Direct Analysis of Lignin Phenols in Freshwater Dissolved Organic

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Direct Analysis of Lignin Phenols in Freshwater Dissolved Organic Matter Hendrik Reuter, Julia Gensel, Marcus Elvert, and Dominik Zak Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b03729 • Publication Date (Web): 15 Nov 2017 Downloaded from http://pubs.acs.org on November 17, 2017

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Analytical Chemistry

Direct Analysis of Lignin Phenols in Freshwater Dissolved Organic Matter Hendrik Reuter,∗,†,⊥ Julia Gensel,†,‡,⊥ Marcus Elvert,¶ and Dominik Zak†,§,k †Department of Chemical Analytics and Biogeochemistry, Leibniz-Institute of Freshwater Ecology and Inland Fisheries, D-12587 Berlin, Germany ‡Department of Chemistry, Humboldt-Universität zu Berlin, D-12489 Berlin, Germany 1

¶MARUM Center for Marine Environmental Sciences & Department of Geosciences, University of Bremen, D-28359 Bremen, Germany §Department of Bioscience, University of Aarhus, DK-8600 Silkeborg, Denmark kInstitute of Landscape Ecology and Site Evaluation, University of Rostock, D-18059 Rostock, Germany ⊥Both authors contributed equally. E-mail: [email protected]

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ACS Paragon Plus Environment


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Abstract

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A novel approach for the analysis of dissolved lignin in freshwaters is presented.

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Lignin concentrations in natural waters are low and a lignin extraction is usually re-

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quired to obtain sufficient sample for analysis. In this method, extraction and dry down

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of the dissolved lignin are omitted and alkaline CuO oxidation is directly performed us-

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ing 15 mL water sample in a microwave digestion system, thus reducing the required

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amount of sample and its preparation time considerably. Low procedural blank values

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are obtained by solid phase extraction (SPE) of the oxidized lignin phenols on HLB

10

sorbents. In combination with a here presented tandem GC-MS method, this leads to

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selective and sensitive lignin phenol quantification. Method detection limits for lignin

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phenols range from 23 to 1259 ng/L, offering applications for wetland, river and lake wa-

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ters with high terrestrial dissolved organic matter inputs as well as leachates. Besides,

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negative effects of dissolved nitrate on the lignin yield are investigated. The addition of

15

EDTA before sample acidification prevents those effects and is presented as a general

16

method improvement if lignin phenols are extracted using SPE. Three natural water

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samples, a leaf leachate and two humic-rich lake waters, were analyzed by the here pre-

18

sented direct method and by the established C18 lignin extraction procedure. Results

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show a similar reproducibility of both methods but higher absolute lignin phenol yields

20

for the here described direct dissolved lignin analysis.

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Lignin is a structural component of vascular plants and, after cellulose, the second most

22

abundant biopolymer on earth. In senescent plant litter it accounts for approximately 20%

23

dry mass. Owing to its decay-resistant structure, intact or moderately altered lignin macro-

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molecules constitute a part of the organic matter in soils, sediments and dissolved organic

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matter (DOM). 1,2 One technique to analyze lignin in such samples is the alkaline CuO oxida-

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tion which releases a set of lignin-derived phenolic monomers that are subsequently separated

27

and quantified mostly using gas chromatography mass spectrometry. 3,4

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For DOM, the quantification of lignin phenols is a powerful approach to estimate the de-

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gree of terrestrial organic matter inputs to an aquatic system. 5–9 Typical lignin yields in

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freshwater DOM range from 0.24 to 3.12 mg/100 mg dissolved organic carbon (DOC). 10,11

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The upper range of dissolved lignin is usually encountered in wetland waters where organic

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matter inputs from the vegetation are high. 5,6,9 In contrast, dissolved lignin in the oceans

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constitutes a smaller part of the total DOM with yields of 0.001-0.01 mg/100 mg DOC in the

34

North Pacific and North Atlantic ocean surface waters. 12,13

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Compared to the analysis of lignin in soils or sediments, the analysis of dissolved lignin in wa-

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ter samples requires higher effort during sample preparation. DOM concentrations in aquatic

37

systems are low (about 1-100 mg/L DOC) and a concentration routine is usually needed to

38

extract enough organic substrate for CuO oxidation. 14 Today, the most widely used DOM

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concentration method is solid-phase extraction (SPE) on C18 functionalized silica sorbents

