Anal. Chem. 2000, 72, 3116-3121
Alkaline CuO Oxidation with a Microwave Digestion System: Lignin Analyses of Geochemical Samples Miguel A. Gon˜i*,† and Shelagh Montgomery‡
Department of Geological Sciences, University of South Carolina, Columbia, South Carolina 29208, and Chaire de recherche en environnement, Universite´ du Que´ bec a` Montre´ al, C.P. 8888, Succursale Centre-Ville, Montre´ al, Que´ bec H3C 3P8, Canada
A novel approach for the analysis of lignin in geochemical samples has been developed as an alternative to the alkaline CuO oxidation procedure first developed in 1982. The new procedure utilizes microwave digestion technology, as opposed to conductive heating, to carry out oxidative hydrolysis of six samples in an oxygen-free atmosphere at 150 °C for 90 min. Ethyl acetate is used as the extraction solvent in place of diethyl ether. Additionally, the new method incorporates a simplified extraction procedure that minimizes solvent handling and the amount of glassware needed. Under these novel conditions, the yields and compositions of lignin phenols from four different samples (modern and ancient sediments; woody and nonwoody tissues) match those obtained by the “traditional” procedure. The significant advantages of this new alkaline CuO oxidation method include faster reaction times, the ability to accurately measure and control reaction conditions, added flexibility for the analyst, and a marked increase in the achievable sample throughput. Lignin is a complex biomacromolecule composed of methoxylated phenol units linked by ether and carbon bonds.1,2 Alkaline CuO oxidation is one technique commonly used to analyze the composition of lignins in complex sample matrixes such as soils and sediments. Upon CuO oxidation, the lignin macromolecule is hydrolyzed, yielding several characteristic methoxylated phenols that are amenable to gas chromatography.3-6 The yields of these CuO reaction products have been extensively used to trace the sources and composition of lignin in plant tissues, soils, sediments, and colloids from a large number of environments.7-17 The use of CuO oxidation in geochemical studies has also grown partially * Corresponding author. Tel: 803-777-3550. Fax: 803-777-6610. E-mail: goni@ geol.sc.edu. † University of South Carolina. ‡ Universite´ du Que´bec a` Montre´al. (1) Sarkanen, K. V. Lignins; Sarkaren, K. V., Ludwig, C. H., Eds.; WileyInterscience: New York, 1971; pp 95-163. (2) Fengel, D.; Wegener, G. Wood: Chemistry, Ultrastructure, Reactions; Walter de Gruyter: New York, 1984. (3) Pearl, I. A.; Beyer, D. L. J. Am. Chem. Soc. 1954, 76, 2224-2226. (4) Pearl, I. A.; Beyer, D. L. J. Am. Chem. Soc. 1954, 76, 6106-6108. (5) Pearl, I. A. J. Am. Chem. Soc. 1956, 78, 5672-5674. (6) Hedges, J. I.; Ertel, J. R. Anal. Chem. 1982, 54, 174-178. (7) Hedges, J. I.; Mann, D. C. Geochim. Cosmochim. Acta 1979, 43, 18091818.
