mangrove tannins in aquatic ecosystems

8/13/2019 Mangrove Tannins in Aquatic Ecosystems

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Mangrove tannins in aquatic ecosystems: Their fate and possible influence on dissolvedorganic carbon and nitrogen cycling

Nagamitsu Maie, 1 Oliva Pisani, and Rudolf Jaffe

Southeast Environmental Research Center and Department of Chemistry and Biochemistry, Florida InternationalUniversity, Miami, Florida 33199

AbstractWe describe the fate of mangrove leaf tannins in aquatic ecosystems and their possible influence on dissolved

organic nitrogen (DON) cycling. Tannins were extracted and purified from senescent yellow leaves of the redmangrove ( Rhizophora mangle ) and used for a series of model experiments to investigate their physical andchemical reactivity in natural environments. Physical processes investigated included aggregation, adsorption toorganic matter-rich sediments, and co-aggregation with DON in natural waters. Chemical reactions includedstructural change, which was determined by excitationemission matrix fluorescence spectra, and the release of proteins from tanninprotein complexes under solar-simulated light exposure. A large portion of tannins can bephysically eliminated from aquatic environments by precipitation in saline water and also by binding tosediments. A portion of DON in natural water can coprecipitate with tannins, indicating that mangrove swampscan influence DON cycling in estuarine environments. The chemical reactivity of tannins in natural waters wasalso very high, with a half-life of less than 1 d. Proteins were released gradually from tanninprotein complexesincubated under light conditions but not under dark conditions, indicating a potentially buffering role of tannin protein complexes on DON recycling in mangrove estuaries. Although tannins are not detected at a significantlevel in natural waters, they play an important ecological role by preserving nitrogen and buffering its cycling inestuarine ecosystems through the prevention of rapid DON export/loss from mangrove fringe areas and/or fromrapid microbial mineralization.

Mangrove forests represent an important ecosystem intropical and subtropical coastal fringes and cover approx-imately 1.7 3 105 km 2 worldwide (Valiela et al. 2001). Thebiogeochemical, ecological, and economical importance of the mangrove ecosystem has been recognized recently(Valiela et al. 2001). For example, mangroves play an

important role in global carbon cycling by acting as a sinkof CO 2 (Ong 1993) and also as a significant source of dissolved organic matter (DOM) to the world oceans(Dittmar et al. 2006). While both aboveground andbelowground productivity in mangrove forests contributeto the high organic matter content in mangrove soils(Comley and McGuinness 2005), litterfall plays a crucialrole in the nutrient cycling of mangrove forests (Odum andHeald 1975; Twilley et al. 1986). Indeed, litterfall, which

consists mainly of leaves, represents about one third of primary production in mangrove forests (Robertson et al.1992). Under submerged conditions the abscised leavesrelease substantial amounts of DOM containing sugars,proteins, and polyphenols, as well as inorganic nutrients, tothe surrounding aquatic environment within a relativelyshort time period (Benner et al. 1990; Maie et al. 2006 a).Sugars and proteins are generally believed to be susceptibleto microbial degradation and thus can be quickly in-corporated into food webs (Gremm and Kaplan 1998;Weiss and Simon 1999). However, tannins, a class of polyphenols, are known to suppress microbial activity(Kuiters 1990; Hattenschwiler and Vitousek 2000; Kraus etal. 2003), and as such can affect biogeochemical cycling inthese ecosystems.

Tannins are major constituents of vascular plants, alongwith cellulose, hemicellulose, and lignins (De Leeuw andLargeau 1993), and sometimes comprise more than 20 % of the dry weight of plant materials. Their concentrationvaries among vegetation types (Hernes and Hedges 2004),

organs (Preston 1999), growing stages (Lin et al. 2006), andenvironmental conditions (Northup et al. 1998). Theconcentration of tannins can exceed that of lignins in softtissues, such as foliage, flowers, and fine roots (Kraus et al.2003).

Tannin polymers are mostly water-soluble and highlyreactive, and their ecological effect in soil sciences has beenwidely investigated (Kuiters 1990; Kraus et al. 2003).Among these ecological effects is their protein-bindingability, influencing nitrogen (N) dynamics in ecosystems(Northup et al. 1998; Schimel et al. 1998; Preston 1999).Tannins form microbially recalcitrant complexes withproteins; they also deactivate exoenzymes, preventing them

1 To whom correspondence should be addressed. Presentaddress: Department of Environmental BioScience, School of Veterinary Medicine, Kitasato University, Towada, Aomori, 034-8628 Japan ([email protected]).

