelson, N.B., Siegel, D.A., Michaels, A.F., 1998. Seasonal dynamicsof colored dissolved material in the Sargasso Sea. Deep-SeaResearch. Part 1. Oceanographic Research Papers 45 (6), 931–957.

Nelson, N.B., Carlson, C.A., Steinberg, D.K., 2004. Production ofchromophoric dissolved organic matter by Sargasso Sea microbes.Marine Chemistry 89 (1–4), 273–287.

Nissenbaum, A., Kaplan, I.R., 1972. Chemical and isotopic evidencefor the in-situ origin of marine humic substances. Limnology andOceanography 17 (4), 570–582.

Pages, J., Gadel, F., 1990. Dissolved organic matter and UVabsorptionin a tropical hypersaline estuary. Science of the Total Environment99 (1–2), 173–204.

Plummer, D.T., 1987. An Introduction to Practical Biochemistry.McGraw-Hill, London.

Reitner, B., Herzig, A., Herndl, G.J., 2002. Photoreactivity andbacterioplankton availability of aliphatic versus aromatic aminoacids and a protein. Aquatic Microbial Ecology 26 (3), 305–311.

Rochelle-Newall, E.J., Fisher, T.R., 2002. Production of chromophoricdissolved organic matter fluorescence in marine and estuarineenvironments: an investigation into the role of phytoplankton.Marine Chemistry 77 (1), 7–21.

Stevenson, F.J., 1994. Humus Chemistry: Genesis, Composition,Reactions. Wiley, New York.

Sun, L., Perdue, E.M., Meyer, J.L., Weis, J., 1997. Use of elementalcomposition to predict bioavailability of dissolved organic matterin a Georgia river. Limnology and Oceanography 42 (4), 714–721.

Thurman, E.M., 1985. Organic Geochemistry of Natural Waters.Kluwer Academic, Dordrecht.

Tissot, B.P., Welte, D.H., 1978. Petroleum Formation and Occurrence.Springer-Verlag, Berlin.

Twardowski, M.S., Boss, E., Sullivan, J.M., Donaghay, P.L., 2004.Modeling the spectral shape of absorption by chromophoricdissolved organic matter. Marine Chemistry 89 (1–4), 69–88.

Uher, G., Andreae, M.O., 1997. Photochemical production of carbonylsulfide in North Sea water: a process study. Limnology andOceanography 42 (3), 432–442.

Vähätalo, A.V., Zepp, R.G., 2005. Photochemical mineralization ofdissolved organic nitrogen to ammonium in the Baltic Sea.Environmental Science and Technology 39 (18), 6985–6992.

Velapoldi, R.A., Mielenz, K.D., 1980. A Fluorescence StandardReference Material: Quinine Sulfate dihydrate. National Bureau ofStandards (U.S.) Special Publication, pp. 260–264.

Vidal, M., Duarte, C.M., Agusti, S., 1999. Dissolved organic nitrogenand phosphorus pools and fluxes in the central Atlantic Ocean.Limnology and Oceanography 44 (1), 106–115.

Xie, H.X., Moore, R.M., Miller, W.L., 1998. Photochemical produc-tion of carbon disulphide in seawater. Journal of GeophysicalResearch – Oceans 103 (C3), 5635–5644.

Zepp, R.G., Schlotzhauer, P.F., 1981. Comparison of photochemicalbehavior of various humic substances in water: III. Spectroscopicproperties of humic substances. Chemosphere 10 (5), 479–486.

Zepp, R.G., Sheldon, W.M., Moran, M.A., 2004. Dissolved organicfluorophores in southeastern US coastal waters: correction methodfor eliminating Rayleigh and Raman scattering peaks in excita-tion–emission matrices. Marine Chemistry 89 (1–4), 15–36.

the role of nitrogen in chromophoric and fluorescent dissolved organic matter formation

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