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by which lignin phenols from 0.5 to 50 L water sample are quantitatively recovered. 15,16 Other

41

concentration methods include SPE on XAD resins, reverse osmosis or the direct dry down

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by freeze-drying or rotary evaporation. 17,18

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In this study, we present a novel approach for the analysis of dissolved lignin that directly

44

uses the aqueous water sample for CuO oxidation, omitting the initial DOM concentration

45

step. We combined several method improvements reported over the last years which lead

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to low blank values and significantly improved detection limits. While still limited to wa-

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ter samples with comparatively high dissolved lignin content, this approach considerably

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reduces the sample preparation time and the amount of water sample required. This makes

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the method particularly suitable for laboratory decomposition experiments.

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Applications as well as limitations of this approach are presented. Furthermore, we inves-

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tigated the impact of DOC concentration on the lignin yield and report negative effects on

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lignin yield, if the water sample contains dissolved nitrate. The addition of EDTA to the

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reaction mixture is presented as an approach to counteract nitrate effects and as a general

54

improvement of the phenol extraction procedure.

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Experimental section

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Chemicals. All used reagents and solvents were of analytical grade, HPLC grade or LC-MS

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grade and were, unless otherwise stated, obtained from Merck KGaA (Darmstadt, Germany),

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Sigma Aldrich Co. (St. Louis, MO, USA) or VWR International GmbH (Darmstadt, Ger-

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many). Pyridine was stored over KOH and freshly distilled every day. Every acidification

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procedure mentioned was performed using 25% HCl. To avoid contamination, all glassware

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R was heated to 450◦ C for 4 h before use. Water was obtained from an arium pro Ultrapure

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Water System from Sartorius (Göttingen, Germany).

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Leaf extracts and natural water samples. Leaf leachates were prepared from senescent

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Phragmites australis leaves collected in autumn 2014 from the rewetted fen Stangenhagen,

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south-west of Berlin, Germany. 19 We only collected brown P. australis leaves that were still

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connected to the plant. Leaves were stored under dry and dark conditions until further

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usage. 5.25 g dry leaves were leached for 24h in 3.5 L 3.5 mM NaCl solution at room temper-

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ature. A second leachate was prepared using 15 g P. australis leaves from the kettle-hole mire

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Kablow-Ziegelei, situated south of Berlin, under the same leaching procedure. That leachate

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was sequentially diluted before CuO oxidation to study the effect of DOC concentration on

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lignin yield.

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Natural water samples were taken from the experimental brown-water lake Grosse Fuchs-

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kuhle (north-eastern Germany) on October 25, 2017. Since 1989 the lake is artificially divided

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into four basins with distinct catchment areas and thus contrasting DOM characteristics. 20

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In the present study we sampled two basins with different hydrochemistry: The south-west

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basin that receives high influxes of humic-rich organic matter from an adjacent Sphagnum

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mire, and the north-east basin that receives no water from the mire. 21,22 Lake water samples

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and leachates were passed through 0.2 µm PTFE membrane filters (Omnipore, Millipore)

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and stored in the dark at 4◦ C until analysis within 48 hours. Subsamples were analyzed for

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DOC concentration on a Shimadzu TOC analyzer.

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Comparative DOM extraction. In order to evaluate the performance of the direct anal-

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ysis of dissolved lignin to an established analytical routine, we analyzed three natural water

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samples directly and after isolation of the dissolved lignin using C18 cartridges after Lou-

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chouarn et al. 16 . Briefly, we extracted lignin from 500 mL water samples and used an aliquot

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of the methanolic lignin extract which corresponded to the amount of dissolved lignin used

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in the direct approach. This volume precludes sample size effects which have been reported

87

to affect intrinsic lignin parameters in former studies. 4

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Prepacked C18 cartridges (Mega-Bond Elut-C18, 10 g, 60 mL, Agilent Technologies) were

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pretreated with 100 mL methanol followed by 50 mL acidified water (pH 2). 500 mL water

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sample, acidified to pH 2, was passed through the C18 cartridges under slightly elevated

91

head-pressure. Loaded C18 cartridges were rinsed with 500 mL acidified water to remove

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salts and dried by passing a mild stream of argon through the sorbent for 10 min. Finally,

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the DOM was eluted from the cartridge into a 100 mL volumetric flask in one fraction of

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90 mL methanol and made up to volume. An aliquot (10 mL) of the methanolic DOM extract

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was used for determination of the DOC recovery. A second aliquot (3 mL), corresponding to

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DOM in 15 mL of the initial water sample, was directly transferred to the PTFE reaction

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vessel and the methanol was removed at 45◦ C under a mild stream of nitrogen. 15 mL water

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(sparged with argon for 60 min) was added to the reaction vessel in addition to the later

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described reagents for the CuO oxidation.