3116 Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
due to the discovery of additional reaction products derived from more condensed lignin structures,18 as well as from nonlignin sources, including cutin,19-21 proteins, and lipids.22,23 In addition, the recent stable carbon isotope measurements of individual lignin CuO oxidation products24,25 have further increased the applicability of the technique. As originally developed by Hedges and Ertel,6 the CuO oxidation method has several drawbacks that have limited its wider utilization. Important constraints include the relatively few samples (four) that can be analyzed during a single oxidation, the long duration of the reaction (3 h of hydrolysis), and the analyst-intensive nature of the procedure. In addition, the choice of diethyl ether as the extracting solvent further constrains the analyst to carry out the extractions soon after hydrolysis because of the development of peroxides in ether over time. These peroxides can efficiently react with oxidation products and drastically alter their distribution. In this paper, we present a new approach to perform CuO oxidations that uses a microwave digestion system and a modified (8) Hedges, J. I.; Ertel, J. R.; Leopold, E. B. Geochim. Cosmochim. Acta 1982, 46, 1869-1877. (9) Wilson, J. O.; Valiela, I.; Swain, T. Mar. Biol. 1985, 90, 129-137. (10) Meyers-Schulte, K. J.; Hedges, J. I. Nature 1986, 321, 61-63. (11) Ziegler, F.; Ko ¨gel, I.; Zech, W. Z. Pflanzenernaehr. Bodenk. 1986, 149, 323331. (12) Moran, M. A.; Pomeroy, L. R.; Sheppard, E. S.; Atkinson, L. P.; Hodson, R. E. Limnol. Oceanogr. 1991, 36, 1134-1149. (13) Requejo, A. G.; Brown, J. S.; Boehm, P. D.; Sauer, T. C. Org. Geochem. 1991, 17, 649-662. (14) Alberts, J. J.; Filip, Z.; Price, M. T.; Hedges, J. I.; Jacobsen, T. R. Org. Geochem. 1992, 18, 171-180. (15) Prahl, F. G.; Ertel, J. R.; Gon ˜i, M. A.; Sparrow, M. A.; Eversmeyer, B. Geochim. Cosmochim. Acta 1994, 58, 3035-3048. (16) Opsahl, S.; Benner, R. Nature 1997, 386, 480-482. (17) Bergamaschi, B. A.; Tsamakis, E.; Keil, R. G.; Eglinton, T. I.; Montlucon, D. B.; Hedges, J. I. Geochim. Cosmochim. Acta 1997, 61, 1247-1260. (18) Gon ˜i, M. A.; Hedges, J. I. Geochim. Cosmochim. Acta 1992, 56, 40254043. (19) Gon ˜i, M. A.; Hedges, J. I. Geochim. Cosmochim. Acta 1990, 54, 30833093. (20) Gough, M. A.; Fauzi, R.; Mantoura, C.; Preston, M. Geochim. Cosmochim. Acta 1993, 57, 945-964. (21) Opsahl, S.; Benner, R. Geochim. Cosmochim. Acta 1995, 59, 4889-4904. (22) Gon ˜i, M. A.; Hedges, J. I. Geochim. Cosmochim. Acta 1995, 59, 29652981. (23) Gon ˜i, M. A.; Ruttenberg, K. C.; Eglinton, T. I. Geochim. Cosmochim. Acta 1998, 62, 3055-3075. (24) Gon ˜i, M. A.; Eglinton, T. I. Org. Geochem. 1996, 24, 601-615. (25) Gon ˜i, M. A.; Ruttenberg, K. C.; Eglinton, T. I. Nature 1997, 389, 275-278. 10.1021/ac991316w CCC: $19.00
© 2000 American Chemical Society Published on Web 05/31/2000
Figure 2. Graph of the temperature and pressure conditions inside the microwave Teflon reaction vessels during an alkaline CuO oxidation run.
Figure 1. Sketch of the frontal view of the microwave digestion system (CEM Corp.). For simplicity, only three reaction vessels are depicted inside the microwave cavity. The electronic controls (temperature and pressure) are depicted below the body of the microwave. All lines connecting the reaction vessels, pump, and pressure transducers are Teflon.
