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Revised black carbon assessment using benzene polycarboxylic acids
Organic Geochemistry Organic Geochemistry 36 (2005) 1299–1310 www.elsevier.com/locate/orggeochem
Revised black carbon assessment using benzene polycarboxylic acids S. Brodowski b
a,c,* ,
A. Rodionov b, L. Haumaier a, B. Glaser a, W. Amelung
c
a Institute of Soil Science and Soil Geography, University of Bayreuth, 95440 Bayreuth, Germany Institute of Soil Science and Plant Nutrition, Martin Luther University Halle-Wittenberg, 06108 Halle, Germany c Institute of Soil Science, University of Bonn, Nussallee 13, 53115 Bonn, Germany
Received 8 January 2004; accepted 31 March 2005 (returned to author for revision 3 March 2004) Available online 26 May 2005
Abstract Black carbon (BC), the ubiquitous stable product of incomplete combustion, is believed to be a potential sink for atmospheric CO2 and therefore a contributor to the EarthÕs radiative heat balance. Nevertheless, analytical procedures to measure BC are inconsistent, giving a non-systematic variation by factors of 14–571 for estimates of its content in soil. We hypothesized that the HCl used to isolate benzene polycarboxylic acids (BPCAs) as markers for BC helps form these compounds, which could then cause an overestimation of the BC content of the soil. We found that indeed up to 90% of BPCA yields may be attributed to this HCl pre-treatment. To correct this error we developed a revised method that uses BPCAs as BC markers but which allows us to eliminate any confusion in the results. This aim was achieved by digestion with 4 M trifluoroacetic acid (TFA). After oxidation with HNO3, the BPCAs were purified using a cation exchange resin and derivatized to form the trimethylsilyl derivatives. Analyses were performed using a gas chromatograph (GC) equipped with a flame ionization detector (FID); constant linearity was obtained at P7 ng BPCA injection amount and peak purity was determined using mass spectrometry (MS). The recovery of the BPCAs averaged 93.5 ± 5.1% for pure standards and 95.0 ± 3.6% for spiked charred plant material. The contribution of BPCAs from aspergillin to soil organic carbon was estimated to be negligible. No close correlation between the results obtained with the original method and our revised procedure was observed. 2005 Elsevier Ltd. All rights reserved.
1. Introduction Black carbon (BC), a product of incomplete combustion of biomass and fossil fuel, is found nearly everywhere due to widespread aeolian transport (Goldberg, 1985; Stoffyn-Egli et al., 1997). The BC particles comprise a *
Corresponding author. Tel.: +49 228 73 2194; fax: +49 228 73 2782. E-mail address: [email protected] (S. Brodowski).
wide continuum of combustion residues ranging from char and charcoal to condensates, such as soot (Jones and Chaloner, 1991; Goldberg, 1985). BC is relatively inert. Because of this, Kuhlbusch (1998), for instance, suggested that it may be a potential sink for atmospheric CO2. If so, it could well contribute to the EarthÕs radiative heat balance (Crutzen and Andreae, 1990). A number of authors have developed an interest in the presumed impact of BC on the global carbon cycle and have begun intensified research into the fate of charred organic material in various environmental compartments: the
0146-6380/$ - see front matter 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.orggeochem.2005.03.011
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atmosphere (Chang et al., 1982; Charlson and Ogren, 1982; Andreae et al., 1994; Bird, 1997), (marine) sediments (Herring, 1985; Novakov et al., 1997; Masiello and Druffel, 1998; Middelburg et al., 1999) and soils (Skjemstad et al., 1996; Schmidt et al., 1999; Schmidt and Noack, 2000; Schmidt et al., 2002). Despite growing interest in BC, both a generally accepted definition and standard analytical procedures are lacking (Schmidt and Noack, 2000; Schmidt et al., 2001). As pointed out by Schmidt et al. (2001), comparative results from six methods revealed a non-systematic variation in soil BC content by factors of 14–571. Thermal and UV-oxidation procedures have been criticized as overestimating BC content of soils and sediments (Simpson and Hatcher, 2004). To accurately quantify BC in the environment there is, therefore, a need to improve current methods. Glaser et al. (1998) proposed the use of benzene polycarboxylic acids (BPCAs) as specific markers for BC (Fig. 1). Their method involves preextraction with hot 32% HCl, oxidation of condensed aromatics to BPCAs using hot HNO3, purification and silylation of the products prior to quantification using GC with FID. Some drawbacks in the method became obvious, however, when it was applied to a large number of samples. Unacceptable analytical errors arose whenever samples with a wide range of weight and containing small amounts of BC were analyzed (Brodowski et al., unpublished data). Moreover, the test was unable to re-
veal whether the BPCAs came from artifactual formation or additional natural sources. For instance, treating organic matter or pinoresinol, a constituent of lignin (Freudenberg et al., 1965), with strong acid resulted in a formation of BPCAs (Brunow, 1965; Rudakov et al., 1986). Strong acid treatment of bacterial biomass or of a mixture of monosaccharides and amino acids produced melanoidin-like artefacts (Allard et al., 1997). Upon permanganate oxidation at least some melanoidins yield BPCAs (Ishiwatari et al., 1986). This finding led us to suspect that artefact formation in the pre-treatment step of the method might cause further methodological errors. Apart from artificial formation, BPCAs could, in principle, also be derived from melanoidins, the BPCA yields from which are unknown. We agree with those who say that the Maillard reaction (non-enzymatic browning) does not appear to contribute any significant amount of SOM formation (Hatcher, 2003) or degradation of algae (Zang et al., 2001). We therefore disagree with those who contend that the process is a possible humification source for soils and marine environments (Nissenbaum and Kaplan, 1972; Stevenson, 1982; Ertel and Hedges, 1983; Rubinsztain et al., 1984; Poirier et al., 2000). The reason for this is that the process involves a number of reactions, some of which take place at temperatures substantially warmer and pH substantially more alkaline than those commonly found in soil. While
Fig. 1. Structures of benzene polycarboxylic acids used as markers for black carbon assessment.
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
high temperatures are known to form heterocyclic N and polyaromatic C from soil organic residues (Almendros et al., 2003), the only time that high temperatures occur naturally in soil is during a wildfire or controlled burning. Maillard reactions are not sustained at any substantial scale when fires are absent. We therefore suggest that any melanoidin-derived aromatic structure formed in soils should be assigned as BC. Reports of BPCA formation from some of the melanoidins may warrant attention. Other natural products in soils, upon oxidation after treatment with strong acids, can produce BPCAs. One of these is aspergillin, a black amorphous pigment from Aspergillus niger (Lund et al., 1953) that is supposed to contribute to the aromaticity of soil humic substances (Kononova and Aleksandrova, 1959). At the latter time, aspergillin had not been analyzed using the method of Glaser et al. (1998). The main aims of the present study were (1) to eliminate the analytical problem in samples with low BC contents when analyzed using the method of Glaser et al. (1998); (2) to examine whether the HCl treatment in this method artificially forms BPCAs and, thus overestimates BC contents of soils and if so, to establish an alternative pre-treatment; (3) to determine the extent to which synthetic melanoidins and naturally occurring aspergillin yield BPCAs using this method.
2. Materials and methods 2.1. Samples We obtained BC-free materials from above-ground biomass (stems and leaves) of seven plants (Astragalus sp., Cymbaria dahurica L., Leperiza sp., Populus pseudosimonii var. patula T.Y. Sun, Secale cereale L., Setaria viridis (L.) P. Beauv., Zea mays L.); and from soilinhabiting microorganisms such as the fungi Aspergillus niger DSM B202, Penicillium citrinum DSM 62830 and the bacteria Pseudomonas putida DSM 291T and Agrobacterium tumefaciens ATCC 11095. Melanoidins were prepared by condensation of one amino acid (glycine, glutamic acid or lysine) and a sugar (D-glucose) in hot alkaline solution as described by Hedges (1978). The sugar to amino acid concentration ratios were 9:1 and 1:9 (v:v), respectively. Pseudomelanoidins were produced in a pure sugar solution without adding amino acids. For alkaline conditions, 0.167 M amino acid or sugar solutions were prepared in 0.1 M Na2CO3. The reaction was carried out by heating 200 ml of reaction solution in glass bottles to 100 C for 170 h. Thereafter, the melanoidins and pseudomelanoidins were precipitated by acidification with 1 M TFA to pH 1. After 16 h the precipitated melanoidins were centrifuged (4420g, 15 min), washed with
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deionized water, which was acidified with TFA to pH 1, again centrifuged, freeze-dried and subjected to BPCA analysis. We failed to produce a precipitate for a sugar to amino acid ratio of 1:9 (v:v). All reactions were carried out in quadruplicate to produce enough material for BC analysis. For the method evaluation, we used ground straw of Z. mays L., the fungus Penicillium citrinum DSM 62830, the two bacteria P. putida DSM 291T and A. tumefaciens ATCC 11095 and various soil samples with different properties from various climatic regimes (Table 1). The maize straw was prepared for charring by air drying and chopping to <5 cm pieces. These materials were placed in stainless steel containers closed with a cap to reduce oxygen supply. The containers were heated to a temperature of 350 C in a muffle furnace. It took 1 h to reach this temperature and heating continued thereafter for 2 h. The mass loss after heating averaged 67.6 ± 0.47% (SE; n = 12) of the initial mass. The charred maize contained 66.35% C, 3.36% H, 16.00% O, and 2.50% N. 2.2. Analytical procedures 2.2.1. Elimination of polyvalent cations, oxidation and purification For metal elimination about 0.5 g of each sample was subjected to digestion either with 2 ml of 32% HCl [4 h, 170 C; Glaser et al. (1998) original method] or with 10 ml of 4 M TFA [4 h at 105 C; revised method]. Additionally, several extractants (32% HCl, 6 M HCl, sodium dithionite, sodium oxalate, hydrofluoric acid) were tested for metal elimination at two temperatures (room temperature, 105 C) and three digestion times (16, 8 and 4 h). These alternatives were, however, unmanageable or gave irreproducible results or both (data not shown). When the sample was expected to contain more than 200 g BC per kg organic C and/or more than 100 g kg1 organic C, we reduced the sample weight to 5 mg organic C. After digestion with either acid, the residue was processed as outlined by Glaser et al. (1998) with additional modifications concerning the reduction of HNO3 sample aliquots to 62 ml, extended storage time of the derivatization solution (P24 h). In brief, samples were processed as follows. The residue was collected by filtration through a glass fibre filter (GF 6, Schleicher and Schuell, Dassel, Germany), rinsed several times with deionized water and dried at 30–40 C for at least 2 h. Thereafter, all of the residue was transferred to a glass digestion tube, 2 ml of 65% HNO3 were added and the mixture was heated to 170 C and held there for 8 h in a high pressure digestion apparatus (Schramel et al., 1980) where BC was oxidized to BPCAs (Fig. 1). Repeated oxidation of BC generally reproduced the BPCA yields with a variation of 3.5% (median) or 6.9
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Table 1 Selected climatic data and properties of soils used for method optimization [benzene polycarboxylic acids (BPCAs) determined using revised method; without conversion factor] No.
