Ken-ichi Kuroda a,, Akiko Nakagawa-izumi b, Bibhuti B. Mazumder c, Yoshito Ohtani c, Kazuhiko Sameshima c. 1. Introduction

Industrial Crops and Products 22 (2005) 223 232 Evaluation of chemical composition of the core and bast lignins of variety Chinpi-3 kenaf (Hibiscus cannabinus L.) by pyrolysis gas chromatography/mass spectrometry and cupric oxide oxidation Ken-ichi Kuroda a,, Akiko Nakagawa-izumi b, Bibhuti B. Mazumder c, Yoshito Ohtani c, Kazuhiko Sameshima c a Department of Forest and Forest Products Sciences, Faculty of Agriculture, Kyushu University, Fukuoka 812-8581, Japan b Institute of Agricultural and Forest Engineering, University of Tsukuba, Tsukuba, Ibaraki 305-8572, Japan c Faculty of Agriculture, Kochi University, Nankoku, Kochi 783-8502, Japan Received 19 August 2004; accepted 6 January 2005 Abstract In order to clarify the chemical composition of in situ lignin of kenaf (Hibiscus cannabinus L.), variety Chinpi-3, the core and bast fibers were fractionated from the 6-month maturated kenaf with small stalks (309.0 cm height 1.3 cm diameter, lignin content 16.6%) and large stalks (390.0 cm height 2.3 cm diameter, lignin content 18.8%). The samples were subjected to conventional pyrolysis gas chromatography/mass spectrometry (500 C/4 s), and cupric oxide oxidation. The quantitative pyrolysis results showed that (1) the differences in stalk size, when harvested at the same maturity, do not influence the lignin composition, (2) in situ core lignin is a mixed lignin comprising 1.4 parts of syringyl-, 1 part of guaiacyl-, and a small part of p-hydroxyphenyl lignin units, and (3) in in situ bast fiber lignin the syringyl lignin units are present in greater quantities than the guaiacyl lignin units. The cupric oxide oxidation results also supported the pyrolysis results. 2005 Elsevier B.V. All rights reserved. Keywords: Pyrolysis gas chromatography/mass spectrometry; Kenaf (Hibiscus cannabinus L.); In situ lignin composition; Syringyl lignin units; Guaiacyl lignin units; Stalk dimensions; Cupric oxide oxidation 1. Introduction A part of this study was presented in the 2nd American Kenaf Society Meeting, San Antonio, TX, USA, February 25 26, 1999. Corresponding author. Tel.: +81 92 642 2988; fax: +81 92 642 2988. E-mail address: kenfumi@agr.kyushu-u.ac.jp (K.-I. Kuroda). Kenaf (Hibiscus cannabinus L.) is a dicotyledonous annual plant grown in temperate and tropical areas with a high fiber yield, consisting of an inner core fiber (75 60%) and an outer bast fiber (25 40%). It has the 0926-6690/$ see front matter 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.indcrop.2005.01.002

224 K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 excellent advantages of being renewable, inexpensive, and is easily grown even under severe conditions such as low water supply and few fertilizers. With a decrease in wood resources, interest has grown in using kenaf as an alternative raw material for wood-based materials such as pulps (Aorigele and Sano, 1997; Kaldor, 1989, 1992; Karakus et al., 2001; Mazumder et al., 1998; Romanoschi et al., 1997), although no high added-value applications have been found for kenaf so far. The chemical compositions and abundances of lignin in lignocellulosic materials greatly influence both pulp yield and ease of delignification which are important issues for the papermaking industries. In particular, lignocellulosic materials with high proportion of the syringyl lignin units are advantageous for delignification during pulping and bleaching processes (Collins et al., 1990; Fergus and Goring, 1969; Rodrigues et al., 1999). An understanding of the chemical composition of kenaf lignin therefore gives insight into the application of kenaf as a raw material for pulp. Results are published on the chemical properties of kenaf lignin (Clark and Wolff, 1969; del Rio et al., 2004; Gutierrez et al., 2004; Kuroda et al., 1999, 2002a,b; Mazumder et al., 2005; Morrison III et al., 1999a; Neto et al., 1996; Nishimura et al., 2002; Pappas et al., 1998; Ralph, 1996; Ralph and Lu, 1998; Seca et al., 1997, 1998). Neto et al. (1996) and Seca et al. (1997, 1998) showed that large differences in chemical composition exist between in situ and isolated lignins of kenaf. That is, both the core and bast fibers in situ lignins are rich in the guaiacyl lignin units, while the lignins isolated from the core and bast fibers are rich in the syringyl lignin units. Such discrepancies between both in situ and isolated lignins are unlike the natural variation of the chemical composition due to the differences in varieties and morphological regions (Morrison III et al., 1999a,b; Nishimura et al., 2002). Variations resulting from isolation procedures, if so, are artifacts and can lead to an erroneous selection of kenaf for kenaf-based materials. Compositional information of lignins in general is available by wet chemical and spectroscopic methods; most of them involve multi-step sample pretreatments which may have a risk to modify the polymeric structures significantly. Therefore, it is often hard to draw in situ structural