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Lignin Oxidation. The lignin oxidation procedure was adapted after Goñi and Mont-

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gomery 23 using a microwave digestion system (Microprep A, MLS GmbH, Germany) equipped

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with 10 PTFE reaction vessels (100 mL volume). 500 mg CuO, 150 mg (NH4 )2 Fe(SO4 )2 ·6H2 O

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and 10 mg glucose were added to each reaction vessel. The aqueous DOM sample was

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sparged with argon for 60 min to remove dissolved oxygen and 15 mL sample was transferred

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to each vessel. Finally, 50 µL internal standard solution (ethylvanilline and cinnamic acid,

106

c = 80 µg/mL in pyridine) and 1.76 mL NaOH (50%) were added and all reaction vessels

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R , Sigma Aldrich) which was thoroughly were transferred into a 520 L glovebag (Atmosbag

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flushed with argon for 5 min. After sealing and intense shaking, the vessels were placed into

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the microwave system and heated to 150◦ C within 10 min with a temperature hold time of

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90 min.

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Lignin phenol extraction. After lignin oxidation the content of each PTFE vessel was

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transferred to a 50 mL glass centrifuge tube (custom-made), centrifuged at 750 x g for 5 min

113

and the supernatant was decanted. The step was repeated after rinsing the PTFE vessel with

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5 mL 2M NaOH. In order to remove remaining traces of CuO the combined supernatants

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were centrifuged once more and transferred to a final 50 mL glass tube. 20 mg ethylenedi-

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aminetetraacetic acid (EDTA) were added to the cooled reaction mixture (ice bath) before

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slowly adding 5.50 mL HCl (25%), thereby avoiding any heating up of the solution. The pH

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of each solution was tested before adding further HCl, if required (pH > 2).

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R HLB extracLignin phenol extraction was adapted from Kaiser and Benner 4 using Oasis

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tion cartridges (60 mg, 3 mL, Waters) placed on a 12-port extraction manifold (J.T. Baker).

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The HLB cartridges were conditioned twice with 2 mL methanol and twice with 2 mL acidi-

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fied water (pH 2). Acidified samples (pH 2) were passed through the HLB cartridges under

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gravity flow and the glass centrifuge tubes were rinsed with 0.5 mL acidified water. Salts

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were removed from the HLB cartridges with two rinses of 2 mL acidified water. Residual

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water was removed from the HLB cartridges by centrifugation at 7000 x g for 5 min. At this

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stage the HLB cartridges were frozen at -20◦ C until further use.

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For elution, teflon needle liners (disposable flow control liners for VisiprepTM DL, Supelco)

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were installed into the extraction manifold to avoid possible contamination. Prepacked anhy-

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drous sodium sulfate drying cartridges (1 g, Agilent) were installed on the disposable liner,

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followed by the HLB cartridge. Lignin phenols were eluted from the HLB cartridge with

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three rinses of 1 mL dichloromethane/methyl acetate/pyridine (70/25/5, v/v/v%). Residual

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solvent was removed from the drying cartridge by applying vacuum to the extraction mani-

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fold. The sodium sulfate drying cartridge was removed and the HLB cartridge was installed

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directly on the disposable flow liner. Phenol elution from the HLB cartridge was completed

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with two rinses of 0.5 mL dry methanol. The solvents in the combined eluates were evap-

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orated under a mild stream of nitrogen at room temperature. Samples were redissolved in

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150 µL dry pyridine, transferred into a 1.5 mL sample vial and stored frozen at -20◦ C until

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further use.