Site identification
Samples used for method evaluation 1a RUSSIA: Kursk 1b 2 RUSSIA: Frolovo 3 BRAZIL: Belterra
MAPa (mm)
MATb (C)
Depth (cm)
Organic C (%)
C/N
BPCAC (g kg1 organic C)
Haplic Chernozem
573
5.3
Haplic Kastanozem Xanthic Ferralsol
300 2050
6.0 26.5
0–10 30–40 10–25 0–10
5.64 3.45 2.29 1.44
12.0 12.3 10.4 10.3
49.3 88.5 65.2 19.8
480
8.7
490
9.1
890
8.2
0–25 0–20 54–64 0–25 0–25 0–30 0–30
2.01 1.33 0.27 1.30 1.25 1.36 1.30
11.2 9.4 6.6 15.5 15.2 8.3 8.2
63.5 53.7 58.7 65.2 63.9 20.1 20.0
Soil type (FAO, 1988)
Further samples used for method validation 5 GERMANY: Bad Lauchsta¨dt Haplic Chernozem 6a 6b 7 GERMANY: Halle Luvic Phaeozem 8 9 GERMANY: Rotthalmu¨nster Stagnic Luvisol 10 a b
MAP, mean annual precipitation. MAT, mean annual temperature.
(mean) ± 2.0% (standard error; n = 13) of the mean. The solution was poured through an ash-less cellulose filter (Schleicher and Schuell 5893; d = 70 mm) into a 10 ml volumetric flask. Deionized water was added to fill the flask. To eliminate remaining polyvalent cations, such as of Fe3+ or Al3+, the following procedure was used. To avoid degradation of the internal standard by the strong acid, an aliquot (0.5–2 ml) of the sample solution was diluted with 4 ml deionized water (the original method processed up to 5 ml aliquots). Subsequently, internal standard 1 was added (100 lg citric acid in 100 ll deionized water). The solution was then dropped on to a cation exchange resin (Dowex 50 W X 8, 200– 400 mesh, Fluka, Steinheim, Germany), which was converted to the H+ form prior to use. After complete infiltration, the BPCAs were eluted into 100 ml conical flasks with 50 ml of deionized water in portions of 10 ml. Finally, the eluates were freeze-dried for water removal. We also learned that the use of an aliquot >2 ml for further sample processing should be avoided. It sometimes resulted in significant decomposition of the citric acid due to strong acid conditions and possibly an incomplete removal of interfering cations during purification and subsequent complexation of BPCAs (data not shown). All reagents were of analytical grade and purchased from Sigma/Aldrich or Fluka. 2.2.2. Derivatization The freeze-dried BPCAs were re-dissolved in 4 · 1 ml methanol and transferred to 5 ml reactivials (Alltech GmbH, Unterhaching, Germany) with Teflon-laminated septum screw caps. One hundred microliters of
the second internal standard solution containing 100 lg of biphenyl-2,2 0 -dicarboxylic acid in methanol were added and the solvent was evaporated with a stream of air. The dried BPCAs were converted to trimethylsilyl derivatives. For this purpose, we added 100–125 ll dry pyridine and 100–125 ll N,O-bis(trimethylsilyl)-trifluoroacetamide, heated the solution at 80 C for 2 h, allowed it to cool and stored it for 24 h. During the 24 h the detector signals increased (see below) and then remained stable. We conclude that derivatization was not complete without the 24 h storage. 2.2.3. GC GC analysis was performed with a Hewlett Packard 6890 gas chromatograph (Hewlett Packard GmbH, Waldbronn, Germany) equipped with a FID and an HP-5 capillary column (30 m · 0.32 mm i.d., 0.25 lm film thickness). Helium was used as carrier gas at a constant flow of 0.8 ml min1. Both the injector and detector temperatures were 300 C. Aliquots (2 ll) of sample solution were injected at a split ratio of 30:1 into a fully deactivated inlet system with silylated liners. The temperature programme was: initial column temperature of 100 C held for 2 min followed by an increase in 20 C min1 to 240 C and held for 7 min. Subsequently, the temperature was raised at 30 C min1 to 300 C and held for an additional 10 min (total run time = 28 min). Peak identity was confirmed with full scan mass spectra obtained using a Hewlett Packard 6890 series GC coupled to a Hewlett Packard 5972A mass selective detector (Hewlett Packard GmbH, Waldbronn, Germany).
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
2.2.4. Standard series Standard series were prepared for each analysis in two- to three-fold replication using standard solutions of 5, 10, 15, 25, 50, 100 lg per vial of BPCA (hemimellitic acid, trimellitic acid, trimesic acid, pyromellitic acid, benzene pentacarboxylic acid, mellitic acid). Linearity was checked with additional standard solutions of 20, 30, 40, 60, 80, 120, and 150 lg per vial of BPCA. These were prepared by dissolving 100 lg of each BPCA in 100 ll methanol and transferring the corresponding volume of standard solution (e.g., 5 ll or 10 ll etc.) into a vial, then containing, e.g., 5 lg or 10 lg of each BPCA. The content of the internal standards was kept constant at 100 lg of citric acid and 100 lg biphenyl-2,2 0 -dicarboxylic acid per vial, both in 100 ll methanol. Subsequently, the standards were completely dried and individually derivatized as described above.