extraction procedure, which result in a marked increase in sample throughput. The temperature and duration of the reaction are optimized to yield results comparable to those in the large body of literature devoted to the “traditional” method. Finally, this procedure represents a novel hydrolysis application of microwave technology, which has been utilized prevalently to conduct acid digestions. The work presented here demonstrates that the potential problem of arching (i.e., sparking) can be avoided at moderately low (2 N) base concentrations. EXPERIMENTAL SECTION Apparatus. Oxidations were performed using a CEM MDS 2000 microwave digestion system fitted with up to 12 all-Teflon PFA reaction vessels (CEM) designed for liquid-phase hydrolysis reactions (Figure 1). The reaction vessels, which are rated to 260 °C and 100 psi, are sealed using Teflon O-rings and connected with Teflon tubing. The temperature of the reaction is monitored via a Thermo-Optic (CEM) fiber-optic probe that is placed inside one of the reaction vessels. Pressure is measured with a transducer that is connected to another reaction vessel via the pressure/vacuum line. Both temperature and pressure are continuously monitored and recorded. The directly measured temperature (and/or pressure) is used to regulate the microwave power output and maintain a precise, user-selected temperature value. The pressure/vacuum line is connected via a valve to both a pump and tank of nitrogen gas. By switching back and forth between the pump and gas tank, it is possible to evacuate the air in the reaction vessel headspace and replace it with a blanket of inert gas before starting the reaction. Procedure. Prior to the oxidation, a freshly prepared 2 N NaOH solution is bubbled overnight with N2 gas to remove
dissolved oxygen in the water. This is important, as most samples are sensitive to the presence of O2 in the reaction solution. A known amount of sample, representing 2-5 mg of organic carbon, is loaded into each vessel along with 500 mg of fine CuO powder and 50 mg of ferrous ammonium sulfate, the latter used to bind any remaining oxygen. After the addition of ∼15 mL of the bubbled 2 N NaOH solution to each sample, each reaction vessel is capped using an automatic capping station (CEM). The closed reaction vessels are placed in the rotating tray, connected to each other via the Teflon tubing, and loaded into the microwave. The fiber-optic temperature probe is connected to the first reaction vessel while the pressure/vacuum line is connected to the last reaction vessel. Once the vessels are in place, a leak check is conducted by pressurizing the whole system to ∼60 psi with N2 gas and monitoring the pressure after it is isolated by closing the pressure/vacuum valve. After the leak check, the whole system is purged several times by alternatively evacuating the headspace in the reaction vessels with a vacuum pump and replacing it with a blanket of N2 gas. Prior to the start of the reaction, a slight positive N2 pressure (∼10 psi) is attained. According to our experiments (see Results and Discussion), the optimal oxidation temperature for comparisons with previous results is 150 °C and the reaction time is 90 min. When six reaction vessels are loaded in the microwave, this reaction temperature is reached within 5 min and maintained for the duration of the procedure at pressures of ∼ 60-70 psi (Figure 2). Because longer times are needed to achieve 150 °C when all 12 vessels are used, we recommend running six samples at a time to ensure reproducible reaction conditions. Analyses of 12 samples per day are easily achieved by carrying out a second oxidation while the postreaction extraction and drying steps are performed for the first six samples. Once the oxidation reactions are complete, the reaction vessels are allowed to cool, removed from the microwave, and opened using the capping station. At this point, known amounts of recovery standards (ethylvanillin and trans-cinnamic acid) are added to each vessel. The contents of each vessel are then transferred to Pyrex centrifuge tubes, which are centrifuged at 3000 rpm for 10 min to separate the solids (sample + CuO) from the hydrolysate. The supernates are decanted into 50 mL Pyrex tubes fitted with Teflon-coated caps. This step is repeated once after the addition of ∼5 mL of 1 N NaOH to each centrifuge tube Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
3117
to ensure efficient transfer of the supernate. The alkaline solutions are acidified to pH 1 by addition of ∼4 mL of concentrated HCl to each hydrolysate. Following acidification, a known volume (6-8 mL) of ethyl acetate is added to each tube. Extraction of the aqueous phase is achieved by thoroughly shaking the closed centrifuge tube and allowing the two phases to separate. The organic phase is carefully transferred to a second tube using a precleaned glass pipet. This step is repeated once more to maximize recovery of the organic extract. At this point, the tubes can be closed with Teflon-lined caps and stored in a refrigerator until further processing. Any excess water present in the ethyl acetate extract is removed by addition of cleaned Na2SO4. The ethyl acetate solution is transferred quantitatively to cleaned glass test tubes that fit into a Zymark Turbo Vap evaporation station. The Turbo Vap contains a water bath and 50 individual gas outlets for automated solvent evaporation. The ethyl acetate in each tube is evaporated to dryness under a constant N2 gas stream (5 psi head pressure) in a ∼45 °C water bath. Under these conditions, the ∼12 mL quantities of ethyl acetate evaporate in 50 min. Immediately after the solvent is evaporated, the samples are redissolved in ∼400 µL of pyridine and the