AcknowledgmentsThe authors thank T. Dittmar, Florida State University, and V.

H. Rivera-Monroy, Louisiana State University, whose valuableadvice and suggestions improved this manuscript. Two anony-mous reviewers and the associate editor, E. A. Canuel, areacknowledged for providing constructive criticism on the originalmanuscript.

Funding for this research was provided by the National ScienceFoundation under the FCE LTER Program (DEB-9910514) andby NOAA (Florida Bay Program) to R. Jaffe. O. Pisani thanksNOAA for an ESRIP Internship and the SERC Endowment fora George Barley Everglades Research Grant for Undergraduatesfor financial support.

This is SERC Contribution number 357.

Limnol. Oceanogr., 53(1), 2008, 160171E 2008, by the American Society of Limnology and Oceanography, Inc.

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from rapid microbial mineralization (Baldwin et al. 1983).As a result, their presence can affect the efficiency and therate of nutrient cycling (Schimel et al. 1998; Bradley et al.2000).

Although the ecological importance of tannins has beenwidely recognized in the field of soil science and plantnutrition, there is no information on their fate in aquaticenvironments. DOM freshly leached from wetland plantbiomass contains significant amounts of dissolved organicnitrogen (DON) (Benner et al. 1990; Davis et al. 2003),which is thought to play a significant role in biogeochemicalprocesses in aquatic ecosystems. Since mangroves, especiallythose involving the Rhizophoraceae family, contain a largeamount of tannins (Basak et al. 1999; Hernes and Hedges2004), it is expected that tannins leached from mangroveleaves may sequester proteins in aquatic ecosystems. Pre-vious studies from our laboratory indicate that (1) mangrove(Rhizophora mangle ) leachate contains a large amount of polyphenols (Maie et al. 2006 a); (2) tannin concentrationsare quite low in natural waters, even in areas surrounded bymangrove forests (Maie et al. 2006 a); and (3) leachates frommangrove leaves are photosensitive (Scully et al. 2004). Thisinformation indicates that mangrove tannins are likely

highly reactive and may undergo both physical and chemicaltransformations after their release into natural waters. Theseprocesses may affect N cycling through the formation of stable tanninprotein complexes. To address this hypothesis,we proposed a conceptual model for the biogeochemicalprocessing of mangrove tannins in coastal ecosystems(Fig. 1). Each process depicted in the figure was examinedby a series of simple model experiments, as described below.

Aggregation of tannins (H1)

Tannin molecules have vicinal hydroxyl groups, whichcan act as chelates with metals (Kraus et al. 2003). Since

natural waters, especially saline waters, contain a highconcentration of metals and salts, tannins may be pre-cipitated out by aggregation (intermolecular associations)through interactions with polyvalent cations in coastal andestuarine zones. In this experiment, we investigate theaggregation/precipitation behavior of tannins along a salin-ity gradient.

Sorption of tannins to sediments (H2)

Tannins have a strong affinity to soils (Schofield et al.1998; Bradley et al. 2000; Kaal et al. 2005). As such, tanninsin natural waters may be sequestered by sorption onsuspended solids and/or underlying peat soils, even underflooded conditions. In this experiment, sorption of tanninson a mangrove peat soil was examined.

Sequestration of DON in natural waters bytannins (H3)

Tannins form insoluble complexes with proteins, whichare resistant to microbial degradation (reviewed by Krauset al. 2003). Although no such studies have been conducted

in aquatic ecosystems, tannins could potentially sequesterprotein-like DON in mangrove estuaries and significantlyreduce protein export into coastal regions. In thisexperiment, we examined the formation of insolublecomplexes between tannins and DON in natural waters.

Alteration of tannin structure (H4)

Tannins are chemically reactive (Kraus et al. 2003; Maieet al. 2003; Nierop et al. 2006); as such, tannins mighttransform quickly in aqueous environments. Furthermore,since tannins have strong light absorptive properties,photochemical reactivity is expected (Forest et al. 2004).

Fig. 1. Conceptual diagram of dynamics of tannins in aquatic environments. The hypothesisof each process, denoted in parentheses, was examined in the corresponding section.

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This experiment focused on the structural alteration of tannins in aquatic environments and the influence of lighton its reactivity.