information on lignin from lignocellulosic materials by wet chemical and spectroscopic methods. The goal of this study was to obtain structural information on kenaf in situ lignin. For this, pyrolysis gas chromatography/mass spectrometry (pyrolysis GC/MS) and cupric oxide oxidation were used. The former is a method of resolving problems such as those which occurred between in situ and isolated lignins because it can directly measure the chemical composition of lignin; therefore, it provides minor modifications (if at all) to the sample structures. This method has been successfully used to analyze lignocellulosic materials and to draw in situ structural information from them (Meier and Faix, 1992; Ralph and Hatfield, 1991). Cupric oxide oxidation is a convenient method capable of obtaining the chemical composition of lignin, as well as alkaline nitrobenzene oxidation: this method is prefer to nitrobenzene oxidation because of its milder and more selective degrading functions (Chang and Allan, 1971). With pyrolysis in the presence of tetramethylammonium hydroxide (TMAH thermochemolysis) (Kuroda et al., 2002a,b), and chemical degradation methods (Nishimura et al., 2002), we showed greatly different results on kenaf (variety Chinpi-3) in situ lignin composition from those by Neto et al. (1996) and Seca et al. (1997, 1998), who analyzed a different variety (Salvador) with different methods from our methods. The pyrolysis GC/MS and cupric oxide oxidation results obtained here will confirm and complement our previous results (Kuroda et al., 2002a; Nishimura et al., 2002). 2. Materials and methods 2.1. Kenaf samples A kenaf cultivar, variety Chinpi-3, was planted in the experimental field (100 m 2 ) of the Faculty of Agriculture, Kochi University, Japan, in late-april 1993, and harvested in mid-november 1993. The harvested kenaf fibers were manually separated from foliage and divided into three groups according to the size of the kenaf stalk: small, medium and large size kenafs. Immediately after harvesting, they were air-dried and cut in three parts (top, middle, and bottom) with

K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 225 the roughly same length. The middle parts of the kenaf stalks were further hand-separated to provide the core and bast fiber samples each. Each sample was ground. Materials finer than 60 mesh were Soxhletextracted with ethanol-benzene (1:2, v/v) for 6 h. After air-dried, the extractive-free small and large size kenafs were subjected to the lignin determination, cupric oxide oxidation, and pyrolysis GC/MS analysis. Until analyzed, the samples were stored in a dark room. 2.2. Lignin determination The lignin content was determined by the 72% sulfuric acid method according to the Tappi standard rule T222 om-8 (TAPPI, 1988). 2.3. Cupric oxide oxidation Cupric oxide oxidations were carried out according to the method described in the previous paper (Kuroda et al., 2002a). Relative standard deviation was 3.5% for duplicate runs. 2.4. Pyrolysis GC/MS The pyrolysis GC/MS system was a combination of a JHP-3 model Curie-point pyrolyzer (Japan Analytical Industry, Tokyo, Japan) and an HP 5890 series II gas chromatograph (Hewlett-Packard, Palo Alto, CA, USA) with an HP 5972A quadrupole mass selective detector (Hewlett-Packard). The kenaf meals ( 200 300 g) were pyrolyzed at 500 C for 4 s. The volatile products were sent to the GC/MS. The pyrolysis GC/MS conditions were the same as those employed in the previous paper (Mazumder et al., 2005): a Quadrex MS fused-silica capillary column (25 m 0.25 mm i.d., film thickness 0.25 m), temperature program of 50 C (1 min) to 300 C ramped at 5 C/min, carrier gas He (flow rate 1.3 ml/min), temperature of injection and detector ports 280 C, 70 ev electron impact voltage, and mass range m/z 50 600. Spectra were collected by an HP ChemStation software package. Peak assignments were carried out on the basis of mass fragmentation patterns, and by comparing the MS data with published data (Ralph and Hatfield, 1991). Relative standard deviation was 4.0% for duplicate runs. 3. Results and discussion 3.1. Sample description After a 6-month maturity, 315 stalks were harvested and grouped into the small (108 stalks, 34.5%), medium (175 stalks, 55.5%), and large (32 stalks, 10.0%) size kenafs according to the stalk dimension. The plants varied in height from 3 to 4 m, and in diameter from 1.3 to 2.3 cm. We expected that the differences in stalk dimension influence the chemical composition, as well as those in harvested time and development of grown stage whose effects are typically observed on the stalk dimensions and chemical compositions (Karakus et al., 2001; Mambelli and Grandi, 1995; Mazumder et al., 2005; Morrison III et al., 1999a,b). Average stalk dimensions (height basal diameter) were 309.0 cm 1.23 cm, 391.0 cm 1.78 cm, and 390.5 cm 2.3 cm for the small, middle, and large size kenafs, respectively. The differences in fiber length, its distribution, and pulp yield among these different size kenafs are reported by Zhou et al. (1995a,b) The small and large size kenafs were selected for this study. The small size kenafs contained a smaller amount of lignin than the large size kenafs: 16.6% versus 18.8%. The average bast/core ratios were 0.66 for the small size kenafs and 0.54 for the large size kenafs. The small size kenaf core and bast fibers contained 20.9 and 10.1% lignin, respectively, and the large size kenaf core and bast fibers contained 22.3 and 12.3% lignin, respectively. 