solutions are quantitatively transferred to amber glass vials, which are capped and stored in a freezer until the samples are needed for gas chromatographic analyses. Because of the ability to store extracts in ethyl acetate without measurable compositional changes, we typically perform the drying steps after several CuO oxidation runs to maximize the capacity of the Turbo Vap system. Gas chromatographic analyses are performed after transferring portions of the pyridine extracts (typically 50 µL) to autosampler GC vials. Internal standard compounds (e.g., 3,4-dihydroxybenzoic acid) can be added to the vials at this point. Prior to injection of a sample into the gas chromatograph, an excess volume (e.g., 50 µL) of BSTFA + 1% TCMS is added to the vial, which is crimpsealed with a Teflon-lined cap, and the contents are allowed to react on a hot plate at 60 °C for 10 min in order to silylate any exchangeable hydrogens present in the extract. Trimethylsilyl ether and esters of CuO oxidation products are analyzed by gas chromatography using a 60 m × 0.320 mm (i.d.) J&W DB-1 column (0.25 µm film thickness) fitted to a flame ionization detector. Chromatographic separation is achieved using a temperature program of 100 °C initial temperature, 4 °C/min temperature ramp, a final temperature of 320 °C, and a final hold of 10 min. In our laboratory, we routinely use several gas chromatographs fitted with cool on-column and/or splitless injectors, as well as constant column flow and/or constant column pressure conditions. The data presented in this study were collected with a Hewlett-Packard 6890 gas chromatograph fitted with an oncolumn injector and used in a constant flow (1.5 mL/min) mode. The data from the gas chromatograph are acquired and analyzed using a Chrom Perfect (Justice Innovations) chromatographic data system. The concentrations of individual lignin phenols are calculated using the response factors of commercially available standards, which are injected periodically, and reported in mg/ 100 mg of organic carbon in the sample. Gas chromatography/ mass spectrometry is used to verify the identity of individual compounds. 3118
Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
RESULTS AND DISCUSSION Extraction Solvent. One of the important modifications of this new procedure is the choice of ethyl acetate over diethyl ether as the extraction solvent. A major reason for this change is the problem associated with the buildup of peroxides in ether and their destructive attack on aromatic compounds stored in this solvent. As a result, with the “traditional” CuO method, it is necessary to produce freshly distilled ether each time an oxidation is performed and it is imperative to avoid storage of extracts in ether for any extended period of time. Both of these requirements increase the overall duration of the procedure and significantly reduce the flexibility of the analyst. With the use of ethyl acetate, these problems are resolved, since ethyl acetate is commercially available in high-purity grade (GC2), thus eliminating the need for solvent distillation, and can be used to store aromatic products without alteration. We compared the extraction efficiencies of ethyl acetate and ethyl ether to ensure that the change in solvent would not negatively affect the results of the oxidation. To perform this comparison, 2 N NaOH solutions containing known amounts of the 13 standard compounds listed in Table 1 were acidified and extracted with similar volumes of either ethyl acetate or diethyl ether according to the procedure outlined above. The organic solvent extracts were handled exactly as previously described except that a known amount of p-hydroxybenzoic acid was added to each vial just prior to derivatization with BSTFA and GC analyses and used as an internal standard for absolute quantification. These experiments show that slightly higher recoveries for syringyl (dimethoxyl) phenols were obtained when ethyl acetate was used instead of ethyl ether. Such a trend may lead to somewhat higher syringyl phenol yields in samples analyzed using this new procedure. Overall, however, our experiments show that ethyl acetate has extraction efficiencies similar to those of diethyl ether for all the lignin-derived compounds routinely quantified by the oxidation procedure. Hence, ethyl acetate offers the possibility to carry out the drying steps in batches at a significant saving of the analyst’s time while at the same time yielding results comparable to those previously produced using ethyl ether. Oxidation Conditions. The method originally developed by Hedges and Ertel6 uses stainless steel reaction vessels (minibombs) that are fitted into a large Parr bomb. This apparatus is placed into a thermal sleeve and heated for 3 h.6 Using this setup, it is impossible to measure the temperature of the NaOH solution inside the minibombs, but optimal lignin phenol recoveries are obtained when the temperature inside the large Parr bomb is set to 155 °C.18 Because of the large thermal mass of the reaction vessels, it usually takes 30 min to achieve the reaction temperature in the Parr bombs, probably longer inside the minibombs. Under these reaction conditions, phenols with aldehyde, ketone, and acid groups are obtained as the oxidation products of lignin building blocks with three carbon side chains present in the original polymer. We used four previously characterized samples to optimize the reaction conditions in the microwave so as to produce results comparable to those presented in the literature. The samples include a recent sediment sample from Lake Washington26 in Seattle, WA, and a ∼40 000-year-old marine sediment from the (26) Hedges, J. I. Geophys. Monogr. 1991, 63, 129-137.