Dissociation of tanninprotein complexes (H5)Tanninprotein complexes have been shown to be

recalcitrant to microbial degradation (Kraus et al. 2003).However, Maie et al. (2003) showed that the molecularstructure of tannins was modified during degradation of foliage, accompanied by a decrease in the protein bindingcapacity. For this reason, it could be expected that biolabileproteins can be re-released into the water column throughthe diagenetic alteration of insoluble tanninprotein com-plexes. This experiment focused on the breakdown of tanninprotein complexes during (photo)chemical alter-ation.

Materials and methodsIsolation and characterization of tannins Tannins were

extracted and purified from nearly senescent, yellow-colored leaves of red mangroves ( Rhizophora mangle ),according to the method of Maie et al. (2003). Briefly,freeze-dried mangrove leaves were soxhlet-extracted withhexane. The tannins were isolated from the solid residue byan acetone/water solution (70 : 30), to which 0.1 % (w/v)ascorbic acid was added. The crude extract was firstcleaned by several washings with dichloromethane andethyl acetate and was then purified by column chromatog-raphy on Sephadex TM LH-20 (Amersham PharmaciaBiotech AB). The purified tannins were freeze-dried and

powdered. A tannin stock solution (2,000 mg L2 1

) wasprepared fresh on the day of the experiment by dissolvingpowder tannins in Milli-Q H (Millipore) water. Tanninsolutions with concentrations of 20 mg L 2 1 or 10.6 mgcarbon (C) L 2 1 were used in most of the experiments. Thisconcentration may be somewhat higher than that in thenatural environment but was intended to facilitate thedetection of tannins.

The purity and chemical characteristics of the tanninswere confirmed by solution 13 C nuclear magnetic resonance(NMR) spectroscopy. The NMR spectrum was obtained at100.62 MHz on a Bruker AM400 spectrometer using a 5-mm probe. Purified tannin power was dissolved in a D 2 O/

acetone-d 6 (1 : 1) mixture. A standard pulse sequence withinverse gated decoupling using a 30 u excitation pulse wasapplied to obtain a quantitative spectrum (Maie et al. 2003).

Natural water and peat samples Coastal wetland sur-face-water samples werecollected from study sites establishedby the Florida Coastal Everglades Long Term EcologicalResearch (FCE-LTER) program (http://fcelter.fiu.edu/) (Ta-ble 1). A low dissolved organic carbon (DOC)concentrationmarine water sample was collected from surface Gulf Streamwater flowing offshore of the Florida Keys. Natural watersamples were filtered through GF/F glass-fiber filters andstored at 4 u C until use. A mangrove peat soil sample wascollected from site SRS-4 (25 u 419N, 80 u 969W), one of theFCE-LTER stations, where the dominant vegetation ismangrove (mixture of R. mangle , Conocarpus erectus, andLaguncularia racemosa ). The soil sample was wet-sievedthrough a 0.5-mm sieve to remove large particles.

Filtration of water samples Precombusted (450 u C for4 h) GF/F glass-fiber filters (nominal pore size, 0.7 mm;Whatman International Ltd.) were used for the filtration of water samples. Although the pore size of the filter was notsmall enough to entirely eliminate all bacteria, the filter waschosen to avoid DOC contamination from the filter andadsorption of tannins onto the filter.

DOC and total dissolved N (TDN) analysis DOCconcentrations were analyzed using a high-temperaturecatalytic combustion method on a Shimadzu TOC-V totalorganic carbon (TOC) analyzer. Samples were acidified(pH , 2) with concentrated HCl in a built-in syringe of the

TOC analyzer and were purged for 5 min with N 2 toremove inorganic C just before the injection. The practicalquantitation limit (PQL) of the measurement was 0.27 mgL2 1 . TDN was measured on an ANTEK 9000 NitrogenAnalyzer using a previously reported modification to thestandard method (Frankovich and Jones 1998). The PQLof the measurement was 0.04 mg L 2 1 .

Colorimetric analysis of total phenols Total phenolconcentration, a proxy for tannin concentration, wasanalyzed using the FolinCiocalteu method (Waterman andMole 1994). A 0.25-mL aliquot was pipetted into a test tubeand the volume was adjusted to 5 mL with Milli-Q water.