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Lignin phenol quantification. Lignin phenol quantification was carried out using a

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7000C Triple Quadrupole GC/MS system from Agilent (Palo Alto, CA, USA) equipped

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with a 7890B GC oven, a multimode inlet, and an automated liquid sampler (ALS). Prior

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to analysis, lignin phenols were derivatized by transferring 5 µL sample into a 2 mL crimp

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top vial equipped with a 250 µL vial insert. 50 µL N,O-bis(trimethylsilyl)trifluoroacetamide

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(BSTFA) with 1% trimethylchlorosilane (TMCS) was added and the reagents were mixed

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by pipetting up and down about 5 times. The vials were closed with crimp caps, stored at

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75◦ C for 20 min to complete the derivatization reaction and transferred to the ALS. 1 µL

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sample was injected under splitless mode with an inlet temperature of 300◦ C. The septum

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purge flow was 3 mL/min. The purge flow to split vent was set to 100 mL/min after 3 min.

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870 µL ultra inert inlet liners with glass wool were used.

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Separation was achieved on a DB-5ms Ultra Inert capillary column (60 m length, 0.25mm

151

inner diameter, 0.25 µm film thickness) at constant flow mode (1.5 mL/min, helium). The

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start temperature of the GC oven was 50◦ C, held for 3 min, followed by a temperature ramp

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of 10◦ C/min, a final temperature of 300◦ C and a hold time of 5 min. The transfer line tem-

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perature was held at 250◦ C. On the MS side, we used the 7000C electron ionization (EI)

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ion source heated at 230◦ C and operated at 70 eV, quadrupoles held at 150◦ C, a nitrogen

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collision flow of 1.5 mL/min, and a helium quench flow of 2.25 mL/min.

157

Measurements were carried out in Multiple Reaction Monitoring (MRM) mode following

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Louchouarn et al. 15 with modifications. The chromatographic run duration was subdi-

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vided into 12 time segments corresponding to the retention times of the different lignin phe-

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nols: 16.5-18.0 min (p-hydroxybenzaldehyde, PAL), 18.0-19.1 min (p-hydroxyacetophenone,

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PON), 19.1-19.9 min (vanillin, VAL & cinammic acid, CiAD), 19.9-20.25 min (ethyl vanillin,

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EVAL), 20.25-20.95 min (p-hydroxybenzaldehyde, PAD & acetovanillon, VON), 20.95-21.75 min

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(syringaldehyde, SAL), 21.75-22.2 min (vanillic acid, VAD & acetosyringone, SON), 22.2-

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22.9 min (3,5-dihydroxy-benzoic acid, DiOHBA), 22.9-23.55 min (syringic acid, SAD), 23.55-

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24.5 min (p-coumaric acid, CAD), 24.5 min-end (ferulic acid, FAD). One quantification and

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two qualification transitions were monitored for each target compound (Table 1). The dwell

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times were set to 50 µs per transition for time segments with three monitored transitions

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and 40 µs for segments with six transitions.

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Lignin phenols were quantified using the Agilent MassHunter Workstation Software (Quan-

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titative Analysis, Version B.07.01 for QQQ). Calibration points were determined for each

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lignin phenol by spiking blanks with increasing amounts of a calibration mixture and 50 µL

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of the internal standard mixture before CuO oxidation. Concentrations were determined as

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relative response factors to the internal standards EVAL (for VAL, VON and SAL) or CiAD

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(remaining phenols). Using a 1:2 dilution pattern, 7 calibration points ranging in concentra-

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tion from 70 to 25.000 pg/µL per phenol were determined to generate quadratic calibration

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curves based on 5 selected data points and dependend on the target analyte concentration

177

in the specific data set.

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Results and discussion

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Method Development Strategy. In previously reported methods, the analysis of dis-

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solved lignin in natural water samples requires a concentration of the DOM. 16 The obtained

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dry DOM extract is redissolved in 2M NaOH which serves as solvent in the subsequent lignin

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oxidation reaction. The critical method modification presented here omits the DOM concen-

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tration. 1.76 mL 50% NaOH is directly added to 15 mL natural water sample to obtain the

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lignin-containing 2M NaOH for the oxidation reaction. Advantages of this approach include

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the use of less water sample and the avoidance of an additional sample preparation step like

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C18 SPE or direct dry down of the water sample.

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Our approach leads to considerably smaller analyte concentrations compared to previously

188

described methods. This was addressed with the use of a microwave digestion system which

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allows higher sample volumes than the often used reaction minibombs. Lignin phenols were

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extracted using polymer-based extraction cartridges 4 to obtain lower procedural blank values

191

compared to the liquid-liquid extraction technique utilizing ethyl acetate. Phenol quantifi-

192

cation was performed by tandem mass spectrometry in MRM mode which leads to higher

193

selectivity and sensitivity compared to single quadrupole instruments.