3. Results and discussion 3.1. Linearity Previous experiments using different sample amounts for analysis according to the method of Glaser et al. (1998) revealed high coefficients of variation (P50% of the mean). In these earlier experiments, however, a one-point calibration was used. Derivatizing standard mixtures of different concentrations revealed an apparent linearity of detection (r2 = 0.9964 and higher). The coefficient of regression was only slightly improved when using a second-order regression curve. However, as previously reported by Amelung and Zhang (2001) for MS analyses of amino acid enantiomers, even an r2 close to unity may blur a non-linearity at low concentrations. Indeed, when calibration curves were forced through the origin, signal response of the FID was non-linear at <7 ng injection amount of silylated BPCAs (data not shown) and this non-linearity could be reproduced. For minimization of analytical errors we recommend, however, that sample weight be adapted to give >7 ng of expected BPCA yield, if possible. 3.2. Method-induced production of benzene polycarboxylic acids When we used the Glaser et al. (1998) method to analyze charred maize straw we found BPCAs, as expected (Fig. 2a). We also unexpectedly found significant amounts of BPCAs (Fig. 2b), however, when we used the same method to analyze plant materials which we knew were free of BC. Therefore, we concluded that the original method of Glaser et al. (1998) produced BPCAs. That method involves acid digestion with hot 32% HCl, and we suspected that there was acid-catalyzed BPCA formation, leading to
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an overestimation of BC contents in environmental samples. No BPCAs were found when oxidation was carried out without the HCl pre-treatment (Fig. 2c). Recently, overestimation has also been reported for thermal and UV-oxidation procedures used to determine BC in the environment (Simpson and Hatcher, 2004). The method-induced BPCA formation was not directly proportional to the amount of plant material added to soil (data not shown) and accounted for 0.1–4.3% of plant and microbial biomass (Table 2). The BPCAs produced from plant material by hot HCl comprised mostly benzene pentacarboxylic acid, indicating that lignin may have been a precursor. We drew this conclusion because Brunow (1965) reported formation of benzene pentacarboxylic acid upon the oxidation of acid-treated pinoresinol. BPCAs were also obtained from lignin-free microbial biomass, indicating that polysaccharides may have played an essential role in the formation of such artefacts (Table 2). The results support findings of Allard et al. (1997). They reported that hot 6 M HCl treatment of bacterial biomass produced melanoidin-like polymers and earlier findings had suggested that some of such polymers may form BPCAs during permanganate oxidation (Ishiwatari et al., 1986). 3.3. Elimination of artifactual benzene polycarboxylic acid formation Several pre-treatments were tested to see if any of them might eliminate the method-induced formation of BPCAs. Lowering of temperature or acid concentration reduced yields but did not eliminate the problem. Pretreatment with sodium dithionite, sodium oxalate or hydrofluoric acid yielded irreproducible results (data not shown) and the samples were difficult to handle. Pre-treating the plants with 4 M TFA (at the recommended 4 h, 105 C; originally used for carbohydrate extraction Amelung et al., 1996) did not result in any detectable BPCA formation, however (Fig. 2d). In contrast, charred plant material (Fig. 2e) and a BC-rich soil (Fig. 2f) readily yielded BPCAs after pre-treatment with TFA. Tests were then required to assure that interfering compounds, such as polyvalent cations, could be sufficiently removed by TFA. For this purpose we added maize straw and charred maize straw to a Chernozem (Soil 1) sample. As shown in Table 3, adding different amounts of plant material (Z. mays) to soil neither led to any additional BPCA formation nor to loss of BPCAs found in the soil during further sample processing. Between 89% and 102% of the added charred plant material was recovered. When HCl was replaced with TFA, it had to be ascertained whether all further steps in the sample processing remained reliable. Therefore, we used various soil
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Fig. 2. Gas chromatograms of benzene polycarboxylic acids obtained from (a) charred Zea mays after HCl digestion and HNO3 oxidation, (b) Zea mays after HCl digestion and HNO3 oxidation, (c) Zea mays without HCl pre-treatment and HNO3 oxidation only. The other chromatograms show the BPCA yields after replacement of the HCl pre-treatment with 4 M TFA (4 h, 105 C) and subsequent HNO3 oxidation of (d) Zea mays, (e) charred Zea mays, and (f) a Haplic Chernozem. (1) Citric acid (internal standard 1); (2) biphenyl-2,2 0 -dicarboxylic acid (internal standard 2); (3) hemimellitic acid; (4) trimellitic acid; (5) trimesic acid; (6) pyromellitic acid; (7) mellophanic acid; (8) prehnitic acid; (9) benzene pentacarboxylic acid; (10) mellitic acid. Table 2 Method-induced formation (wt.%) of benzene polycarboxylic acids after digestion with 32% HCl at 170 C for 4 h, followed by 8 h treatment with 65% HNO3 at 170 C according to the protocol of Glaser et al. (1998) Sample
Benzene polycarboxylic acid Hemimellitic acid
Trimellitic acid
Trimesic acid
Pyromellitic acid
Benzene pentacarboxylic acid
Mellitic acid
Sum
Plants Astragalus sp. Cymbaria dahurica Leperiza sp. Populus pseudosimonii Secale cereale Setaria viridis Zea mays
0.02 0.01 0.01 0.01 0.04 0.00 0.03
0.07 0.03 0.02 0.01 0.12 0.00 0.11
0.01 0.00 0.00 0.00 0.01 0.00 0.01
0.27 0.10 0.07 0.04 0.71 0.01 0.39
1.39 0.42 0.26 0.16 2.14 0.05 1.24
0.64 0.22 0.17 0.04 1.25 0.03 0.63
2.41 0.78 0.53 0.26 4.28 0.09 2.40
Bacteria Pseudomonas putida Agrobacterium tumefaciens
0.00 0.02
0.01 0.02
0.00 0.02
0.48 0.30
0.21 0.14
0.18 0.13
0.88 0.62
Fungi Penicillium citrinum
0.05
0.06
0.04
0.35
0.25
0.23
0.98
materials to re-evaluate the recovery of added BPCAs. The mean recovery of the added standard solutions ranged from 75% to 114% (Table 4). Neither the presence of lime (Kastanozem) nor interfering Fe cations (Ferralsol)
reduced the recovery significantly. Neither did the amount of standard solution added have an impact on the percentage of BPCAs recovered (Table 4). As a result, when HCl is replaced with TFA, the purification
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procedure as suggested by Glaser et al. (1998) with the changes mentioned above needs no further adaptation.
101.7 96.9 89.3
Recovery (%)
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
23.69 29.47 40.80 ZM, Zea mays. Pseudomonas putida DSM 291T, Agrobacterium citrinum ATCC 11095, Penicillium citrinum DSM 62830 did not yield any BPCA as well. a
b
0 5.45 6.77 6.43 6.45 6.67 8.30 9.12 10.74 ZMa,b Charred ZM (20.5 mg) Soil Soil + ZM (5.5 mg) Soil + ZM (10.2 mg) Soil + ZM (20 mg) Soil + charred ZM (4.9 mg) Soil + charred ZM (10 mg) Soil + charred ZM (20 mg)
0 0.78 0.43 0.41 0.42 0.44 0.60 0.82 1.15
0 1.10 0.52 0.51 0.50 0.54 0.79 1.04 1.51
0 0.09 0.04 0.03 0.05 0.06 0.06 0.07 0.10
0 1.64 1.00 0.96 0.95 1.05 1.32 1.69 2.28
0 1.76 0.95 0.90 0.90 0.98 1.31 1.73 2.40
0 2.78 1.72 1.60 1.66 1.76 2.41 3.01 3.99
0 9.64 6.71 6.39 6.39 6.79 9.30 11.06 14.25
0 23.23 18.14 17.24 17.31 18.29 24.09 28.55 36.43
Calculated sum (lg injected amount1) Sum (lg injected amount1) Mellitic acid Benzene pentacarboxylic acid Prehnitic acid Mellophanic acid Pyromellitic acid Trimesic acid Trimellitic acid Hemimellitic acid
Benzene polycarboxylic acid (lg injected amount1) Sample
Table 3 Yield and recovery of benzene polycarboxylic acids from soil (Haplic Chernozem), charred maize, and soil spiked with charred maize (using modified method)
3.4. Correction of data obtained with the original method Because the original method has been used extensively (e.g., Glaser et al., 2000; Schmid et al., 2002; Kleber et al., 2003), we conducted additional analyses to see if the data obtained by that method could be corrected. When different sample sets between digestion procedures with TFA and HCl were compared no clear correlation was observed (r2 = 0.24). Even though the overall correlation between the former and modified methods was low, exceptions are possible. For instance, when Glaser and Amelung (2003) compared data from the old and modified methods they observed similar trends in BC content across the studied sample sequence. Nevertheless, further evaluation is warranted before the modified method can be used for estimating the function of BC as a sink for atmospheric CO2 in a range of environments. 3.5. Conversion factor Of course, not all BC C is recovered as BPCA C. Therefore, a conversion factor was needed to allow the calculation of the absolute BC content of a given sample. Glaser et al. (1998) analyzed four commercially available charcoals and inferred that actual BC content exceeded BPCA yield by a factor of 2.27. These analyses were carried out without HCl digestion. But because coals and charcoals are defined as BC, no artefact formation should occur using HCl. We tested whether analyses of coals and charcoals that include the TFA pre-treatment produce similar results as analyses that include the HCl pre-treatment and found that they do (data not shown). Our alterations to the method show an agreement with Glaser et al. (1998) in that a conversion or correction factor is needed, but we failed to agree with the 2.27 they established. Yet, none of our analyses found a factor lower than 2.27, so we believe that to be the conservative estimate of the BC content of soils. The maximum we found exceeded 4.5, which is too high for subsoils (Brodowski et al., unpublished data), where the factor 2.27 allows BC to explain up to 50% of total C, but a factor of 4.5 or greater would have accounted for more than 100% of total C. Hence, we refute the hypothesis that using the factor 2.27 always makes calculation of BC content accurate. The conversion of BC to BPCAs depends on the degree of aromatic condensation (see Glaser et al., 1998). This degree is unknown and we are not aware of any accurate method available to reliably estimate the degree of BC condensation in the solid state. Applying C/H ratios for estimating BC condensation requires,
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3.6. Other sources of benzene polycarboxylic acids
for instance, that the sample be free of impurities. (A reliable re-evaluation of the conversion factor is impossible without suitable standards with known amount of quaternary C atoms. If any readers are able to provide us with such material in the near future we invite them to do so). Therefore, we recommend, in cases where absolute estimations of BC contents in soil are required, that either authentic BC from an individual scientific context is used as a standard or that working with the conversion factor of 2.27 of Glaser et al. (1998) is continued because it is (i) not affected by our method alterations, and (ii) appears to be the minimum conversion factor assessed so far. Nevertheless, one has to be aware that this calculation is only an approximation and the actual BC contents in soil may be greater. Besides, this method may not include graphitic carbon which warrants further attention.