Table 1. Percentages of Lignin Phenols Recovereda Using Different Solvents compound name
code
ethyl acetate
diethyl ether
Recovery Standards CnAd EVl
72.9 ( 2.7 71.3 ( 5.8
78.1 ( 4.2 76.5 ( 5.3
vanillin (3-methoxy-4-hydroxybenzaldehyde) acetovanillone (3-methoxy-4-hydroxyacetophenone) vanillic acid (3-methoxy-4-hydroxybenzoic acid)
Vanillyl Phenols Vl Vn Vd
68.5 ( 6.3 75.7 ( 4.0 85.0 ( 3.3
68.9 ( 5.6 75.7 ( 4.6 81.3 ( 3.1
syringealdehyde (3,5-dimethoxy-4-hydroxybenzaldehyde) acetosyringone (3,5-dimethoxy-4-hydroxyacetophenone) syringic acid (3,5-dimethoxy-4-hydroxybenzoic acid)
Syringyl Phenols Sl Sn Sd
80.0 ( 4.0 82.0 ( 4.5 84.0 ( 3.9
68.3 ( 2.9 68.8 ( 3.1 73.3 ( 1.8
87.0 ( 3.1 77.6 ( 3.8
86.8 ( 1.4 77.0 ( 3.2
45.1 ( 12 86.4 ( 4.3 82.2 ( 2.9
57.3 ( 10 83.4 ( 2.8 68.8 ( 1.9
cinnamic acid (trans-phenylacrylic acid) ethylvanillin (3-ethoxy-4-hydroxybenzaldehyde)
Cinnamyl Phenols p-coumaric acid (trans-(4-hydroxyphenyl)acrylic acid) pCd ferulic acid (trans-(3-methoxy-4-hydroxyphenyl)acrylic acid) Fd Benzoic Acids benzoic acid m-hydroxybenzoic acid 3,5-dihydroxybenzoic acid
Bd mBd 3,5-Bd
a Recoveries were calculated for 2 N NaOH solutions containing ∼1.3 ng/µL concentrations of each commercially available compound. The solutions were acidified to pH 1 with concentrated HCl and extracted according to the CuO oxidation procedure. Tabulated data are averages ( standard deviations from measurements performed in triplicate.
Table 2. Comparison of Lignin CuO Oxidation Data Obtained Using the Microwave Procedure vs the Traditional Published Method Lake Washington sediments parameter
microwave (n ) 6)
publisheda (n ) 8)
Amazon fan sediments microwave (n ) 4)
Amazon wood
publishedb (n ) 1)d
microwave (n ) 5)
oak leaves
publishedc (n ) 1)d
microwave (n ) 4)
publishedc (n ) 2)
Lignin Phenol Yields (mg/100 mg of Organic Carbon) Vl Vn Vd total V
0.82 ( 0.08 0.26 ( 0.02 0.39 ( 0.03 1.47 ( 0.10
0.82 ( 0.04 0.27 ( 0.01 0.34 ( 0.02 1.44 ( 0.04
Vanillyl Phenols (V) 0.87 ( 0.06 0.91 ( 0.09 0.30 ( 0.02 0.28 ( 0.04 0.38 ( 0.04 0.28 ( 0.04 1.55 ( 0.09 1.47 ( 0.15
6.53 ( 0.18 1.15 ( 0.07 1.74 ( 0.43 9.42 ( 0.46
5.24 ( 0.52 0.89 ( 0.13 1.00 ( 0.15 7.13 ( 0.71
1.39 ( 0.08 0.29 ( 0.01 0.35 ( 0.16 2.03 ( 0.20
1.77 ( 0.18 0.42 ( 0.06 0.35 ( 0.05 2.54 ( 0.25
Sl Sn Sd total S
0.34 ( 0.03 0.24 ( 0.07 0.14 ( 0.05 0.71 ( 0.10
0.35 ( 0.02 0.14 ( 0.01 0.12 ( 0.01 0.61 ( 0.03