3.2. Pyrolysis GC/MS characterization of the kenaf core fibers Fig. 1a shows the pyrogram of the core fiber fractionated from the small size kenaf; the pyrograms of the core and bast fiber fractions of the large size kenaf are shown in a previous paper (Kuroda et al., 1999). The identified products are listed in Table 1 with the relative abundances (%) and pyrolytic sources. The GC/MS signal areas were integrated to derive the relative abundances of the individual compounds (total GC/MS signal area equals 100) in order to determine the relative abundance of the product. The products are classified as carbohydrate-derived products, products possessing p-hydroxyphenyl- (H), guaiacyl- (G), and syringyl- (S) nuclei, and miscellaneous products (M). The chemical structures of the phenols are shown in

226 K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 Fig. 1. Pyrolysis GC/MS traces of (a) core fibers and (b) bast fibers from small size variety Chinpi-3. Note: Product numbers and names refer to those in Table 1. The structures of the phenols are shown in Fig. 2. ( ) Carbohydrate-derived products, ( ) p-hydroxyphenyl lignin-derived products, ( ) guaiacyl lignin-derived products, ( ) syringyl lignin-derived products, ( ) miscellaneous products. Fig. 2. The differences in stalk dimension produce no large qualitative differences in pyrolysis product profile of the core fiber, but produce quantitative differences (see Table 1). Fig. 1a reveals many S type products. The major S type products observed are 2,6-dimethoxy- 4-vinylphenol (29, 9.3%) and 2,6-dimethoxyphenol (20, 8.3%) followed by (E)-2,6-dimethoxy-4- propenylphenol (37, 4.8%). Sinapaldehyde (45), syringaldehyde (32), 2,6-dimethoxy-4-methylphenol (24), and (E)-sinapyl alcohol (46) are present in about 2.7 3.4% relative abundance each (order of abundance). The relative abundance of 2,6-dimethoxy- 4-ethylphenol (28), propiosyringone (42) and 2,6- dimethoxy-4-allylphenol (30) each is about 1.4 1.6% (order of abundance). Other 2,6-dimethoxyphenols 33 and 43 are found in a relative abundance of <1%. Acetosyringone (39, 1.69%) co-eluted with coniferaldehyde (38, 2.10%). We tentatively

K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 227 Table 1 Assignment of the pyrolysis GC/MS peaks of kenaf Peak Product Relative abundance (%) a Source b Small size kenaf Large size kenaf Core Bast Core Bast 1 Furfural 3.00 5.68 2.50 4.55 C 2 2-Hydroxymethylfuran 1.06 2.93 0.69 1.34 C 3 (5H)-Furan-2-one 1.56 2.84 0.56 n.d. C 4 5-Methyl-3-hydrofuran-2-one 2.63 5.18 1.11 2.40 C 5 5-Methylfuran-2-carbaldehyde 1.38 2.72 0.66 n.d. C 6 4-Hydroxy-5,6-dihydro-2H-pyran-2-one 3.73 3.93 1.38 1.51 C 7 Phenol 0.82 1.20 0.45 1.19 H 8 2-Hydroxy-3-methylcyclopent-2-en-1-one 1.22 3.82 0.73 3.41 C 9 Methylphenol n.d. c n.d. 0.36 1.07 H 10 Guaiacol 4.69 3.64 4.30 2.97 G 11 3-Hydroxy-2-methyl-4H-pyran-4-one n.d. 1.62 n.d. n.d. C 12 Dimethyldihydropyranone isomer n.d. 1.37 n.d. n.d. C 13 2,3-Dihydroxybenzaldehyde 0.79 n.d. 0.68 1.89 M 14 4-Methylguaiacol 1.76 1.79 1.13 1.43 G 15 Catechol n.d. n.d. n.d. n.d. M 16 4-Vinylphenol 7.66 n.d. 7.66 n.d. M 17 3-Methoxybenzene-1,2-diol 1.08 2.06 1.19 3.00 M 18 4-Ethylguaiacol 0.84 0.80 0.95 1.25 G 19 4-Vinylguaiacol 6.97 2.76 5.63 2.95 G 20 2,6-Dimethoxyphenol 8.31 10.01 7.93 12.25 S 21 Eugenol 1.65 n.d. 0.81 1.13 G 22 Vanillin 1.95 1.16 1.58 0.70 G 23 (Z)-Isoeugenol 0.51 n.d. 0.49 n.d G 24 2,6-Dimethoxy-4-methylphenol 3.15 3.58 2.78 2.87 S 25 (E)-Isoeugenol 3.19 1.67 3.10 1.73 G 26 Acetoguaiacone 0.69 n.d. 0.84 n.d. G 27 Propioguaiacone 1.18 n.d. 0.80 0.91 G 28 2,6-Dimethoxy-4-ethylphenol 1.64 4.10 1.45 1.87 S 29 2,6-Dimethoxy-4-vinylphenol 9.26 10.07 8.57 11.67 S 30 2,6-Dimethoxy-4-allylphenol 1.37 2.97 1.52 8.06 S 31 2,6-Dimethoxy-4-propylphenol n.d. n.d. n.d. n.d. S 32 Syringaldehyde 3.16 2.63 3.09 3.45 S 33 (Z)-2,6-Dimethoxy-4-propenylphenol 0.87 1.51 1.17 2.29 S 34 Dihydroconiferyl alcohol 0.02 n.d. 0.02 n.d. G 35 (Z)-Coniferyl alcohol 0.37 n.d. 2.01 n.d. G 36 Homosyringaldehyde n.d. n.d. 0.46 n.d. S 37 (E)-2,6-Dimethoxy-4-propenylphenol 4.80 9.11 5.19 11.95 S 38 Coniferaldehyde 2.10 n.d. 1.27 n.d. G 39 Acetosyringone 1.69 1.66 1.93 2.78 S 40 (E)-Coniferyl alcohol 5.18 1.54 10.96 2.18 G 41 Syringylacetone 2.07 3.12 1.94 3.51 S 42 Propiosyringone 1.55 0.96 0.40 1.00 S 43 Dihydrosinapyl alcohol n.d. n.d. 1.11 n.d. S 44 (Z)-Sinapyl alcohol n.d. n.d. 1.50 n.d. S 45 Sinapaldehyde 3.38 1.83 2.71 n.d. S 46 (E)-Sinapyl alcohol 2.74 2.12 6.34 n.d. S Total 100.0 100.4 100.0 97.3 a Based on the relative GC/MS peak area (%). b C: carbohydrate, H: p-hydroxyphenyl lignin units, G: guaiacyl lignin units, S: syringyl lignin units, M: miscellaneous. c Not determined or not found.

