Lignin monomer composition affects Arabidopsis cell-wall degradability after liquid hot water pretreatment
- Xu Li†1,
- Eduardo Ximenes†2,
- Youngmi Kim2,
- Mary Slininger2,
- Richard Meilan3,
- Michael Ladisch2, 4Email author and
- Clint Chapple1Email author
© Li et al; licensee BioMed Central Ltd. 2010
Received: 27 September 2010
Accepted: 2 December 2010
Published: 2 December 2010
Lignin is embedded in the plant cell wall matrix, and impedes the enzymatic saccharification of lignocellulosic feedstocks. To investigate whether enzymatic digestibility of cell wall materials can be improved by altering the relative abundance of the two major lignin monomers, guaiacyl (G) and syringyl (S) subunits, we compared the degradability of cell wall material from wild-type Arabidopsis thaliana with a mutant line and a genetically modified line, the lignins of which are enriched in G and S subunits, respectively.
Arabidopsis tissue containing G- and S-rich lignins had the same saccharification performance as the wild type when subjected to enzyme hydrolysis without pretreatment. After a 24-hour incubation period, less than 30% of the total glucan was hydrolyzed. By contrast, when liquid hot water (LHW) pretreatment was included before enzyme hydrolysis, the S-lignin-rich tissue gave a much higher glucose yield than either the wild-type or G-lignin-rich tissue. Applying a hot-water washing step after the pretreatment did not lead to a further increase in final glucose yield, but the initial hydrolytic rate was doubled.
Our analyses using the model plant A. thaliana revealed that lignin composition affects the enzymatic digestibility of LHW pretreated plant material. Pretreatment is more effective in enhancing the saccharification of A. thaliana cell walls that contain S-rich lignin. Increasing lignin S monomer content through genetic engineering may be a promising approach to increase the efficiency and reduce the cost of biomass to biofuel conversion.
Utilization of lignocellulosic biomass for biofuel production requires the hydrolysis of cellulose and other cell-wall polysaccharides to their component monosaccharides. This process is affected by many structural and compositional characteristics of the biomass, including the presence of lignin, a phenolic polymer composed of three major types of building blocks: p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units. The association between lignin and the recalcitrance of biomass materials has long been recognized in forage feeding and tree pulping practices, and led to earlier lignin-engineering efforts aimed at improving feedstock performance in these processes . For example, it has been shown that lignin content (the total amount of lignin in the tissue) greatly influences forage digestibility , and that lignin composition (the relative ratio of its component subunits) strongly affects the efficiency of the chemical pulping process. Specifically, transgenic poplar trees with higher S/G ratios show a greatly enhanced pulping efficiency . A recent analysis of transgenic alfalfa revealed that high lignin content is correlated with the recalcitrance of cell-wall materials to enzymatic saccharification during biofuel production . There was little variation in the S:G ratios of these alfalfa lines and as a result, the effects of lignin composition on the efficiency of biomass to biofuel conversion remain to be determined.
To investigate the effect of S-lignin composition on biomass degradability, we chose to analyze two A. thaliana lines in which the activity or expression of ferulate 5-hydroxylase (F5H), a key enzyme required for the synthesis of S-lignin monomer, is eliminated or enhanced, respectively. Both lines have been generated previously and characterized in detail [5–8]. The fah1-2 mutant is defective in F5H and does not deposit S lignin, whereas overexpression of F5H under the control of the cinnamate 4-hydroxylase (C4H) promoter in the C4H:F5H transgenic line results in lignin with an S-unit content in excess of 90% . These lines represent two extremes in lignin composition and are each distinct from the wild type, which deposits a G/S copolymer with an S-subunit content of approximately 20 mol%. Despite their lignin difference, these two lines show similar growth to that of wild-type Arabidopsis.
In the biomass to biofuel conversion processes, pretreatment is generally used to increase the accessibility of cell-wall polysaccharides to enzymes. Pressure-cooking in liquid hot water (LHW) has been shown to be a cost-effective pretreatment to enhance the enzymatic digestibility of cellulose in a variety of feedstocks [9–12]. In this study, we assessed the cell-wall degradability of mutant and transgenic A. thaliana lines by enzyme hydrolysis without pretreatment, after LHW pretreatment, or after LHW pretreatment followed by hot water washing. Our results indicate that increasing the proportion of S subunits in Arabidopsis lignin decreases the recalcitrance of cell walls to enzymatic hydrolysis.