Syringyl Phenols (S) 0.66 ( 0.04 0.79 ( 0.04 10.05 ( 0.86 0.28 ( 0.04 0.23 ( 0.03 2.73 ( 0.28 0.15 ( 0.10 0.23 ( 0.02 1.82 ( 0.52 1.08 ( 0.15 1.25 ( 0.13 14.60 ( 1.21
8.70 ( 0.44 1.71 ( 0.26 1.51 ( 0.15 11.90 ( 1.19
1.39 ( 0.08 0.45 ( 0.15 0.37 ( 0.11 2.22 ( 0.11
1.65 ( 0.08 0.48 ( 0.07 0.32 ( 0.03 2.45 ( 0.25
pCd Fd total C
0.16 ( 0.02 0.05 ( 0.04 0.21 ( 0.03
0.11 ( 0.01 0.07 ( 0.01 0.19 ( 0.01
Cinnamyl Phenols (C) 0.09 ( 0.01 0.08 ( 0.01 0.03 ( 0.02 0.05 ( 0.01 0.13 ( 0.03 0.13 ( 0.02
0.01 ( 0.01 0.07 ( 0.05 0.09 ( 0.04
0.01 ( 0.00 0.08 ( 0.01 0.09 ( 0.01
0.39 ( 0.07 0.12 ( 0.03 0.51 ( 0.07
0.43 ( 0.06 0.24 ( 0.04 0.67 ( 0.10
total lignin (Λ)
2.40 ( 0.13
2.23 ( 0.05
2.76 ( 0.24
24.11 ( 1.19
19.12 ( 1.91
4.75 ( 0.19
5.66 ( 0.57
0.42 ( 0.02 0.13 ( 0.01 0.42 ( 0.03 0.34 ( 0.03
Lignin Parameters 0.70 ( 0.08 0.85 ( 0.09 0.08 ( 0.02 0.08 ( 0.01 0.43 ( 0.05 0.31 ( 0.05 0.22 ( 0.14 0.29 ( 0.04
1.55 ( 0.16 0.01 ( 0.00 0.27 ( 0.07 0.18 ( 0.04
1.67 ( 0.17 0.01 ( 0.00 0.19 ( 0.03 0.17 ( 0.03
1.10 ( 0.14 0.25 ( 0.06 0.25 ( 0.12 0.27 ( 0.08
0.96 ( 0.10 0.26 ( 0.04 0.20 ( 0.03 0.19 ( 0.03
S/V C/V [Ad/Al]ve [Ad/Al]sf
0.49 ( 0.09 0.15 ( 0.03 0.47 ( 0.05 0.41 ( 0.13
2.85 ( 0.28
a Data produced by Hedges’ laboratory for interlaboratory comparisons. b Data from ref 27. c Data from ref 18. d For samples with n ) 1, standard deviations were based on the average analytical reproducibility of each individual lignin CuO reaction product. e Vanillic acid/vanillin ratio. f Syringic acid/syringealdehyde ratio.
Amazon deep sea fan27 off the coast of Brazil. In addition, we analyzed samples of vascular plant tissues, including a tropical hardwood (Zanthoxylum compactum) from the Amazon River drainage basin28 and green leaves from northern red oak (Quercus rubra) collected in the Seattle, WA, arboretum.18 The primary variables investigated were the temperature and duration of the oxidation. We sought to minimize the reaction
time and, after several experiments, ascertained an optimal reaction temperature of 150 °C for 90 min (Table 2). The data in Table 2 show extremely good agreement between the “traditional” method and the new microwave technique. Overall, both techniques show excellent agreement in the lignin phenol yields and compositional parameters (i.e., S/V, C/V, and [Ad/Al] ratios) for the two sediment samples analyzed. Furthermore, the lignin yields
(27) Gon ˜i, M. A. Proceedings of the Ocean Drilling Program. Scientific Results; Ocean Drilling Program, College Station, TX, 1997; Vol. 155, pp 519-530.