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Chromatography, MRM transitions and detection limits. A tandem GC-MS chro-

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matogram of the lake Grosse Fuchskuhle water sample is presented in Figure 1. No chromato-

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graphic separation was achieved for VON and PAD, which was not required for quantification

197

due to the high selectivity of the MRM transitions (Figure 1, inserted picture). An overview

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of the MRM method settings of the silylated lignin phenols is presented in Table 1. Tandem

199

mass spectrometry is based upon the selective filtering of a specific precursor ion in the first

200

quadrupole followed by the collision-activated dissociation of the precursor in the second

201

quadrupole and the quantification of a specific product ion in the third quadrupole. Such

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effective ion filtering results in very low signal-to-noise values and a sensitive and selective

203

quantification of the target analytes. Three transitions were measured for each analyte and

204

consistent peak ratios of the quantifier and qualifier signals were constantly monitored by the 9



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Table 1: Retention times, collision energies and m/z-values with proposed fragmentation patterns of the silylated lignin phenols.

CiAD

RTb (min) 19.45

EVAL

20.07

PAL

17.30

PON

18.55

PAD

20.37

VAL

19.35

VON

20.35

VAD

21.89

SAL

21.27

SON

22.01

SAD

23.27

DiOHBA

22.39

CAD

23.78

FAD

25.22

Compound

Precursor Ionc (m/z ) 205 [M-CH3 ]+ 205 [M-CH3 ]+ 161 [M-CH3 -CO2 ]+ 195 [M-CH3 -C2 H4 ]+ 195 [M-CH3 -C2 H4 ]+ 167 [M-CH3 -C2 H4 -CO]+ 179 [M-CH3 ]+ 151 [M-CH3 -CO]+ 151 [M-CH3 -CO]+ 208 [M]+· 208 [M]+· 193 [M-CH3 ]+ 282 [M]+· 267 [M-CH3 ]+ 267 [M-CH3 ]+ 209 [M-CH3 ]+ 209 [M-CH3 ]+ 194 [M-C2 H6 ]+· 223 [M-CH3 ]+ 223 [M-CH3 ]+ 193 [M-CH3 -C2 H6 ]+ 312 [M]+· 297 [M-CH3 ]+ 297 [M-CH3 ]+ 254 [M]+· 254 [M]+· 224 [M-C2 H6 ]+· 268 [M]+· 253 [M-CH3 ]+ 238 [M-C2 H6 ]+· 342 [M]+· 327 [M-CH3 ]+ 327 [M-CH3 ]+ 370 [M]+· 355 [M-CH3 ]+ 355 [M-CH3 ]+ 293 [M-CH3 ]+ 293 [M-CH3 ]+ 219 [M-C3 H9 SiO]+ 323 [M-CH3 ]+ 323 [M-CH3 ]+ 308 [M-C2 H6 ]+·

Product Ionc (m/z ) 161 [M-CH3 -CO2 ]+ 131 [M-CH3 -C2 H6 SiO]+ 145 [M-CH3 -CO2 -CH4 ]+ 179 [M-CH3 -C2 H4 -CH4 ]+ 167 [M-CH3 -C2 H4 -CO]+ 151 [M-CH3 -C2 H4 -CO-CH4 ]+ 151 [M-CH3 -CO]+ 95 [M-CH3 -CO-C2 H4 Si]+ 75 [M-CH3 -CO-C6 H4 ]+ 193 [M-CH3 ]+ 73 [M-C8 H7 O2 ]+ 73 [M-C8 H7 O2 ]+ 267 [M-CH3 ]+ 223 [M-CH3 -CO2 ]+ 193 [M-CH3 -C2 H6 SiO]+ 193 [M-CH3 -CH4 ]+ 165 [M-CH3 -C3 H8 ]+ 137 [M-C2 H6 -C2 H5 Si]+· 208 [M-CH3 -CH3 ]+· 193 [M-CH3 -C2 H6 ]+ 137 [M-CH3 -C2 H6 -CH3 -CHO]+ 297 [M-CH3 ]+ 282 [M-CH3 -CH3 ]+· 267 [M-CH3 -C2 H6 ]+ 239 [M-CH3 ]+ 224 [M-C2 H6 ]+· 195 [M-C2 H6 -CHO]+ 238 [M-C2 H6 ]+· 238 [M-CH3 -CH3 ]+· 195 [M-C2 H6 -C2 H3 O]+ 327 [M-CH3 ]+ 253 [M-CH3 -C2 H6 SiO]+ 223 [M-CH3 -C3 H9 SiO-CH3 ]+ 281 [M-C3 H9 SiO]+ 311 [M-CH3 -CO2 ]+ 281 [M-CH3 -C2 H6 SiO]+ 249 [M-CH3 –CO2 ]+ 219 [M-CH3 -C2 H6 SiO]+ 191 [M-C3 H9 SiO-CO]+ 293 [M-CH3 -C2 H6 ]+ 249 [M-CH3 -C2 H6 SiO]+ 293 [M-C2 H6 -CH3 ]+