Applying our revised method to synthetic melanoidins and pseudo-melanoidins gave a mean yield of 13 mg BPCA C per g melanoidin C. The main BPCAs detected were tetra- and pentacarboxylic acids (Table 5). This finding apparently contradicts those of Glaser et al. (1998) who reported that BPCAs were not produced from melanoidins. However, the procedure they used for melanoidin formation differed from that of Hedges (1978) in that the pH was not adjusted to alkaline conditions with Na2CO3. The values we found for BPCA C in melanoidins exceed the amounts found by Ishiwatari et al. (1986). Recently, melanoidins were hypothesized to make up a substantial part of the refractory, resistant organic residue in soils (Poirier et al., 2000). Nevertheless, clear evidence for the existence of
Table 4 Recovery (% added amount) of spiked benzene polycarboxylic acids after hydrolysis with 4 M TFA (4 h, 105 C; n = 3–6; standard error in parentheses) Sample
Depth (cm)
Hemimellitic acid
Trimesic acid
Pyromellitic acid
Benzene pentacarboxylic acid
Mellitic acid
Spike with 100 lg BPCA standard before the purification step Haplic Chernozem 0–10 75 (0.4) 77 (0.4) Haplic Chernozem 30–40 71 (0.9) 73 (0.9) Haplic Kastanozem 10–25 76 (0.3) 77 (0.1) Xanthic Ferralsol 0–10 84 (5.6) 87 (5.6)
71 68 72 87
76 74 72 90
99 87 92 86
103 (15.5) 99 (9.3) 97 (2.9) 93 (6.0)
Mean
75
77
Trimellitic acid
79
(0.4) (2.3) (0.3) (6.1)
(1.9) (0.5) (0.1) (5.8)
78
91
Spike with 100 lg BPCA standard after the purification step Haplic Chernozem 0–10 80 (3.9) 84 (4.1) Haplic Chernozem 30–40 84 (5.8) 88 (5.7) Haplic Kastanozem 10–25 85 (6.7) 88 (7.6) Xanthic Ferralsol 0–10 85 (4.7) 89 (4.9)
83 86 90 89
Mean
84
87
Spike with 25 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
before the purification step 97 (2.6) 96 (1.8) 96 (0.3) 95 (1.2) 96 (3.1) 96 (2.5) 94 (2.3) 95 (0.1)
95 94 95 93
Mean
96
94
Spike with 10 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
before the purification step 93 (3.4) 94 (3.6) 94 (2.7) 95 (1.4) 95 (0.4) 95 (1.2) 98 (4.6) 97 (5.2)
95 95 95 96
Mean
95
95
114
Spike with 10 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
after the purification step 97 (3.4) 100 (3.0) 92 (2.4) 95 (1.6) 93 (3.7) 96 (3.7) 94 (2.3) 96 (2.5)
102 (1.0) 99 (1.5) 99 (1.4) 100 (0.6)
121 120 108 105
Mean
94
100
114
87
96
95
97
(4.9) (6.1) (8.7) (5.3)
86 90 91 92
(5.7) (6.3) (13.0) (5.1)
90 (0.7) (0.7) (2.6) (0.5)
108 109 113 108
(1.7) (0.1) (1.0) (0.8)
110 (3.9) (1.5) (0.5) (4.0)
(15.2) (6.2) (3.6) (5.5)
112 122 110 111
(5.2) (3.3) (2.3) (4.6)
(5.1) (3.0) (5.0) (2.1)
79 93 87 81
98 (6.3) (6.8) (4.6) (6.3)
90 (5.6) 102 (8.6) 101 (7.5) 88 (6.7)
85
95
110 (10.2) 102 (0.7) 101 (7.5) 94 (11.6)
112 107 105 112
102
109
83 (3.1) 83 (15.0) 101 (23.3) 92 (7.6)
95 (1.4) 107 (14.6) 116 (19.4) 83 (19.9)
90
100
104 (8.6) 99 (13.9) 107 (10.9) 98 (4.4)
102 (9.5) 108 (4.7) 106 (28.1) 84 (24.4)
102
100
(11.4) (0.3) (1.9) (0.4)
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
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Table 5 Benzene polycarboxylic acids yields (g C kg1 C) from pseudo-melanoidins, melanoidins and Aspergillus niger (dry weight; without conversion factor) Sample
B3CAa
B4CAb
B5CAc
B6CAd
Sum
Glucose (pseudo-melanoidin) Glucose–Glutamic acid Glucose–Lysine Glucose–Glycine
1.51 3.42 1.98 0.62
7.43 7.49 4.68 1.71
9.06 4.26 2.76 0.91
2.95 1.51 1.55 0.30
20.96 16.68 10.96 3.54
1.88 0.48
5.33 1.34
4.25 0.73
1.58 6.01
13.03 8.55
9:1 9:1 9:1
Mean Aspergillus niger a b c d
B3CA, B4CA, B5CA, B6CA,
benzene tricarboxylic acids. benzene tetracarboxylic acids. benzene pentacarboxylic acid. mellitic acid.
(pseudo-)melanoidins in soils and an unambiguous distinction between melanoidins and BC are still lacking (see above). We conclude, therefore, that Ô(pseudo-)melanoidinsÕ do not contribute significantly to the BPCAs found in soils and if melanoidins are generated during vegetation fires, these melanoidin-derived BPCAs can be assigned as BC. We did not find any BPCAs after oxidation of bacteria or the fungus Penicillium citrinum. Only Aspergillus niger yielded 0.86% BPCA C of the fungal C, mainly
(>70%) as mellitic acid derived from the black amorphous pigment aspergillin (Fig. 3, Table 5), indicating its aromatic nature as reported by Lund et al. (1953) and Ray and Eakin (1975). Hence, other sources, all of which need to be evaluated, contributed to the BPCAs from soil. The BPCA pattern could be an indicator of its source. In this study, the BC produced by charring exhibited a BPCA pattern similar to that of melanoidins formed at high temperatures, but with more mellitic acid
Fig. 3. Benzene polycarboxylic acid pattern from soils (Haplic Chernozem at the depths 0–10 and 30–40 cm, Haplic Kastanozem at a depth of 10–25 cm, Xanthic Ferralsol at a depth of 0–10 cm; for details P see Table 1, No. 1a–3), charred plant material (oak P wood, Zea mays, Secale cereale), melanoidins and Aspergillus niger (B3CA = hemimellitic, trimellitic, trimesic acids; B4CA = pyromellitic, mellophanic, prehnitic acids; B5CA = benzene pentacarboxylic acid; B6CA = mellitic acid).
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and less benzene tetracarboxylic acids. The BPCA pattern of the soils seemed to result from a complex mixture of charred plant material and unburned Aspergillus niger (Fig. 3). Thus, the importance of biologically produced BPCA C needed to be estimated. We calculated a maximum amount of BPCAs that might have been produced by the fungus. It was negligibly small; only 0.26 g highly aromatic carbon per kg organic C can be derived from aspergillin (without conversion factor), that being the case only if we assume a living fungal biomass of 5 mg C kg1 soil, i.e. 10 times higher than supposed by Dunger (1974), 20 g kg1 organic C in soil, a worst-case contribution of 10% of Aspergillus niger to the fungal biomass and an enrichment factor of 1000 for the selective preservation of aspergillin during humification and litter decomposition. Hence, only when the BPCA yield is less than or equal to 1.2 g kg1 C (without conversion factor) will competing BPCA sources such as aspergillin significantly add (up to 20%) to the total BPCA yield. The soils used for method evaluation and validation had BPCA yields of more than 19.8 g kg1 C (Table 1). This is far beyond the range where BPCA sources other than BC may play a significant role. Thus, in these soils BPCAs remain reliable markers for BC.
4. Conclusions The original method of Glaser et al. (1998), which uses BPCAs as markers for the determination of BC in complex matrices like soil, contains an intrinsic error and requires revision. The major constraint in the original method is method-induced BPCA formation from the use of hot 32% HCl for the removal of interfering cations. Minor constraints are unreliable quantification of low BPCA yields with one-point calibrations, nonadjusted HNO3 aliquots for further sample processing (it results in significant losses of the surrogate standard citric acid under strong acid conditions) and lack of additional sample storage to allow complete derivatization. It is strongly recommended the hot HCl treatment be replaced with 4 M TFA (4 h, 105 C; Amelung et al., 1996) because it eliminates artefact formation. The other constraints are solved by using overall linear calibration only at P7 ng BPCA injection amount and by using a maximum 2 ml HNO3 sample aliquot only for further sample and internal standard processing. The samples are stored 24 h prior to analysis using GC. Data obtained with the original method (Glaser et al., 1998) cannot be corrected for different sample sets. We found evidence that BPCAs in soils may not originate solely from BC but also from biological sources like the pigment aspergillin. Elucidating the ecological significance of such biological BC production might warrant further attention. We estimate that the error in using BPCAs as markers for BC is lower than 20%
when BPCA yields are not lower than 1.2 g kg1 organic C. Such low BPCA yields are not common. Thus, using BPCAs as specific markers for BC in soils is still recommended.