228 K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 Fig. 2. Chemical structures of the identified phenols. estimated the abundance of 38 to be 1.2 times that of 39 based on the ratio of the molecular ion intensity of 38 (m/z 178) to that of 39 (m/z 196). Of 2-methoxyphenols coming from the G lignin units, the main products are 4-vinylguaiacol (19, 7.0%), (E)-coniferyl alcohol (40, 5.2%), and guaiacol (10, 4.7%) followed by (E)-isoeugenol (25, 3.2%). Compared to these products, 4- methylguaiacol (14), 4-ethylguaiacol (18), vanillin (22), acetoguaiacone (26), and coniferaldehyde (38) are present in a small abundance (0.7 2.1%). 4-Vinylphenol (16, 7.7%) co-eluted with catechol (15); the intensity of the molecular ion of the former was more than 20 times that of the latter. 16 comes from either etherified or esterified p-coumaric acids, similar to 16 in rice pyrolyzates (Kuroda et al., 1995). The support was provided from the TMAH thermochemolysis- GC/MS analysis of the same large size core sample whose pyrogram showed 4-methoxycinnamic acid methyl ester in a considerable abundance and 3,4- dimetoxycinnamic acid methyl ester in a rather small abundance (Kuroda et al., 2002a). Therefore, in the core fiber of variety Chinpi-3 the p-coumaric acid moieties are present in greater abundances than the ferulic acid moieties, similar to in the core fibers of varieties Tainung-1 and Everglades-41 (Morrison III et al., 1999b). This suggests that 19 mostly comes from the lignin, not from the ferulic acid moieties. Catechols 13, 15, 17 are produced in a 1.2% relative abundance. Tannin contained as an impurity may give catechol (15) (van Bergen et al., 1997). Several pyrolytic sources are proposed for 3-methoxybenzene- 1,2-diol (17): the 5-hydroxyguaiacyl moiety (Suzuki et al., 1997), microbial demethylated S lignin units (van Bergen et al., 2000) and benzodioxane structures (Ralph et al., 2001) which is produced by the incorporation of 5-hydroxyconiferyl alcohol. Exact sources for 17 are unclear at present. We grouped 13 and 15 17 to the miscellaneous products (M) based on the above discussion, and 7 and 9 to the H lignin unit-derived products. To assess the chemical composition of kenaf lignin, the S/G ratio was determined based on the relative abundances of the products listed in Table 1. Based on the C:H:G:S:M ratio of about 14.6:0.8:31.1:44.0:9.5 in the core fiber pyrolyzate of the small size kenaf, the relative abundances of the S type products are 1.4 times those of the G type products, showing that in situ core fiber lignin of small size variety Chinpi-3 is a mixed lignin comprising the G and S lignin units with a 1.4 S/G ratio. A similar S/G ratio was obtained with the large size in situ core lignin. 3.3. Pyrolysis GC/MS characterization of the kenaf bast fiber Fig. 1b shows the pyrogram of the bast fiber fractionated from the small size kenaf fiber. No large qualitative differences in products were essentially observed between the bast fibers fractionated from

K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 229 the small and large size kenaf fibers, although quantitative differences were observed (Table 1). The distribution profile of the products of the bast fiber lignin is simple compared to that of the core fiber lignin. Fig. 1b reveals three intense peaks attributable to the S type products: 2,6-dimethoxyphenol (20), 2,6-dimethoxy-4-vinylphenol (29), and 2,6- dimethoxy-4-propenylphenol (37) whose relative abundances are more than 9% each. The less than 4% relative abundances were given by 45, 46, 10, 19 and 40 (order of abundance). The most abundant G type product is 19 ( 2.8%) followed by 25 ( 1.7%). Consequently, the C:H:G:S:M ratio in the small size bast pyrolyzate is 30.1: 1.2:13.4:53.7:2.1, showing that this lignin is of S type with an S/G ratio of 4.0. Similarities were obtained with the large size in situ bast lignin. Unlike the core fiber pyrogram, the pyrogram of the bast fiber lacks 16 (compare Fig. 1b with Fig. 1a). This and the TMAH thermochemolysis-gc/ms results of the large size kenaf bast fiber (Kuroda et al., 2002a) show that the bast fiber of variety Chinpi-3 contains a trace abundance of etherified and/or esterified p-coumaric and ferulic acid moieties. Similarities are observed with the bast fibers of varieties Tainung-2 and Everglades-41 (Morrison III et al., 1999b), and of the kenaf (not specified variety) of del Rio and