b

CE (eV ) 5 17 5 8 8 11 8 8 17 8 29 20 8 11 20 20 29 29 5 17 23 8 5 14 5 17 20 14 5 40 8 23 32 23 11 17 8 17 11 14 17 5

Retention time on DB-5ms Ultra Inert column (60 m, 0.25 mm ID, 0.25 µm film thickness; Agilent Technologies). c Underlined ions highlight the quantification transitions.

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Figure 1: Tandem mass chromatogram of the lake Grosse Fuchskuhle (south-east) water sample in MRM mode (black) and in full scan mode (grey, multiplied with 0.2). The inserted picture highlights the coelution of VON and PAD and the three MRM transitions recorded for each analyte. 205

quantification software. Only PAL and PAD required manual inspection of a correct peak

206

integration in a few natural samples with low lignin concentrations. These analytes pro-

207

vided a comparably poor fragmentation pattern due to the low number of functional groups

208

present, thus more unspecific product ions had to be chosen (m/z = 75, (CH3 )2 SiOH+ and

209

m/z = 73, (CH3 )3 Si+ ).

210

Instrument detection limits (IDL) and method detection limits (MDL) were determined

211

following guidelines of the U.S. Environmental Protection Agency (EPA). 24 The IDL was

212

calculated from 7 replicate injections of the lowest calibration standard (4.2-7.4 pg/µL) as

213

IDL = tα ·σ where tα refers to the critical value of the t distribution (α = 0.01) and σ is the

214

standard deviation. The MDL was calculated analogously using 7 spiked samples (10-50 ng

215

phenol) that ran through the complete sample preparation procedure. IDLs are presented

216

in pg analyte per µL pyridine/BSTFA solution (Table 2), ranging from 0.15 to 0.60 pg/µL.

217

MDL values are presented as lignin phenol concentrations in a water sample after CuO ox-

218

idation and range from 23 to 1259 ng/L. The lowest values between 23 and 94 ng/L were

219

determined for the phenols bearing a ketone group (PON, VON and SON). These analytes

220

showed a high and reproducible extraction recovery and low blank values. MDL values allow 11



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Table 2: Instrumental detection limit (IDL) and method detection limit (MDL) of single lignin phenols using MRM by tandem mass spectrometry. Compound PAL PON PAD VAL VON VAD SAL SON SAD DiOHBA CAD FAD a b

221

222

223

224

IDLa (pg/µL) 0.37 0.24 0.28 0.40 0.15 0.39 0.29 0.28 0.55 0.24 0.43 0.60

MDLb (ng/L) 230 64.2 225 57.7 22.7 91.3 166 93.5 294 1100 1260 836

IDL refers to unsilylated analyte concentration in the injected pyridin/BSTFA mixture. MDL refers to concentration of lignin oxidation products in a water sample.

a rough estimation of the minimal DOC content in order to analyze a water sample directly for dissolved lignin. The total dissolved lignin content of a water sample is defined as the P P sum of vanillyl, syringyl and cinnamyl phenols ( 8 =V+S+C). This leads to a 8 -MDL of P 2.8 µg/L and a method quantification limit 8 -MQL of 8.5 µg/L. In a natural aquatic system

225

with terrestrial DOM sources, total dissolved lignin accounts for about 0.5 mg/100mg DOC

226

what would lead to a minimal DOC concentration of 1.7 mg/L in order to quantify lignin.