Acknowledgments The authors acknowledge the helpful support of Prof. Dr. Wolfgang Zech. We also thank Elisabeth Keese for providing the microbial biomass and Tanja Gonter and Katja Poxleitner for their help in the laboratory. We gratefully acknowledge the constructive advice of Dr. Jose´ A. Gonza´lez, an anonymous reviewer and Dr. Geoff Abbott. This work was funded by the Deutsche Forschungsgemeinschaft (DFG Ze 154/48-1 and 48-2) and the Hanns-Seidel-Stiftung e.V. Associate Editor—G.D. Abbott
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Revised black carbon assessment using benzene polycarboxylic acids S. Brodowski b
a,c,* ,
A. Rodionov b, L. Haumaier a, B. Glaser a, W. Amelung
c
a Institute of Soil Science and Soil Geography, University of Bayreuth, 95440 Bayreuth, Germany Institute of Soil Science and Plant Nutrition, Martin Luther University Halle-Wittenberg, 06108 Halle, Germany c Institute of Soil Science, University of Bonn, Nussallee 13, 53115 Bonn, Germany
Received 8 January 2004; accepted 31 March 2005 (returned to author for revision 3 March 2004) Available online 26 May 2005
Abstract Black carbon (BC), the ubiquitous stable product of incomplete combustion, is believed to be a potential sink for atmospheric CO2 and therefore a contributor to the EarthÕs radiative heat balance. Nevertheless, analytical procedures to measure BC are inconsistent, giving a non-systematic variation by factors of 14–571 for estimates of its content in soil. We hypothesized that the HCl used to isolate benzene polycarboxylic acids (BPCAs) as markers for BC helps form these compounds, which could then cause an overestimation of the BC content of the soil. We found that indeed up to 90% of BPCA yields may be attributed to this HCl pre-treatment. To correct this error we developed a revised method that uses BPCAs as BC markers but which allows us to eliminate any confusion in the results. This aim was achieved by digestion with 4 M trifluoroacetic acid (TFA). After oxidation with HNO3, the BPCAs were purified using a cation exchange resin and derivatized to form the trimethylsilyl derivatives. Analyses were performed using a gas chromatograph (GC) equipped with a flame ionization detector (FID); constant linearity was obtained at P7 ng BPCA injection amount and peak purity was determined using mass spectrometry (MS). The recovery of the BPCAs averaged 93.5 ± 5.1% for pure standards and 95.0 ± 3.6% for spiked charred plant material. The contribution of BPCAs from aspergillin to soil organic carbon was estimated to be negligible. No close correlation between the results obtained with the original method and our revised procedure was observed. 2005 Elsevier Ltd. All rights reserved.
1. Introduction Black carbon (BC), a product of incomplete combustion of biomass and fossil fuel, is found nearly everywhere due to widespread aeolian transport (Goldberg, 1985; Stoffyn-Egli et al., 1997). The BC particles comprise a *
Corresponding author. Tel.: +49 228 73 2194; fax: +49 228 73 2782. E-mail address: [email protected] (S. Brodowski).
wide continuum of combustion residues ranging from char and charcoal to condensates, such as soot (Jones and Chaloner, 1991; Goldberg, 1985). BC is relatively inert. Because of this, Kuhlbusch (1998), for instance, suggested that it may be a potential sink for atmospheric CO2. If so, it could well contribute to the EarthÕs radiative heat balance (Crutzen and Andreae, 1990). A number of authors have developed an interest in the presumed impact of BC on the global carbon cycle and have begun intensified research into the fate of charred organic material in various environmental compartments: the
0146-6380/$ - see front matter 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.orggeochem.2005.03.011
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atmosphere (Chang et al., 1982; Charlson and Ogren, 1982; Andreae et al., 1994; Bird, 1997), (marine) sediments (Herring, 1985; Novakov et al., 1997; Masiello and Druffel, 1998; Middelburg et al., 1999) and soils (Skjemstad et al., 1996; Schmidt et al., 1999; Schmidt and Noack, 2000; Schmidt et al., 2002). Despite growing interest in BC, both a generally accepted definition and standard analytical procedures are lacking (Schmidt and Noack, 2000; Schmidt et al., 2001). As pointed out by Schmidt et al. (2001), comparative results from six methods revealed a non-systematic variation in soil BC content by factors of 14–571. Thermal and UV-oxidation procedures have been criticized as overestimating BC content of soils and sediments (Simpson and Hatcher, 2004). To accurately quantify BC in the environment there is, therefore, a need to improve current methods. Glaser et al. (1998) proposed the use of benzene polycarboxylic acids (BPCAs) as specific markers for BC (Fig. 1). Their method involves preextraction with hot 32% HCl, oxidation of condensed aromatics to BPCAs using hot HNO3, purification and silylation of the products prior to quantification using GC with FID. Some drawbacks in the method became obvious, however, when it was applied to a large number of samples. Unacceptable analytical errors arose whenever samples with a wide range of weight and containing small amounts of BC were analyzed (Brodowski et al., unpublished data). Moreover, the test was unable to re-
veal whether the BPCAs came from artifactual formation or additional natural sources. For instance, treating organic matter or pinoresinol, a constituent of lignin (Freudenberg et al., 1965), with strong acid resulted in a formation of BPCAs (Brunow, 1965; Rudakov et al., 1986). Strong acid treatment of bacterial biomass or of a mixture of monosaccharides and amino acids produced melanoidin-like artefacts (Allard et al., 1997). Upon permanganate oxidation at least some melanoidins yield BPCAs (Ishiwatari et al., 1986). This finding led us to suspect that artefact formation in the pre-treatment step of the method might cause further methodological errors. Apart from artificial formation, BPCAs could, in principle, also be derived from melanoidins, the BPCA yields from which are unknown. We agree with those who say that the Maillard reaction (non-enzymatic browning) does not appear to contribute any significant amount of SOM formation (Hatcher, 2003) or degradation of algae (Zang et al., 2001). We therefore disagree with those who contend that the process is a possible humification source for soils and marine environments (Nissenbaum and Kaplan, 1972; Stevenson, 1982; Ertel and Hedges, 1983; Rubinsztain et al., 1984; Poirier et al., 2000). The reason for this is that the process involves a number of reactions, some of which take place at temperatures substantially warmer and pH substantially more alkaline than those commonly found in soil. While
Fig. 1. Structures of benzene polycarboxylic acids used as markers for black carbon assessment.
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
high temperatures are known to form heterocyclic N and polyaromatic C from soil organic residues (Almendros et al., 2003), the only time that high temperatures occur naturally in soil is during a wildfire or controlled burning. Maillard reactions are not sustained at any substantial scale when fires are absent. We therefore suggest that any melanoidin-derived aromatic structure formed in soils should be assigned as BC. Reports of BPCA formation from some of the melanoidins may warrant attention. Other natural products in soils, upon oxidation after treatment with strong acids, can produce BPCAs. One of these is aspergillin, a black amorphous pigment from Aspergillus niger (Lund et al., 1953) that is supposed to contribute to the aromaticity of soil humic substances (Kononova and Aleksandrova, 1959). At the latter time, aspergillin had not been analyzed using the method of Glaser et al. (1998). The main aims of the present study were (1) to eliminate the analytical problem in samples with low BC contents when analyzed using the method of Glaser et al. (1998); (2) to examine whether the HCl treatment in this method artificially forms BPCAs and, thus overestimates BC contents of soils and if so, to establish an alternative pre-treatment; (3) to determine the extent to which synthetic melanoidins and naturally occurring aspergillin yield BPCAs using this method.
2. Materials and methods 2.1. Samples We obtained BC-free materials from above-ground biomass (stems and leaves) of seven plants (Astragalus sp., Cymbaria dahurica L., Leperiza sp., Populus pseudosimonii var. patula T.Y. Sun, Secale cereale L., Setaria viridis (L.) P. Beauv., Zea mays L.); and from soilinhabiting microorganisms such as the fungi Aspergillus niger DSM B202, Penicillium citrinum DSM 62830 and the bacteria Pseudomonas putida DSM 291T and Agrobacterium tumefaciens ATCC 11095. Melanoidins were prepared by condensation of one amino acid (glycine, glutamic acid or lysine) and a sugar (D-glucose) in hot alkaline solution as described by Hedges (1978). The sugar to amino acid concentration ratios were 9:1 and 1:9 (v:v), respectively. Pseudomelanoidins were produced in a pure sugar solution without adding amino acids. For alkaline conditions, 0.167 M amino acid or sugar solutions were prepared in 0.1 M Na2CO3. The reaction was carried out by heating 200 ml of reaction solution in glass bottles to 100 C for 170 h. Thereafter, the melanoidins and pseudomelanoidins were precipitated by acidification with 1 M TFA to pH 1. After 16 h the precipitated melanoidins were centrifuged (4420g, 15 min), washed with
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deionized water, which was acidified with TFA to pH 1, again centrifuged, freeze-dried and subjected to BPCA analysis. We failed to produce a precipitate for a sugar to amino acid ratio of 1:9 (v:v). All reactions were carried out in quadruplicate to produce enough material for BC analysis. For the method evaluation, we used ground straw of Z. mays L., the fungus Penicillium citrinum DSM 62830, the two bacteria P. putida DSM 291T and A. tumefaciens ATCC 11095 and various soil samples with different properties from various climatic regimes (Table 1). The maize straw was prepared for charring by air drying and chopping to <5 cm pieces. These materials were placed in stainless steel containers closed with a cap to reduce oxygen supply. The containers were heated to a temperature of 350 C in a muffle furnace. It took 1 h to reach this temperature and heating continued thereafter for 2 h. The mass loss after heating averaged 67.6 ± 0.47% (SE; n = 12) of the initial mass. The charred maize contained 66.35% C, 3.36% H, 16.00% O, and 2.50% N. 2.2. Analytical procedures 2.2.1. Elimination of polyvalent cations, oxidation and purification For metal elimination about 0.5 g of each sample was subjected to digestion either with 2 ml of 32% HCl [4 h, 170 C; Glaser et al. (1998) original method] or with 10 ml of 4 M TFA [4 h at 105 C; revised method]. Additionally, several extractants (32% HCl, 6 M HCl, sodium dithionite, sodium oxalate, hydrofluoric acid) were tested for metal elimination at two temperatures (room temperature, 105 C) and three digestion times (16, 8 and 4 h). These alternatives were, however, unmanageable or gave irreproducible results or both (data not shown). When the sample was expected to contain more than 200 g BC per kg organic C and/or more than 100 g kg1 organic C, we reduced the sample weight to 5 mg organic C. After digestion with either acid, the residue was processed as outlined by Glaser et al. (1998) with additional modifications concerning the reduction of HNO3 sample aliquots to 62 ml, extended storage time of the derivatization solution (P24 h). In brief, samples were processed as follows. The residue was collected by filtration through a glass fibre filter (GF 6, Schleicher and Schuell, Dassel, Germany), rinsed several times with deionized water and dried at 30–40 C for at least 2 h. Thereafter, all of the residue was transferred to a glass digestion tube, 2 ml of 65% HNO3 were added and the mixture was heated to 170 C and held there for 8 h in a high pressure digestion apparatus (Schramel et al., 1980) where BC was oxidized to BPCAs (Fig. 1). Repeated oxidation of BC generally reproduced the BPCA yields with a variation of 3.5% (median) or 6.9
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Table 1 Selected climatic data and properties of soils used for method optimization [benzene polycarboxylic acids (BPCAs) determined using revised method; without conversion factor] No.