(28) Hedges, J. I.; Clark, W. A.; Quay, P. D.; Richey, J. E.; Devol, A. H.; Santos, U. d. M. Limnol. Oceanogr. 1986, 31, 717-738.
Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
3119
Figure 3. Plots of lignin phenol yields obtained from the alkaline CuO oxidation analyses of four different sample types at different reaction temperatures using the microwave digestion system. The absence of Sd/Sl data for the Amazon fan sediment at 140 °C is due to the belowdetection yields of syringyl phenols obtained at this temperature. Error bars represent standard deviations of replicate analyses.
and compositions for the oak leaves obtained by the microwave technique are also statistically similar (P ) 0.95; t test) to those produced by the traditional technique. The microwave and traditional methods, on the other hand, give significantly different yields of lignin phenols from the Amazon wood. The higher recovery of lignin phenols by the microwave technique is in part due to the enhanced efficiency for extraction of syringyl phenols by ethyl acetate. However, the solvent effect alone does not explain the differences for this sample. It may be the case that microwave energy is more effective in the cleavage of ether and carbon bonds in a polysaccharide-rich matrix such as wood. Despite these differences in absolute yields, both approaches give comparable ratios of lignin phenols that characterize this sample. Overall, these data show that the microwave-based procedure favorably reproduces the results obtained by the traditional method.6 The effects of reaction temperature on the yields and parameters from the four samples are illustrated in Figure 3. It is clear that the type of sample matrix plays a critical role in the recovery of lignin phenols at different temperatures. For example, while the total lignin yields do not change markedly with increasing temperature in the case of Lake Washington sediments and oak leaves, the yields obtained from Amazon fan sediments and Amazon wood are highly diminished at low temperatures. For example, the syringyl phenol yields obtained from the oxidation of Amazon fan sediments at 140 °C are below detection limits. Most samples display similar trends of increasing acid/aldehyde ratios with increasing temperature. The character of the CuO oxidation reaction helps to explain these trends. The alkaline CuO 3120
Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
reaction is a relatively mild oxidation that involves hydrolysis and one-electron-transfer steps.29 During the CuO procedure, the primary reaction is the oxidative depolymerization of the lignin macromolecule into primary phenolic products (i.e., aldehydes, ketones, and acids). However, as additional energy is provided at elevated temperatures, the oxidation of primary products into secondary ones can become more important and alter the resulting phenolic compositions. According to this explanation, the observed increases in acid/aldehyde ratios with temperature may be due to the oxidation of aldehyde and ketone products into acids. In addition, the high acid/aldehyde ratios obtained from Amazon fan sediments and oak leaves at low temperatures (140 °C) are probably due to the fact that both of these samples contain esterbound vanillic acid.21,30 At these low temperatures, the oxidative hydrolysis of lignin is not very efficient, and therefore, ester-bound vanillic acid dominates the phenol yields. These data indicate that the reaction conditions of the microwave procedure (150 °C, 90 min) are optimal because they result in high lignin phenol yields while at the same time minimize secondary reactions. CONCLUSIONS A newly developed technique to perform alkaline CuO oxidations incorporates the use of a microwave digestion system to hydrolyze lignin phenols from macromolecular precursors in complex geochemical samples. In contrast to the original proce(29) Chang, H. M.; Allan, G. G. Lignins; Sarkaren, K. W., Ludwig, C. H., Eds.; Wiley-Interscience: New York, 1971; pp 433-485. (30) Opsahl, S.; Benner, R. Mar. Ecol.: Prog. Ser. 1993, 94, 191-205.
dure,6 the new method uses ethyl acetate as the extracting solvent and a simplified extraction procedure to recover lignin phenols. At the optimal reaction temperature (150 °C) and time (90 min), this novel approach satisfactorily reproduces the results of the traditional method6 while significantly increasing the overall sample throughput. ACKNOWLEDGMENT Funding for this work was provided to M. Gon ˜i through an NSF grant (OCE-9701940) and to S. Montgomery through a Funds
FCAR research internship. This paper benefited from comments by E. Gordon, M. Teixeira, T. Kastner, H. Aceves, and two anonymous reviewers.
Received for review November 16, 1999. Accepted March 30, 2000.
AC991316W
Analytical Chemistry, Vol. 72, No. 14, July 15, 2000
3121