co-workers (del Rio et al., 2004; Gutierrez et al., 2004). Ralph (1996) and Ralph and Lu (1998) reported that kenaf bast fiber (variety Tainung and other three varieties) lignin contains a large abundance of syringylglycerol- -aryl ether subunits with the -CH 2 OAc group. With pyrolysis GC/MS del Rio and co-workers (del Rio et al., 2004; Gutierrez et al., 2004) also confirmed the presence of both coniferyl and sinapyl alcohol acetates in the kenaf bast pyrolyzate, but in a small level. However, our large and small size kenafs, including the core-, bast-, and whole fractions, did not provide detectable abundances of such acetylated products. This may be due to the differences between the samples employed and/or between the pyrolytic efficiencies of the pyrolysis system employed. 3.4. Cupric oxide oxidation of kenafs Table 2 shows the cupric oxide oxidation results of the kenaf. Responsible for the formation of the products are mainly uncondensed -aryl ether substructures, Table 2 Yields (%) a and molar ratios of cupric oxide oxidation products from variety Chinpi-3 Product Small size Chinpi-3 Large size Chinpi-3 Core Bast Core Bast Vanillin (22) 7.0 2.6 7.7 2.8 Acetoguaiacone 1.6 0.7 1.5 0.7 (26) Syringaldehyde 14.0 18.3 17.5 16.3 (32) Acetosyringone 4.5 3.7 3.7 4.7 (39) Total b 27.1 25.3 30.4 24.5 S/G c 1.79 5.62 1.92 5.06 a Based on Klason lignin. b 22 + 26 + 32 + 39. c Molar ratio of 32 + 39/22 + 26. and parts of -1 and -5 substructures. Therefore, the products stem from several substructures. The yields of vanillin (22) and acetoguaiacone (26) were determined as the parameters of the G lignin units, and those of syringaldehyde (32) and acetosyringone (39) were determined as the parameters of the S lignin units. The summed yield of the products is lower than that of nitrobenzene oxidation products of another variety Chinpi-3 harvested in year 1994 (Ohtani et al., 2001), as in the oxidation of wood samples (Chang and Allan, 1971). However, the observed S/G ratios are enough to show the high proportion of the S lignin units in variety Chinpi-3 in situ lignin: 1.79 and 5.62 S/G ratios for the small size core and bast fibers, respectively, and 1.92 and 5.06 S/G ratios for the large size core and bast fiber lignins, respectively. 3.5. Assessment of lignin composition of variety Chinpi-3 Table 3 summarizes the chemical composition of variety Chinpi-3 in situ lignin determined by pyrolysis GC/MS. The core fibers of the small and large size kenafs provide the same S/G ratio (a 1.4 S/G ratio), showing that variety Chinpi-3 in situ core fiber lignin is a mixed lignin comprising the 1.0 part of the G lignin units and 1.4 part of the S lignin units. The bast fibers of the small and large size kenafs provide the same S/G ratio (a 4.0 S/G ratio), showing that in bast fiber lignin the S lignin units overwhelm the G lignin units.

230 K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 Table 3 Chemical composition of kenaf in situ lignin by different methods Kenaf Relative abundance (%) a S/G ratio by: H G S Conventional pyrolysis b TMAH thermochemolysis b Wet chemical c Chinpi-3 (small size) Core 1.0 41.0 58.0 1.41 1.79 d Bast 1.8 19.4 78.8 4.06 5.62 d Chinpi-3 (large size) Core 1.0 40.9 58.1 1.42 1.53 e 1.92 d Bast 2.9 19.3 77.8 4.04 3.42 e 5.06 d Chinpi-3 f Core n.d. g 44.8 55.2 1.23 1.35 h Bast n.d. g 20.4 79.6 3.89 3.72 h del Rios in situ kenaf i Bast 1.3 15.4 83.3 5.4 Salvador j Core 14 20 57 74 12 23 0.21 0.31 k Bast 9 13 55 60 27 34 0.49 0.57 k a p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin units. b Relative abundance of syringyl type pyrolysis products/that of guaiacyl type pyrolysis products. c Molar ratio of syringyl type products/that of guaiacyl type products. d By cupric oxide oxidation. e From the data of Kuroda et al. (2002b). f From the data of Kuroda et al. (2002b) and Nishimura et al. (2002). g Not determined. h By thioacidolysis (Kuroda et al., 2002b; Nishimura et al., 2002). i From the data of del Rio et al. (2004) and Gutierrez et al. (2004). j From the data of Seca and co-workers (Neto et al., 1996; Seca et al., 1998). k By KMnO 4 oxidation. TMAH thermochemolysis-gc/ms of the large size kenaf sample also supported the predominance of the S lignin units in variety Chinpi-3 lignin (Kuroda et al., 2002a), although it provides different S/G ratios from those determined by conventional pyrolysis GC/MS: 1.53 for the core fiber lignin