Site identification
Samples used for method evaluation 1a RUSSIA: Kursk 1b 2 RUSSIA: Frolovo 3 BRAZIL: Belterra
MAPa (mm)
MATb (C)
Depth (cm)
Organic C (%)
C/N
BPCAC (g kg1 organic C)
Haplic Chernozem
573
5.3
Haplic Kastanozem Xanthic Ferralsol
300 2050
6.0 26.5
0–10 30–40 10–25 0–10
5.64 3.45 2.29 1.44
12.0 12.3 10.4 10.3
49.3 88.5 65.2 19.8
480
8.7
490
9.1
890
8.2
0–25 0–20 54–64 0–25 0–25 0–30 0–30
2.01 1.33 0.27 1.30 1.25 1.36 1.30
11.2 9.4 6.6 15.5 15.2 8.3 8.2
63.5 53.7 58.7 65.2 63.9 20.1 20.0
Soil type (FAO, 1988)
Further samples used for method validation 5 GERMANY: Bad Lauchsta¨dt Haplic Chernozem 6a 6b 7 GERMANY: Halle Luvic Phaeozem 8 9 GERMANY: Rotthalmu¨nster Stagnic Luvisol 10 a b
MAP, mean annual precipitation. MAT, mean annual temperature.
(mean) ± 2.0% (standard error; n = 13) of the mean. The solution was poured through an ash-less cellulose filter (Schleicher and Schuell 5893; d = 70 mm) into a 10 ml volumetric flask. Deionized water was added to fill the flask. To eliminate remaining polyvalent cations, such as of Fe3+ or Al3+, the following procedure was used. To avoid degradation of the internal standard by the strong acid, an aliquot (0.5–2 ml) of the sample solution was diluted with 4 ml deionized water (the original method processed up to 5 ml aliquots). Subsequently, internal standard 1 was added (100 lg citric acid in 100 ll deionized water). The solution was then dropped on to a cation exchange resin (Dowex 50 W X 8, 200– 400 mesh, Fluka, Steinheim, Germany), which was converted to the H+ form prior to use. After complete infiltration, the BPCAs were eluted into 100 ml conical flasks with 50 ml of deionized water in portions of 10 ml. Finally, the eluates were freeze-dried for water removal. We also learned that the use of an aliquot >2 ml for further sample processing should be avoided. It sometimes resulted in significant decomposition of the citric acid due to strong acid conditions and possibly an incomplete removal of interfering cations during purification and subsequent complexation of BPCAs (data not shown). All reagents were of analytical grade and purchased from Sigma/Aldrich or Fluka. 2.2.2. Derivatization The freeze-dried BPCAs were re-dissolved in 4 · 1 ml methanol and transferred to 5 ml reactivials (Alltech GmbH, Unterhaching, Germany) with Teflon-laminated septum screw caps. One hundred microliters of
the second internal standard solution containing 100 lg of biphenyl-2,2 0 -dicarboxylic acid in methanol were added and the solvent was evaporated with a stream of air. The dried BPCAs were converted to trimethylsilyl derivatives. For this purpose, we added 100–125 ll dry pyridine and 100–125 ll N,O-bis(trimethylsilyl)-trifluoroacetamide, heated the solution at 80 C for 2 h, allowed it to cool and stored it for 24 h. During the 24 h the detector signals increased (see below) and then remained stable. We conclude that derivatization was not complete without the 24 h storage. 2.2.3. GC GC analysis was performed with a Hewlett Packard 6890 gas chromatograph (Hewlett Packard GmbH, Waldbronn, Germany) equipped with a FID and an HP-5 capillary column (30 m · 0.32 mm i.d., 0.25 lm film thickness). Helium was used as carrier gas at a constant flow of 0.8 ml min1. Both the injector and detector temperatures were 300 C. Aliquots (2 ll) of sample solution were injected at a split ratio of 30:1 into a fully deactivated inlet system with silylated liners. The temperature programme was: initial column temperature of 100 C held for 2 min followed by an increase in 20 C min1 to 240 C and held for 7 min. Subsequently, the temperature was raised at 30 C min1 to 300 C and held for an additional 10 min (total run time = 28 min). Peak identity was confirmed with full scan mass spectra obtained using a Hewlett Packard 6890 series GC coupled to a Hewlett Packard 5972A mass selective detector (Hewlett Packard GmbH, Waldbronn, Germany).
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
2.2.4. Standard series Standard series were prepared for each analysis in two- to three-fold replication using standard solutions of 5, 10, 15, 25, 50, 100 lg per vial of BPCA (hemimellitic acid, trimellitic acid, trimesic acid, pyromellitic acid, benzene pentacarboxylic acid, mellitic acid). Linearity was checked with additional standard solutions of 20, 30, 40, 60, 80, 120, and 150 lg per vial of BPCA. These were prepared by dissolving 100 lg of each BPCA in 100 ll methanol and transferring the corresponding volume of standard solution (e.g., 5 ll or 10 ll etc.) into a vial, then containing, e.g., 5 lg or 10 lg of each BPCA. The content of the internal standards was kept constant at 100 lg of citric acid and 100 lg biphenyl-2,2 0 -dicarboxylic acid per vial, both in 100 ll methanol. Subsequently, the standards were completely dried and individually derivatized as described above.