Table 1. Characteristics of natural water samples collected from the Florida Coastal Everglades.*

Location EcosystemDOC

(mg C L 2 1 )DON

(mg N L 2 1 )TP

(mg P L2 1 ) Salinity Dominant vegetation Location

T2 Freshwatermarsh

6.8 0.45 3.1 0 Sawgrass ( Cladium jamaicense );spikerush ( Eleocharis spp.);periphyton

25 u 249N,80 u 369W

T6 Mangroveswamp

7.7 0.46 3.7 0.3 Mangrove ( Rhizophora mangle ) 25u 139N,80 u 399W

T10 Estuarine 7.9 0.61 8.1 32 Seagrass; turtle grass(Thalassia testudinum )

25 u 019N,80 u 419W

* DOC, dissolved organic carbon; DON, dissolved organic nitrogen; TP, total phosphorus.

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within the FCE-LTER (T2, T6, and T10) in a 150-mL glassbeaker (inner diameter, 50 mm) covered with a quartz plateor in an amber high density polyethylene (HDPE) Nalgenebottle and incubated under light and dark conditions,respectively, for designated periods (0, 0.5, 1, 2, 4, and7 d). The light incubation was conducted under solar-simulated light produced by a Suntest XLS + solar simulator(Atlas Material Testing Technology LLC) set at 765 W m 2 2(about 1.2 times of solar noon in Miami, 25 u 459N, 80 u 229W).Thus, a total dosing of simulated light for a continuous 7 dwas estimated to correspond to that of about 25 d of sunlightin low latitudes, assuming a daytime of 8 h. After in-cubation, the water samples were filtered, and the DOC andtotal phenol concentrations and EEM fluorescence spectrawere determined. An additional experiment was conductedusing Milli-Q water and Gulf Stream water. In thisexperiment, the incubation period under dark conditionswas extended to 28 d to betterunderstand the decomposition

of tannins. The incubations were conducted in duplicate.

Dissociation of tanninprotein complexes (H5)

The experimental design was the same as for H4 exceptthat we spiked 0.3 mL of a 2,000 mg L 2 1 protein stocksolution (bovine serum albumin; BSA) to produce tannin protein complexes. Gulf Stream water was used in thisexperiment. BSA was used in this experiment since thisprotein is commonly used to estimate the protein-bindingability of tannins (e.g., Kraus et al. 2003). The structuralvariation between BSA and mangrove-leached proteinmight influence the binding strength of proteintannin

complexes and thereby the dissociation rate of protein fromtanninprotein complexes. However, our objective in thisexperiment was to obtain a general idea of the influence of sunlight exposure on the stability of tanninproteincomplexes. Incubation time was set for longer periods (upto 28 d) for dark incubation. DOC and TDN concentra-tions in the filtrate were measured. The experiment wasconducted in triplicate.

Results

Structural characteristics of the purified tannins The Cand N concentrations of tannins used in a series of experiments were 53.0 % and 0.27 % , respectively. Thesolution 13 C NMR spectrum of the tannins (Fig. 2; withassignment of chemical shifts by Czochanska et al. [1980])was typical of pure condensed tannins composed of flavan-3-ol units. The average number of eight monomer units wasestimated from the ratio of C3 extending units (6768 ppm)and C3 terminal units (7273 ppm). The ratio of procyan-idin (PC) to prodelphinidin (PD) in tannin polymer, whichdiffer in the oxygenation pattern of the B ring, waselucidated to 90 : 10 from the relative signal intensities at145 ppm (C3 9 and C4 9 of PC) and 146 ppm (C3 9 and C5 9 of PD) (Czochanska et al. 1980).

Aggregation of tannins (H1) The dissolved tanninconcentration decreased with the increase in salinity, andat salinities above 15 % , about 75 % of mixed tannins wereeliminated from the aqueous phase (Fig. 3). Therefore, weexpect that a major portion of tannins can be eliminatedwithin a day through the physical aggregation processunder high-saline water conditions.

Sorption of tannins to sediments (H2) DOC concentra-tions increased proportionally to the amount of tanninsadded for the treatment without peat (Fig. 4). However, forthe treatment with peat, no appreciable increase was

Fig. 2. Solution 13 C NMR spectrum of purified tannins usedin a series of experiments with assignment of chemical shifts.R 5 H, procyanidin (PC); R 5 OH, prodelphinidin (PD).

Fig. 3. Self-aggregation of tannins in artificial seawater withdifferent salinity (H1). Tannin concentrations were measured as

DOC. Symbol and error bar are the average and range of duplicates, respectively.