and 3.42 for the bast fiber lignin. The differences in S/G ratio between the two pyrolysis methods may be due to the differences in that conventional pyrolysis GC/MS provides nonspecific structural information due to the random cleavage of the polymer, and TMAH thermochemolysis-gc/ms prefers to provide structural information on -aryl ether subunits (Challinor, 2001; Filley, 2003; Filley et al., 1999). The proportion of the H lignin units was smaller than that of Netos kenaf (14 20 mol% in core fibers and 9 13 mol% in bast fibers) (Neto et al., 1996; Seca et al., 1998) because we evaluated only 7 and 9 as the products derived from the H lignin units. Table 3 also contains S/G ratios determined by cupric oxide oxidation and thioacidolysis to corroborate the pyrolysis results. Although the S/G ratios obtained by cupric oxide oxidation are about 1.2 1.4 times larger than those by the pyrolysis method because of an increase of the condensed G lignin units during the process (Chang and Allan, 1971), data comparable and complementary to the pyrolysis results are obtained. Thioacidolysis, as well as TMAH thermochemolysis, effectively cleaves -aryl ether linkages (48 and 60% of total inter-unit linkages in spruce and birch lignins, respectively (Adler, 1977)) to provide -aryl ether subunit-derived products amenable to GC/MS analysis (Lapierre, 1993; Rolando et al., 1992). Therefore, it provides information on the chemical composition of the most abundant substructures. Nishimura et al. (2002) applied thioacidolysis to another variety Chinpi-3, which provided 1.35 and 3.72 S/G ra-

K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 231 tios for the upper middle core and bast fibers, respectively. With the same kenaf TMAH thermochemolysis provides very close S/G values to those by thioacidolysis: 1.23 for the core fiber lignin and 3.89 for the bast fiber lignin (Kuroda et al., 2002b). That is, different methodologies draw the same S/G ratios from the same sample. Consequently, the richness of the S lignin units in variety Chinpi-3 in situ lignin was proven both by analytical pyrolysis and wet chemical methods. In summary, variety Chinpi-3 core fiber in situ lignin is a mixed G-S lignin and bast fiber in situ lignin is of S type, similar to the results by other researchers (Abbott and Bagby, 1986; Abbott et al., 1987; Morrison III et al., 1999a,b; Nishimura et al., 2002). Therefore, variety Chinpi-3 lignin is S-rich lignin. The S/G ratios of our kenaf in situ lignin greatly differ from those of variety Salvador in situ lignin determined by permanganate oxidation (see Table 3) (Neto et al., 1996; Seca et al., 1997, 1998), which showed that both core and bast lignins are rich in the G lignin units. This may be due to a variation in inter-plant, namely the difference between varieties Salvador and Chinpi- 3. However, we noted nitrobenzene oxidation results on variety Salvador in situ lignin shown by the same authors (Seca et al., 1997). The nitrobenzene oxidation provides different S/G ratios (2.13 and 3.95 for core and bast in situ lignins, respectively) from those by the permanganate oxidation. The values by the nitrobenzene oxidation are close to our S/G ratios and to those of variety Salvador isolated lignin determined by 13 C NMR spectrometry (Neto et al., 1996; Seca et al., 1998) (1.55 and 3.38 S/G ratios for core and bast dioxane lignins, respectively). Although permanganate oxidation, as well as nitrobenzene oxidation, is a method frequently employed for structural elucidation of lignin, several improved procedures are proposed (Chang and Allan, 1971; Gellerstedt, 1992). The procedures involving no cleavage of -aryl ether linkages often give products in low yields and limit their sources because in such procedures only phenolic substructures (30 35% of the total substructures in a spruce milled wood lignin (Lai and Sarkanen, 1971)) protected against the oxidation by the methylation are responsible for the oxidation products (Chang and Allan, 1971; Gellerstedt, 1992). It is clear that analytical methods employed influenced the S/G ratio determinations of variety Salvador in situ lignin. References Abbott, T.P., Bagby, M.O., 1986. A 14 C balance on nitrobenzene oxidized kenaf lignin. J. Wood Chem. Technol. 6, 487 494. Abbott, T.P., Tjarks, L.W., Bagby, M.O., 1987. Kenaf lignin structure by correlation of CMR, FTIR and chemical analysis. 