3. Results and discussion 3.1. Linearity Previous experiments using different sample amounts for analysis according to the method of Glaser et al. (1998) revealed high coefficients of variation (P50% of the mean). In these earlier experiments, however, a one-point calibration was used. Derivatizing standard mixtures of different concentrations revealed an apparent linearity of detection (r2 = 0.9964 and higher). The coefficient of regression was only slightly improved when using a second-order regression curve. However, as previously reported by Amelung and Zhang (2001) for MS analyses of amino acid enantiomers, even an r2 close to unity may blur a non-linearity at low concentrations. Indeed, when calibration curves were forced through the origin, signal response of the FID was non-linear at <7 ng injection amount of silylated BPCAs (data not shown) and this non-linearity could be reproduced. For minimization of analytical errors we recommend, however, that sample weight be adapted to give >7 ng of expected BPCA yield, if possible. 3.2. Method-induced production of benzene polycarboxylic acids When we used the Glaser et al. (1998) method to analyze charred maize straw we found BPCAs, as expected (Fig. 2a). We also unexpectedly found significant amounts of BPCAs (Fig. 2b), however, when we used the same method to analyze plant materials which we knew were free of BC. Therefore, we concluded that the original method of Glaser et al. (1998) produced BPCAs. That method involves acid digestion with hot 32% HCl, and we suspected that there was acid-catalyzed BPCA formation, leading to
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an overestimation of BC contents in environmental samples. No BPCAs were found when oxidation was carried out without the HCl pre-treatment (Fig. 2c). Recently, overestimation has also been reported for thermal and UV-oxidation procedures used to determine BC in the environment (Simpson and Hatcher, 2004). The method-induced BPCA formation was not directly proportional to the amount of plant material added to soil (data not shown) and accounted for 0.1–4.3% of plant and microbial biomass (Table 2). The BPCAs produced from plant material by hot HCl comprised mostly benzene pentacarboxylic acid, indicating that lignin may have been a precursor. We drew this conclusion because Brunow (1965) reported formation of benzene pentacarboxylic acid upon the oxidation of acid-treated pinoresinol. BPCAs were also obtained from lignin-free microbial biomass, indicating that polysaccharides may have played an essential role in the formation of such artefacts (Table 2). The results support findings of Allard et al. (1997). They reported that hot 6 M HCl treatment of bacterial biomass produced melanoidin-like polymers and earlier findings had suggested that some of such polymers may form BPCAs during permanganate oxidation (Ishiwatari et al., 1986). 3.3. Elimination of artifactual benzene polycarboxylic acid formation Several pre-treatments were tested to see if any of them might eliminate the method-induced formation of BPCAs. Lowering of temperature or acid concentration reduced yields but did not eliminate the problem. Pretreatment with sodium dithionite, sodium oxalate or hydrofluoric acid yielded irreproducible results (data not shown) and the samples were difficult to handle. Pre-treating the plants with 4 M TFA (at the recommended 4 h, 105 C; originally used for carbohydrate extraction Amelung et al., 1996) did not result in any detectable BPCA formation, however (Fig. 2d). In contrast, charred plant material (Fig. 2e) and a BC-rich soil (Fig. 2f) readily yielded BPCAs after pre-treatment with TFA. Tests were then required to assure that interfering compounds, such as polyvalent cations, could be sufficiently removed by TFA. For this purpose we added maize straw and charred maize straw to a Chernozem (Soil 1) sample. As shown in Table 3, adding different amounts of plant material (Z. mays) to soil neither led to any additional BPCA formation nor to loss of BPCAs found in the soil during further sample processing. Between 89% and 102% of the added charred plant material was recovered. When HCl was replaced with TFA, it had to be ascertained whether all further steps in the sample processing remained reliable. Therefore, we used various soil
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Fig. 2. Gas chromatograms of benzene polycarboxylic acids obtained from (a) charred Zea mays after HCl digestion and HNO3 oxidation, (b) Zea mays after HCl digestion and HNO3 oxidation, (c) Zea mays without HCl pre-treatment and HNO3 oxidation only. The other chromatograms show the BPCA yields after replacement of the HCl pre-treatment with 4 M TFA (4 h, 105 C) and subsequent HNO3 oxidation of (d) Zea mays, (e) charred Zea mays, and (f) a Haplic Chernozem. (1) Citric acid (internal standard 1); (2) biphenyl-2,2 0 -dicarboxylic acid (internal standard 2); (3) hemimellitic acid; (4) trimellitic acid; (5) trimesic acid; (6) pyromellitic acid; (7) mellophanic acid; (8) prehnitic acid; (9) benzene pentacarboxylic acid; (10) mellitic acid. Table 2 Method-induced formation (wt.%) of benzene polycarboxylic acids after digestion with 32% HCl at 170 C for 4 h, followed by 8 h treatment with 65% HNO3 at 170 C according to the protocol of Glaser et al. (1998) Sample
Benzene polycarboxylic acid Hemimellitic acid
Trimellitic acid
Trimesic acid
Pyromellitic acid
Benzene pentacarboxylic acid
Mellitic acid
Sum
Plants Astragalus sp. Cymbaria dahurica Leperiza sp. Populus pseudosimonii Secale cereale Setaria viridis Zea mays
0.02 0.01 0.01 0.01 0.04 0.00 0.03
0.07 0.03 0.02 0.01 0.12 0.00 0.11
0.01 0.00 0.00 0.00 0.01 0.00 0.01
0.27 0.10 0.07 0.04 0.71 0.01 0.39
1.39 0.42 0.26 0.16 2.14 0.05 1.24
0.64 0.22 0.17 0.04 1.25 0.03 0.63
2.41 0.78 0.53 0.26 4.28 0.09 2.40
Bacteria Pseudomonas putida Agrobacterium tumefaciens
0.00 0.02
0.01 0.02
0.00 0.02
0.48 0.30
0.21 0.14
0.18 0.13
0.88 0.62
Fungi Penicillium citrinum
0.05
0.06
0.04
0.35
0.25
0.23
0.98
materials to re-evaluate the recovery of added BPCAs. The mean recovery of the added standard solutions ranged from 75% to 114% (Table 4). Neither the presence of lime (Kastanozem) nor interfering Fe cations (Ferralsol)
reduced the recovery significantly. Neither did the amount of standard solution added have an impact on the percentage of BPCAs recovered (Table 4). As a result, when HCl is replaced with TFA, the purification
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procedure as suggested by Glaser et al. (1998) with the changes mentioned above needs no further adaptation.
101.7 96.9 89.3
Recovery (%)
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
23.69 29.47 40.80 ZM, Zea mays. Pseudomonas putida DSM 291T, Agrobacterium citrinum ATCC 11095, Penicillium citrinum DSM 62830 did not yield any BPCA as well. a
b
0 5.45 6.77 6.43 6.45 6.67 8.30 9.12 10.74 ZMa,b Charred ZM (20.5 mg) Soil Soil + ZM (5.5 mg) Soil + ZM (10.2 mg) Soil + ZM (20 mg) Soil + charred ZM (4.9 mg) Soil + charred ZM (10 mg) Soil + charred ZM (20 mg)
0 0.78 0.43 0.41 0.42 0.44 0.60 0.82 1.15
0 1.10 0.52 0.51 0.50 0.54 0.79 1.04 1.51
0 0.09 0.04 0.03 0.05 0.06 0.06 0.07 0.10
0 1.64 1.00 0.96 0.95 1.05 1.32 1.69 2.28
0 1.76 0.95 0.90 0.90 0.98 1.31 1.73 2.40
0 2.78 1.72 1.60 1.66 1.76 2.41 3.01 3.99
0 9.64 6.71 6.39 6.39 6.79 9.30 11.06 14.25
0 23.23 18.14 17.24 17.31 18.29 24.09 28.55 36.43
Calculated sum (lg injected amount1) Sum (lg injected amount1) Mellitic acid Benzene pentacarboxylic acid Prehnitic acid Mellophanic acid Pyromellitic acid Trimesic acid Trimellitic acid Hemimellitic acid
Benzene polycarboxylic acid (lg injected amount1) Sample
Table 3 Yield and recovery of benzene polycarboxylic acids from soil (Haplic Chernozem), charred maize, and soil spiked with charred maize (using modified method)
3.4. Correction of data obtained with the original method Because the original method has been used extensively (e.g., Glaser et al., 2000; Schmid et al., 2002; Kleber et al., 2003), we conducted additional analyses to see if the data obtained by that method could be corrected. When different sample sets between digestion procedures with TFA and HCl were compared no clear correlation was observed (r2 = 0.24). Even though the overall correlation between the former and modified methods was low, exceptions are possible. For instance, when Glaser and Amelung (2003) compared data from the old and modified methods they observed similar trends in BC content across the studied sample sequence. Nevertheless, further evaluation is warranted before the modified method can be used for estimating the function of BC as a sink for atmospheric CO2 in a range of environments. 3.5. Conversion factor Of course, not all BC C is recovered as BPCA C. Therefore, a conversion factor was needed to allow the calculation of the absolute BC content of a given sample. Glaser et al. (1998) analyzed four commercially available charcoals and inferred that actual BC content exceeded BPCA yield by a factor of 2.27. These analyses were carried out without HCl digestion. But because coals and charcoals are defined as BC, no artefact formation should occur using HCl. We tested whether analyses of coals and charcoals that include the TFA pre-treatment produce similar results as analyses that include the HCl pre-treatment and found that they do (data not shown). Our alterations to the method show an agreement with Glaser et al. (1998) in that a conversion or correction factor is needed, but we failed to agree with the 2.27 they established. Yet, none of our analyses found a factor lower than 2.27, so we believe that to be the conservative estimate of the BC content of soils. The maximum we found exceeded 4.5, which is too high for subsoils (Brodowski et al., unpublished data), where the factor 2.27 allows BC to explain up to 50% of total C, but a factor of 4.5 or greater would have accounted for more than 100% of total C. Hence, we refute the hypothesis that using the factor 2.27 always makes calculation of BC content accurate. The conversion of BC to BPCAs depends on the degree of aromatic condensation (see Glaser et al., 1998). This degree is unknown and we are not aware of any accurate method available to reliably estimate the degree of BC condensation in the solid state. Applying C/H ratios for estimating BC condensation requires,
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3.6. Other sources of benzene polycarboxylic acids
for instance, that the sample be free of impurities. (A reliable re-evaluation of the conversion factor is impossible without suitable standards with known amount of quaternary C atoms. If any readers are able to provide us with such material in the near future we invite them to do so). Therefore, we recommend, in cases where absolute estimations of BC contents in soil are required, that either authentic BC from an individual scientific context is used as a standard or that working with the conversion factor of 2.27 of Glaser et al. (1998) is continued because it is (i) not affected by our method alterations, and (ii) appears to be the minimum conversion factor assessed so far. Nevertheless, one has to be aware that this calculation is only an approximation and the actual BC contents in soil may be greater. Besides, this method may not include graphitic carbon which warrants further attention.