227

This estimation, however, underestimates the DOC concentration as it is solely based on

228

the MDL and neglects the natural distribution of lignin phenols released during oxidation

229

of a natural water sample. Consequently, the direct approach of dissolved lignin analysis

230

can be applied to lignin rich waters from wetlands, 18 to riverine water with high terrestrial

231

DOM concentrations, such as the Congo river, 9 as well as to soil porewater or to laboratory

232

experiments studying dissolved lignin decomposition on appropriate sample types. However,

233

waters from many rivers, such as the arctic rivers, 7 as well as estuarine and marine wa-

234

ters 12,13 generally do not contain sufficient dissolved lignin for direct analysis.

235

Lignin phenol extraction. The liberated lignin phenols are extracted from the aqueous

236

phase after CuO oxidation. Alternatively to the liquid-liquid extraction with ethyl acetate we

237

R HLB catridges after Kaiser and Benner 4 . Deviating followed the SPE approach with Oasis

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Figure 2: SPE recoveries of a lignin phenol standard mixture (0.5-1.5 µg of each phenol) extracted from 2, 10,P25 and 50 mL water. Error bars P represent standard deviations of duplicate analyses. Aldehydes = PAL + VAL + SAL, Ketones = PON + VON + SON, P Acids = PAD + VAD + SAD + CAD + FAD. 238

from the original method, the microwave digestion systems used here processes higher sample

239

volumes than the classical reaction minibombs. We therefore investigated the phenol reten-

240

tion capacity of the HLB resin by spiking equal amounts of a standard containing between

241

0.5 and 1.5 µg of each phenol into different water volumes (i.e. 2, 10, 25 and 50 mL) treated

242

with base, followed by acidification and finally extracted according to the lignin protocol.

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Syringic phenols and those bearing an acidic functional group revealed a lower retention on

244

the resin which led to considerable analyte breakthrough at higher sample volumes (Figure 2

245

and Table 5 in the Supporting Information). In particular SAD, FAD and DiOHBA showed

246

recoveries below 50% when extracted from 50 mL water volume, whereas VAD and PAD

247

were quantitatively recovered.

248

In accordance with the original lignin oxidation protocol in a microwave digestion system 23

249

we routinely used a volume of 15 mL for CuO oxidation of water samples. Recovery rates

250

for calibration standards that ran through the complete analysis ranged from 76.7 to 111.6%

251

except DiOHBA with a low recovery of 48.5% (Table 4 in the Supporting Information).

252

Loading time of the lignin phenols on the HLB sorbent was between 2-4 h. Even though

253

no residual CuO was visually detected in the samples after centrifugation, we noticed the 13



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accumulation of CuO residuals on the SPE sorbent during loading thus reducing sample flow

255

through the sorbent. Addition of EDTA to the sample before the acidification step counter-

256

acted this effect as no accumulation of CuO on the sorbent occurred. Intentionally added

257

to prevent phenol losses in water samples that contain nitrate (see below), we therefore sug-

258

gest the general use of EDTA before acidification. No negative effects of EDTA on phenol

259

extraction were observed, but the HLB cartridge loading time is reduced and catalytically

260

active copper(II) is removed at an early stage.

261

Nitrate interference. The C18 extraction of DOM from water samples not only concen-

262

trates lignin phenols, but also removes dissolved inorganic salts from the sample matrix. As

263

we omitted this step, inorganic salts remain present in the reaction mixture during CuO

264

oxidation. Thus, potential matrix effects of the latter need to be addressed which we con-

265

fidently believe have no effect on the lignin phenol analysis of aqueous DOM samples for

266

two reasons. Firstly, the method is commonly applied to dried sediments, soils or freeze-

267

dried DOM, thus to samples that similarly bear a strong inorganic matrix; 4,17 and secondly,

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(NH4 )2 Fe(SO4 )2 ·6H2 O is routinely added to the reaction mixture prior to the CuO oxidation

269

step in order to remove residual oxygen, a step that generally increases the inorganic matrix

270

considerably.

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As one exception to the otherwise low matrix effects, however, we noticed discrepancies in

272

the lignin yield of water samples that contained high amounts of dissolved nitrate. To specif-

273

ically investigate this effect, we spiked a P. australis leaf leachate with increasing amounts of

274

sodium nitrate before CuO oxidation. Results of this experiment indicate a significant drop

275

in lignin yield at nitrate concentrations of 15 mg/L nitrate-N or higher (t-test, p0.05) (Figure 4a). The

314

sample with 5 mg/L DOC, however, indicated a significant decrease in syringyl and cinnamyl

315

phenol yield (t-test, p

Direct Analysis of Lignin Phenols in Freshwater Dissolved Organic

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