1987 Pulping Conference, pp. 177 183. Adler, E., 1977. Lignin chemistry-past, present and future. J. Wood Sci. Technol. 11, 169 218. Aorigele, X., Sano, Y., 1997. Fractionation and utilization of kenaf (Hibiscus cannabinus) components for conversion of biomass II. Alkaline oxygen pulping of bast fibers with epidermis. Mokuzai Gakkaishi 43, 171 177. Challinor, J.M., 2001. Review: the development and applications of thermally assisted hydrolysis and methylation reactions. J. Anal. Appl. Pyrol. 61, 3 34. Chang, H.-M., Allan, G.G., 1971. Oxidation. In: Sarkanen, K.V., Ludwig, C. (Eds.), Lignins: Occurrence, Formation, Structure and Reactions. John Wiley & Sons, Inc., New York, pp. 433 485. Clark, T.F., Wolff, I.A., 1969. A search for new fiber crops XI. Compositional characteristics of Illinois kenaf at several population densities and maturities. Tappi J. 11, 2111 2116. Collins, D., Pilotti, C., Wallis, A., 1990. Correlation of chemical composition and kraft pulping properties of some Papua New Guinea reforestation woods. Appita J. 43, 193 198. del Rio, J.C., Gutierrez, A., Martinez, A.T., 2004. Identifying acetylated lignin units in non-wood fibers using pyrolysis-gas chromatography/mass spectrometry. Rapid Commun. Mass Spectrom. 18, 1181 1185. Fergus, B.J., Goring, D.A.I., 1969. The topochemistry of delignification in kraft and neutral sulphite pulping of birch wood. Pulp Pap. Mag. Can. 19, 314 320. Filley, T.R., 2003. Assessment of fungal wood decay by lignin analysis using tetramethylammonium hydroxide (TMAH) and 13 C- labeled TMAH thermochemolysis. In: Goodell, B., Nicholas, D.D., Schultz, T.P. (Eds.), Wood Deterioration and Preservation. vol. 845, ACS Symposium Series, ACS, Washington, DC, pp. 119 139. Filley, T.R., Minard, R.D., Hatcher, P.G., 1999. Tetramethylammonium hydroxide (TMAH) thermochemolysis: proposed mechanisms based upon the application of 13 C-labeled TMAH to a synthetic model lignin dimer. Org. Geochem. 30, 607 621. Gellerstedt, G., 1992. Chemical degradation methods: permanganate oxidation. In: Lin, S.Y., Dence, C.W. (Eds.), Methods in Lignin Chemistry. Springer-Verlag, Berlin-Heidelberg, pp. 322 333. Gutierrez, A., Rodriguez, I.M., del Rio, J.C., 2004. Chemical characterization of lignin and lipid fractions in kenaf bast fibers used for manufacturing high-quality papers. J. Agric. Food Chem. 52, 4764 4773. Kaldor, A., 1989. Preparation of kenaf bark and core fibers for pulping by the Ankal method. Tappi J. 58, 137 140. Kaldor, A., 1992. A strategy for the development of a kenaf-based pulp and pulp industry. Tappi J. 75, 141 145. Karakus, S., Roy, D.N., Goel, K., 2001. Chemical and soda pulping properties of kenaf as a function of growth. J. Wood Chem. Technol. 21, 157 167.

232 K.-I. Kuroda et al. / Industrial Crops and Products 22 (2005) 223 232 Kuroda, K., Izumi, A., Mazumder, B.B., Ohtani, Y., Sameshima, K., 2002a. Characterization of kenaf (Hibiscus cannabinus) lignin by pyrolysis-gas chromatography-mass spectrometry in the presence of tetramethylammonium hydroxide. J. Anal. Appl. Pyrolysis 64, 453 463. Kuroda, K., Mazumder, B.B., Ohtani, Y., Sameshima, K., 25 26 February 1999. Characterization of kenaf (Hibiscus cannabinus L.) lignin by analytical pyrolysis. In: Proceedings of the 2nd Annual Conference of the American Kenaf Society, San Antonio, TX, USA, pp. 125 137. Kuroda, K., Nishimura, N., Nakagawa, A., Dimmel, D.R., 2002b. Pyrolysis of lignin in the presence of tetramethylammonium hydroxide (TMAH): a convenient method for S/G ratio determination. J. Agric. Food Chem. 50, 1022 1027. Kuroda, K., Suzuki, A., Kato, M., Imai, K., 1995. Analysis of rice (Oryza sativa L.) lignin by pyrolysis-gas chromatography. J. Anal. Appl. Pyrolysis 34, 1 12. Lai, Y.-Z., Sarkanen, K.V., 1971. Isolation and structural studies. In: Sarkanen, K.V., Ludwig, C.H. (Eds.), Lignins-Occurrence, Formation, Structure and Reactions. John Wiley & Sons, Inc., New York, pp. 165 200. Lapierre, C., 1993. Application of new methods for the investigation of lignin structure. In: Jung, H.G., Buxton, D.R., Hatfield, R.D., Ralph, J. (Eds.), Forage Cell Wall Structure and Digestibility. ASA, CSSA, SSSA, Madison, Winsconsin, USA, pp. 133 166. Mambelli, S., Grandi, S., 1995. Yield and quality of kenaf (Hibiscus cannabinus L.) stem as affected by harvest date and irrigation. Ind. Crops Prod. 4, 97 104. Mazumder, B.B., Nakagawa-izumi, A., Kuroda, K., Ohtani, Y., Sameshima, K., 2005. Evaluation