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observed in the DOC concentration for solutions withtannin concentrations below 20 mg L 2 1 , and more than80% of the added tannins were adsorbed at 80 mg L 2 1 . Thesorption of tannins to peat was fast, occurring in less than0.5 h after mixing (data not shown).

Coprecipitation of tannins with DON in natural water (H3) Although the DOC concentrations increasedas the amount of spiked tannins increased, the rate waslower than that estimated from the amount of tannins

added, indicating the precipitation of a portion of theadded tannins (Fig. 5a). A higher precipitation rate forT10, compared with T2 and T6, waters was attributed totheir high salinity ( see H1 ). While DOC data did not provethe occurrence of the coprecipitation of tannins with DOMin natural waters, TDN data (Fig. 5b,c) demonstratedclearly that the addition of tannins at a level of 20 mg L 2 1sequestered TDN in natural water by 0.050.08 mg N L 2 1 ,or 817 % .

Structural alteration of tannins (H4) DOC concentra-tions fluctuated but did not show any consistent trend forT2 and T6 under dark incubations over a 7-d period(Fig. 6). In contrast, they decreased to 4664 % of 0-

d concentrations, followed by a slight increase under lightconditions. A quick decrease in the DOC concentration byca. 40 % was observed under dark and light conditions forT10 as a result of aggregation ( see H1 ), which was followedby an increase under light conditions.

The ratio of absorption at 300 nm and 350 nm (A 300 /A 350 ) decreased throughout the dark incubation, while itfirst decreased and then increased for the light incubation(Fig. 7). The fluorescence emission peak intensity (Ex/Em5 280/314 nm), which was used as a proxy for nonalteredtannins, decreased soon after incubation for both dark andlight exposure (Fig. 8). The decrease was faster under lightexposure than under dark exposure for all of the water

Fig. 4. Adsorption of tannins on sediment (H2): open circlesare for treatments without peat (reference), and open squares are

for those with peat. Tannin concentrations were measured asDOC. Symbol and error bar are the average and range of duplicates, respectively.

Fig. 5. Sequestration of dissolved organic matter in naturalwater by tannins (H3): (a) self-/co-aggregation of tanninsmeasured by DOC; (b) DON sequestration by tannins shown inmg N L 2 1 ; (c) DON sequestration by tannins shown as % oforiginal DON. The circle, triangle, and square symbols refer to T2(freshwater marsh), T6 (mangrove swamp), and T10 (estuarine),respectively. Symbols will be the same in subsequent figures. Linesin figures represent theoretical values with no sequestration. Sincetannins contained trace amount of N, DON sequestration wascalculated by subtracting the measured value from the expectedvalue.



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samples. These results indicate that the decrease of tanninsignal is not only due to physical removal of tannins butalso to the chemical alteration of the tannin structure.

When tannins were incubated in Gulf Stream water, theDOC concentration decreased soon after the start of thedark incubation and remained at a low level until the end of

the incubation period (Fig. 9a). In contrast, the DOCconcentration increased after 0.5 d of incubation underlight conditions. Tannin concentrations detected by fluo-rescence decreased rapidly, with a half-life of less than 1 dfor both treatments (Fig. 9b). When tannins were in-cubated in Milli-Q water, the DOC concentration de-creased by , 15% and by ca. 30 % after 28 d of dark and 7 dof light incubations, respectively (Fig. 9a). The tanninconcentration detected by fluorescence decreased graduallyto 50 % after 28 d of incubation under dark conditions, butit decreased rapidly after light exposure (half-life being lessthan a day; Fig. 9b). The trend in the decrease of tanninconcentration measured by the FolinCiocalteu method

was similar to that detected by the fluorescence method,although the latter was more rapid (Fig. 9c).