Applying our revised method to synthetic melanoidins and pseudo-melanoidins gave a mean yield of 13 mg BPCA C per g melanoidin C. The main BPCAs detected were tetra- and pentacarboxylic acids (Table 5). This finding apparently contradicts those of Glaser et al. (1998) who reported that BPCAs were not produced from melanoidins. However, the procedure they used for melanoidin formation differed from that of Hedges (1978) in that the pH was not adjusted to alkaline conditions with Na2CO3. The values we found for BPCA C in melanoidins exceed the amounts found by Ishiwatari et al. (1986). Recently, melanoidins were hypothesized to make up a substantial part of the refractory, resistant organic residue in soils (Poirier et al., 2000). Nevertheless, clear evidence for the existence of
Table 4 Recovery (% added amount) of spiked benzene polycarboxylic acids after hydrolysis with 4 M TFA (4 h, 105 C; n = 3–6; standard error in parentheses) Sample
Depth (cm)
Hemimellitic acid
Trimesic acid
Pyromellitic acid
Benzene pentacarboxylic acid
Mellitic acid
Spike with 100 lg BPCA standard before the purification step Haplic Chernozem 0–10 75 (0.4) 77 (0.4) Haplic Chernozem 30–40 71 (0.9) 73 (0.9) Haplic Kastanozem 10–25 76 (0.3) 77 (0.1) Xanthic Ferralsol 0–10 84 (5.6) 87 (5.6)
71 68 72 87
76 74 72 90
99 87 92 86
103 (15.5) 99 (9.3) 97 (2.9) 93 (6.0)
Mean
75
77
Trimellitic acid
79
(0.4) (2.3) (0.3) (6.1)
(1.9) (0.5) (0.1) (5.8)
78
91
Spike with 100 lg BPCA standard after the purification step Haplic Chernozem 0–10 80 (3.9) 84 (4.1) Haplic Chernozem 30–40 84 (5.8) 88 (5.7) Haplic Kastanozem 10–25 85 (6.7) 88 (7.6) Xanthic Ferralsol 0–10 85 (4.7) 89 (4.9)
83 86 90 89
Mean
84
87
Spike with 25 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
before the purification step 97 (2.6) 96 (1.8) 96 (0.3) 95 (1.2) 96 (3.1) 96 (2.5) 94 (2.3) 95 (0.1)
95 94 95 93
Mean
96
94
Spike with 10 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
before the purification step 93 (3.4) 94 (3.6) 94 (2.7) 95 (1.4) 95 (0.4) 95 (1.2) 98 (4.6) 97 (5.2)
95 95 95 96
Mean
95
95
114
Spike with 10 lg BPCA standard Haplic Chernozem 0–10 Haplic Chernozem 30–40 Haplic Kastanozem 10–25 Xanthic Ferralsol 0–10
after the purification step 97 (3.4) 100 (3.0) 92 (2.4) 95 (1.6) 93 (3.7) 96 (3.7) 94 (2.3) 96 (2.5)
102 (1.0) 99 (1.5) 99 (1.4) 100 (0.6)
121 120 108 105
Mean
94
100
114
87
96
95
97
(4.9) (6.1) (8.7) (5.3)
86 90 91 92
(5.7) (6.3) (13.0) (5.1)
90 (0.7) (0.7) (2.6) (0.5)
108 109 113 108
(1.7) (0.1) (1.0) (0.8)
110 (3.9) (1.5) (0.5) (4.0)
(15.2) (6.2) (3.6) (5.5)
112 122 110 111
(5.2) (3.3) (2.3) (4.6)
(5.1) (3.0) (5.0) (2.1)
79 93 87 81
98 (6.3) (6.8) (4.6) (6.3)
90 (5.6) 102 (8.6) 101 (7.5) 88 (6.7)
85
95
110 (10.2) 102 (0.7) 101 (7.5) 94 (11.6)
112 107 105 112
102
109
83 (3.1) 83 (15.0) 101 (23.3) 92 (7.6)
95 (1.4) 107 (14.6) 116 (19.4) 83 (19.9)
90
100
104 (8.6) 99 (13.9) 107 (10.9) 98 (4.4)
102 (9.5) 108 (4.7) 106 (28.1) 84 (24.4)
102
100
(11.4) (0.3) (1.9) (0.4)
S. Brodowski et al. / Organic Geochemistry 36 (2005) 1299–1310
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Table 5 Benzene polycarboxylic acids yields (g C kg1 C) from pseudo-melanoidins, melanoidins and Aspergillus niger (dry weight; without conversion factor) Sample
B3CAa
B4CAb
B5CAc
B6CAd
Sum
Glucose (pseudo-melanoidin) Glucose–Glutamic acid Glucose–Lysine Glucose–Glycine
1.51 3.42 1.98 0.62
7.43 7.49 4.68 1.71
9.06 4.26 2.76 0.91
2.95 1.51 1.55 0.30
20.96 16.68 10.96 3.54
1.88 0.48
5.33 1.34
4.25 0.73
1.58 6.01
13.03 8.55
9:1 9:1 9:1
Mean Aspergillus niger a b c d
B3CA, B4CA, B5CA, B6CA,
benzene tricarboxylic acids. benzene tetracarboxylic acids. benzene pentacarboxylic acid. mellitic acid.
(pseudo-)melanoidins in soils and an unambiguous distinction between melanoidins and BC are still lacking (see above). We conclude, therefore, that Ô(pseudo-)melanoidinsÕ do not contribute significantly to the BPCAs found in soils and if melanoidins are generated during vegetation fires, these melanoidin-derived BPCAs can be assigned as BC. We did not find any BPCAs after oxidation of bacteria or the fungus Penicillium citrinum. Only Aspergillus niger yielded 0.86% BPCA C of the fungal C, mainly
(>70%) as mellitic acid derived from the black amorphous pigment aspergillin (Fig. 3, Table 5), indicating its aromatic nature as reported by Lund et al. (1953) and Ray and Eakin (1975). Hence, other sources, all of which need to be evaluated, contributed to the BPCAs from soil. The BPCA pattern could be an indicator of its source. In this study, the BC produced by charring exhibited a BPCA pattern similar to that of melanoidins formed at high temperatures, but with more mellitic acid
Fig. 3. Benzene polycarboxylic acid pattern from soils (Haplic Chernozem at the depths 0–10 and 30–40 cm, Haplic Kastanozem at a depth of 10–25 cm, Xanthic Ferralsol at a depth of 0–10 cm; for details P see Table 1, No. 1a–3), charred plant material (oak P wood, Zea mays, Secale cereale), melanoidins and Aspergillus niger (B3CA = hemimellitic, trimellitic, trimesic acids; B4CA = pyromellitic, mellophanic, prehnitic acids; B5CA = benzene pentacarboxylic acid; B6CA = mellitic acid).
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and less benzene tetracarboxylic acids. The BPCA pattern of the soils seemed to result from a complex mixture of charred plant material and unburned Aspergillus niger (Fig. 3). Thus, the importance of biologically produced BPCA C needed to be estimated. We calculated a maximum amount of BPCAs that might have been produced by the fungus. It was negligibly small; only 0.26 g highly aromatic carbon per kg organic C can be derived from aspergillin (without conversion factor), that being the case only if we assume a living fungal biomass of 5 mg C kg1 soil, i.e. 10 times higher than supposed by Dunger (1974), 20 g kg1 organic C in soil, a worst-case contribution of 10% of Aspergillus niger to the fungal biomass and an enrichment factor of 1000 for the selective preservation of aspergillin during humification and litter decomposition. Hence, only when the BPCA yield is less than or equal to 1.2 g kg1 C (without conversion factor) will competing BPCA sources such as aspergillin significantly add (up to 20%) to the total BPCA yield. The soils used for method evaluation and validation had BPCA yields of more than 19.8 g kg1 C (Table 1). This is far beyond the range where BPCA sources other than BC may play a significant role. Thus, in these soils BPCAs remain reliable markers for BC.
4. Conclusions The original method of Glaser et al. (1998), which uses BPCAs as markers for the determination of BC in complex matrices like soil, contains an intrinsic error and requires revision. The major constraint in the original method is method-induced BPCA formation from the use of hot 32% HCl for the removal of interfering cations. Minor constraints are unreliable quantification of low BPCA yields with one-point calibrations, nonadjusted HNO3 aliquots for further sample processing (it results in significant losses of the surrogate standard citric acid under strong acid conditions) and lack of additional sample storage to allow complete derivatization. It is strongly recommended the hot HCl treatment be replaced with 4 M TFA (4 h, 105 C; Amelung et al., 1996) because it eliminates artefact formation. The other constraints are solved by using overall linear calibration only at P7 ng BPCA injection amount and by using a maximum 2 ml HNO3 sample aliquot only for further sample and internal standard processing. The samples are stored 24 h prior to analysis using GC. Data obtained with the original method (Glaser et al., 1998) cannot be corrected for different sample sets. We found evidence that BPCAs in soils may not originate solely from BC but also from biological sources like the pigment aspergillin. Elucidating the ecological significance of such biological BC production might warrant further attention. We estimate that the error in using BPCAs as markers for BC is lower than 20%
when BPCA yields are not lower than 1.2 g kg1 organic C. Such low BPCA yields are not common. Thus, using BPCAs as specific markers for BC in soils is still recommended.
Acknowledgments The authors acknowledge the helpful support of Prof. Dr. Wolfgang Zech. We also thank Elisabeth Keese for providing the microbial biomass and Tanja Gonter and Katja Poxleitner for their help in the laboratory. We gratefully acknowledge the constructive advice of Dr. Jose´ A. Gonza´lez, an anonymous reviewer and Dr. Geoff Abbott. This work was funded by the Deutsche Forschungsgemeinschaft (DFG Ze 154/48-1 and 48-2) and the Hanns-Seidel-Stiftung e.V. Associate Editor—G.D. Abbott
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