of harvesting time effects on kenaf bast lignin by pyrolysis-gas chromatography. Ind. Crops Prod. 21, 17 24. Mazumder, B.B., Ohtani, Y., Sameshima, K., 1998. Normal pressure pulping of jute, kenaf and mesta bast fibers. Sen i Gakkaishi 54, 654 669. Meier, D., Faix, O., 1992. Pyrolysis-gas chromatography-mass spectrometry. In: Lin, S.Y., Dence, C.W. (Eds.), Methods in Lignin Chemistry. Springer-Verlag, Berlin-Heidelberg, pp. 177 199. Morrison III, W.H., Akin, D.E., Archibald, D., Raymer, P.L., 1999a. In: Sellers Jr., T., Reichert, N.A. (Eds.), Characterization of Kenaf Core and Bast using Pyrolysis Mass Spectrometry-Kenaf Properties, Processing and Products. Mississippi State University, Mississippi, pp. 169 186. Morrison III, W.H., Akin, D.E., Archibald, D.D., Dodd, R.B., Raymer, P.L., 1999b. Chemical and instrumental characterization of maturing kenaf core and bast. Ind. Crops Prod. 10, 21 34. Neto, C.P., Seca, A., Fradinho, D., Coimbra, M.A., Domingues, F., Evtuguin, D., Silvestre, A., Cavaleiro, J.A.S., 1996. Chemical composition and structure features of the macromolecular components of Hibiscus cannabinus grown in Portugal. Ind. Crops Prod. 5, 189 196. Nishimura, N., Izumi, A., Kuroda, K., 2002. Structural characterization of kenaf lignin: differences among kenaf varieties. Ind. Crops Prod. 15, 115 122. Ohtani, Y., Mazumder, B.B., Sameshima, K., 2001. Influence of the chemical composition of kenaf bast core on the alkaline pulping response. J. Wood Sci. 47, 30 35. Pappas, C., Tarantilis, P.A., Polissiou, M., 1998. Determination of kenaf (Hibiscus cannabinus L.) lignin in crude plant material using diffuse refectance infrared Fourier transform spectroscopy. Appl. Spectrosc. 52, 1399 1402. Ralph, J., 1996. An unusual lignin from kenaf. J. Nat. Prod. 59, 341 342. Ralph, J., Hatfield, R.D., 1991. Pyrolysis-GC-MS characterization of forage materials. J. Agric. Food Chem. 39, 1426 1437. Ralph, J., Lapierre, C., Lu, F.C., Marita, J.M., Pilate, G., van Doorsselaere, J., Boerjan, W., Jouanin, L., 2001. NMR evidence for benzodioxane structures resulting from incorporation of 5- hydroxyconiferyl alcohol into lignins of O-methyltransferasedeficient poplars. J. Agric. Food Chem. 49, 86 91. Ralph, J., Lu, F.C., 1998. The DFRC method for lignin analysis. Part 6. A simple modification for identifying natural acetates on lignins. J. Agric. Food Chem. 46, 4616 4619. Rodrigues, J., Meier, D., Faix, O., Pereira, H., 1999. Determination of tree to tree variation in syringyl/guaiacyl ratio of Eucalyptus globulus wood lignin by analytical pyrolysis. J. Anal. Appl. Pyrol. 48, 121 128. Rolando, C., Monties, B., Lapierre, C., 1992. Chapter 6.4: Thioacidolysis. In: Lin, S.Y., Dence, C.W. (Eds.), Methods in Lignin Chemistry. Springer-Verlag, Berlin-Heidelberg, pp. 334 349. Romanoschi, O., Romanoschi, S., Coller, J.R., Collier, B.J., 1997. Kraft alkali processing. Cellul. Chem. Technol. 31, 347 359. Seca, A.M.L., Cavaleiro, J.A.S., Domingues, D.E., Silvestre, A.J.D., Evtuguin, D.V., Neto, C.P., 1997. Structural characteristics of the bark and core kenaf lignin (variety Salvador). In: Proceedings of the 9th International Symposium on Wood and Pulping Chemistry, Montreal, Canada, pp. 1321 1324. Seca, A.M.L., Cavaleiro, J.A.S., Domingues, F.M.J., Silverstre, A.J.D., Evtuguin, D., Neto, C.P., 1998. Structural characterization of the bark and core lignins from kenaf (Hibiscus cannabinus). J. Agric. Food Chem. 46, 3100 3108. Suzuki, S., Lam, T.B.T., Iiyama, K., 1997. 5-Hydroxyguaiacyl nuclei as aromatic constituents of native lignin. Phytochemistry 46, 695 700. TAPPI, 1988. Pulp testing. In: Tappi Testing Procedures. Tappi Press, Atlanta, GA, USA. van Bergen, P.F., Hatcher, P.G., Boon, J.J., Collinson, M.E., de Leeuw, J.W., 1997. Macromolecular composition of the propagule wall of Nelumbo nucifera. Phytochemistry 45, 601 610. van Bergen, P.F., Poole, I., Ogilvie, T.M.A., Caple, C., Evershed, R.P., 2000. Evidence for demethylation of syringyl moieties in archaeological wood using pyrolysis-gas chromatography/mass spectrometry. Rapid Commun. Mass Spectrom. 14, 71 79. Zhou, C., Zhengguo, L., Sameshima, K., Ohtani, Y., Zhen, M., 1995a. On agronomic foundation of kenaf for papermaking. Acta Agriculturae Zheijiangensis 7, 38 41. Zhou, C., Zhengguo, L., Sameshima, K., Ohtani, Y., Zhen, M., 1995b. On the fiber length and its distribution in kenaf. Acta Agriculturae Zheijiangensis 7, 293 296.