The EEM fluorescence spectra of the diagenetic productsof tannins in Gulf Stream water differed for samplesincubated under dark versus light conditions (Fig. 10).Tannin freshly dissolved into Milli-Q water showed a singlepeak at Ex/Em 5 280/314 nm (Fig. 10a). However, becauseof their high reactivity in natural waters, partial change in

the chemical structure was already apparent for tanninsfreshly dissolved in Gulf Stream water (EEM spectra weremeasured within 2 h after dissolution) (Fig. 10b). TheEEM properties of tannin solutions changed further duringincubation under both dark and light conditions; a domi-nant peak was observed at Ex/Em 5 430/598 nm after 1 d,which disappeared and was replaced by peaks at , 260/470 nm and 385/479 nm after 7 d (Fig. 10c,e). The positionof the latter peak shifted slightly to a shorter wavelength(Ex/Em 5 370/470 nm) after 28 d (Fig. 10g). The majorpeaks of tannin solution incubated under light wereobserved at Ex/Em 5 , 260/468 nm, 305/461 nm, and365/469 nm after 1 d of incubation (Fig. 10d). The position

Fig. 6. Change in the DOC concentrations of natural waterthat was mixed with tannins and incubated under (a) dark and (b)light conditions (H4). The circle, triangle, and square symbolsrefer to T2 (freshwater marsh), T6 (mangrove swamp), and T10(estuarine), respectively. The symbols and error bars represent theaverage of duplicate analyses and the range, respectively.

Fig. 7. Change in the A 300 /A 350 values of natural water that

was mixed with tannins and incubated under (a) dark and (b) lightconditions (H4). The circle, triangle, and square symbols refer toT2 (freshwater marsh), T6 (mangrove swamp), and T10 (estua-rine), respectively. The symbols and error bars represent theaverage of duplicate analyses and the range, respectively.

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of the latter two peaks shifted to a shorter wavelengthduring incubation, and the intensity increased and thendecreased after 2 d and 7 d, respectively (Fig. 10f,h).

Dissociation of tanninprotein complexes (H5) Thedissolved THAA concentration was lower for treatments

with tannin than for those without, by 45%

a t 0 d(Fig. 11a,b). The dissolved THAA concentration decreasedfor 14 d and thereafter remained at a low level under darkconditions. The gradual decrease of THAA concentrationduring the incubation period was ascribed to the microbialmineralization of dissolved THAA remaining in thesolution. Dissolved THAA concentration did not increaseduring 28 d of dark incubation for treatments with tannin,indicating the refractory nature of tanninprotein com-plexes. In contrast, the dissolved THAA concentrationstarted to increase consistently after 1 d of incubationunder light exposure (Fig. 11). After 7 d of exposure, theTHAA concentration for treatments with tannins wassimilar to that without tannins.

Discussion

A series of model experiments revealed that bothphysical and chemical processes control the fate of tanninsin aquatic environments. Physical processes include aggre-gation and adsorption on particulate/peat/sediment (H1and H2). Aggregation was prominent under high salinity(H1), indicating that rain events, estuarine salinity gradi-ents, tidal conditions, and seasonal variations in freshwaterloading can influence this process. Under low salinity,adsorption of tannins on sediment will be an importantprocess (H2). This is likely to be conditioned by pH and

Fig. 8. Change in the tannin concentration in natural waterincubated under dark and light conditions (H4). Tanninconcentration was detected by a fluorescence emission intensity

at Ex/Em 5 280/314 nm. The fluorescence intensity of a corre-sponding water sample incubated without tannins for the sameperiod was subtracted as a blank. Closed and open symbols arefor treatments incubated under dark and light conditions,respectively. The symbols and error bars represent the averageof duplicate analyses and the range, respectively.

Fig. 9. Change in the (a) DOC and (b) tannin concentrations

incubated in Milli-Q or Gulf Stream water under dark and lightconditions (H4). Tannin concentration was detected by a fluores-cence emission intensity at Ex/Em 5 280/314 nm. The DOCconcentration or fluorescence intensity of corresponding watersamples incubated without tannins for the same period wassubtracted as a blank. Circle and triangle symbols refer to tanninsincubated in Milli-Q and Gulf Stream waters, respectively. Closedand open symbols refer to tannins incubated under dark and lightconditions, respectively. The symbols and error bars represent theaverage of duplicate analyses and the range, respectively.



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Fig. 10. Change in the EEM spectra of tannin solution incubated under dark and lightconditions (H4). (a) 0 d in Milli-Q water (MQ); (b) 0 d in Gulf Stream water (GS); (c, e, g) 1 d,7 d, and 28 d after incubation in Gulf Stream water under dark conditions (GSD), respectively;(d, f, h) 1 d, 2 d, and 7 d after incubation in Gulf Stream water under light conditions (GSL),respectively. Background EEM spectra of GS water, which was incubated for the same periodwithout tannins, were subtracted from each EEM spectra.

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peat/sediment characteristics, such as quality and quantityof organic matter (Kaiser 2003) and clay minerals,including sesquioxides (Varadachari et al. 1994; Kaal etal. 2005), grain size, and surface area (Kiem and Kogel Knabner 2002).

Tannins were also found to be chemically reactive innatural waters, and they undergo a two-stage diagenesis,polymerization followed by depolymerization in aquatic

environments soon after leaching (H4). Light seems toinitially stimulate polymerization of tannins, but it bringsabout depolymerization when tannins are exposed exten-sively. This is indicated by the V-shaped, time-baseddistribution of DOC concentration and the A 300 /A 350 valuefor samples incubated in natural waters under lightconditions (Figs. 6, 7, 9). It is noteworthy that precipitatedtannins can become soluble again after a certain degree of decomposition and contribute to the DOC pool in coastalestuarine water. This bidirectional change was not observedfor tannins incubated in Milli-Q water under light. Insteadthey decreased throughout the 7-d incubation (result notshown for A 300 /A 350 ). As such, the data indicate that the

diagenetic progresses were slower in Milli-Q water and thatthe polymerization of tannins seems to be the dominantprocess. The difference in the reactivity of tannins innatural waters may thus result from differences in pH,trace-metal concentrations (especially transition metals;Shindo and Huang 1984), and coexisting DOM quality andquantity (Sandvik et al. 2000).

The contrasting nature (polymerization vs. depolymer-ization) in the reactivity of light-exposed DOM has beenobserved by several researchers and attributed to differ-ences in the quality of DOM (reviewed by Moran andCovert 2003; Anesio et al. 2005). Our results indicate thatthe total dosing of light and water quality will also affectthe fate of tannins. The half-life of detectable tannins innatural water, as determined by fluorescence (Ex/Em 5280/314 nm) and the FolinCiocalteu method, was lessthan 1 d, regardless of the exposure to light (Figs. 8, 9b,c).The results indicate that a major portion of tannins leachedfrom mangrove leaves is being processed (transformed/eliminated) within a day in aquatic environments throughbiotic and/or abiotic chemical reactions. Differences in thelevels of decrease of tannins detected by the two methodsare due to the difference in the mechanisms involved in thedetection (i.e., changes in the molecular orbitals forfluorescence vs. changes in reducing power for the Folin Ciocalteu method).

It is notable that the EEM spectra of diagenetic productsof tannins had fluorescence signals overlapping with thevisible humic-like peak (peak C; Ex/Em, 320360/420 460 nm) and marine humic-like peak (peak M; 290310/370410 nm) according to the classification proposed byCoble et al. (1998) (Fig. 10). The peak position of thefluorescence signal observed near the peak M was,however, shifted to a longer wavelength compared to thetypical peak M position. Peaks C and M are considered tooriginate from terrestrial and marine sources, respectively.Our results indicate that diagenetic products of tannins cancontribute to peak C, and also peak M after extensiveexposure to light, and as such may be important sources of these peaks in tropical and subtropical estuaries and coastalregions.

Mangrove roots sometimes associate with mycorrhizae(Sengupta and Chaudhuri 2002; Kothamasi et al. 2006).Despite the refractive character of tanninprotein com-plexes against microbial degradation, some mycorrhizaecan produce exoenzymes that degrade tanninproteincomplexes and utilize the released N (Bending and Read

1996; Wu et al. 2003). Therefore, mangroves might be ableto utilize the N in tanninprotein complexes through theirsymbiotic mycorrhizae.

Furthermore, insoluble tanninprotein complexes de-grade slowly under exposure to sunlight, releasing pre-viously bound proteins into the aquatic environment(Fig. 11; H5). Therefore, the tanninprotein complexesmay serve as a long-lasting source of N in such ecosystems,which may improve synchrony between supply of N and itsuptake by mangroves. This process will be quicker in openestuaries, where exposure to sunlight is more intense. Inmangrove forests, the dense canopy attenuates sunlightintensity reaching the forest floor. The degree of light

Fig. 11. Change in the THAA concentration in Gulf Streamwater mixed with proteins (and tannins) and incubated under (a)dark and (b) light conditions (H5). The circle and triangle symbolsrefer to the treatments incubated with and without tannin, respec-tively. Error bars are the standard deviation of triplicate analyses.



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Received: 4 December 2006 Accepted: 4 July 2007

Amended: 10 September 2007


mangrove tannins in aquatic ecosystems

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