Derivatization followed by reductive cleavage (DRFC) lignin and compositional analysis of Arabidopsis thaliana samples1
DFRC lignin (mol%)
Percentage by dry mass
Enzyme hydrolysis of untreated samples
Enzyme hydrolysis with LHW pretreatment
We further tested the cell-wall degradability of the lines by applying LHW pretreatment before enzyme hydrolysis. Regardless of genotype, the hydrolytic rate and glucose yield of pretreated samples was greater than for the corresponding untreated samples (Figure 1B). Incubation of LHW-pretreated wild-type samples with enzymes for 1 hour released the same amount of glucose (20% of total glucan) as 6 hours of hydrolysis on untreated samples. More importantly, significant differences in cellulose hydrolysis were observed for LHW-pretreated fah1-2 and C4H:F5H samples (Figure 1B).No differences in glucose yield between the three genotypes were observed after 1 hour of incubation, but after 3 hours of incubation, C4H:F5H yielded more glucose and fah1-2 less glucose, compared with wild type. After 24 hours, the glucose yield of the C4H:F5H sample was nearly 90%.
Scanning electronic microscopy
Our enzyme hydrolysis results suggest that LHW pretreatment is differentially effective against A. thaliana samples with altered lignin composition.
Enzyme hydrolysis after LHW pretreatment and hot-water washing
It has been shown that hot-water washing of LHW pretreated poplar can further improve its enzymatic digestibility . Therefore, we also tested the cell-wall degradability of the A. thaliana samples after this additional step (Figure 1C). Unlike poplar, hot-water washing did not have a significant effect on the total glucose yield as calculated based on the initial glucan content of the Arabidopsis samples tested; however, the initial rate of cellulose hydrolysis of hot-water washed C4H:F5H samples was almost double that of unwashed samples. By contrast, hot-water washing has little effect on the initial hydrolytic rate of the wild-type and fah1-2 samples.
Phenolic inhibitors analysis
Concentration of total phenolics in liquid collected after liquid hot water pre-treatment.1,2
Concentration of total phenolics, mg TAE/ml3
0.30 ± 0.19
0.35 ± 0.01
0.30 ± 0.08
Lignin is a major contributor to the recalcitrance of biomass, and has been a target for feedstock improvement through genetic engineering. It has been demonstrated in several plant species that a reduction in lignin content using transgenic approaches enhances cell-wall degradability; however, significant improvement in conversion efficiency has often been accompanied by abnormal plant growth and development [1, 4]. By contrast, plants seem to be amenable to wide ranges in lignin composition changes, including variation in the content of conventional monomers and the incorporation of atypical precursors [6, 16–18]. Our findings of improved cell-wall degradability in Arabidopsis stems with high S-lignin content demonstrate the potential of lignin composition modification for the improvement of cellulosic feedstock performance.
LHW treatment had a dramatic effect on enzyme hydrolysis of biomass from the high S-lignin line. As much as 90% of the maximum theoretical glucose yield was achieved for C4H:F5H tissue, whereas less than 60% was obtained from wild-type material exposed to the same treatment. SEM analysis detected no obvious anatomical differences between the low and high S-lignin samples after LHW pretreatment. By contrast, after LHW pretreatment and enzyme hydrolysis, stem cross-sections from the high S-lignin stems had a distinct deformity, presumably due to enhanced hydrolysis of the cell wall.
How alteration of lignin composition relates to the observed increase in the effectiveness of LHW pretreatment is unclear. Recently, it was proposed that during high-temperature pretreatment, lignin is melted and relocalized to outer surface of the cell wall, increasing the accessibility of the cellulose within [19, 20]. The S:G ratio is known to have profound effects on lignin structure. Whereas G-rich lignin has a branched structure, S-rich lignin is more linear and has a lower degree of polymerization . It is tempting to speculate that S-rich lignin may have a lower melting point and is more easily relocated than G-rich lignin and, thereby, leads to improved enzymatic digestibility.
The observation that hot-water washing after LHW pretreatment significantly increases the initial saccharification rate of the high-S sample suggests removal of some inhibitory compounds. It has been shown LHW pretreatment of wet cake (solids left after corn is fermented to ethanol) releases some phenolics and water soluble xylo-oligosaccharides that can inhibit cellulases and β-glucosidases . However, in this study, the concentration of phenolics in the liquid after LHW pretreatment was below the level reported to cause significant inhibition. Moreover, we did not see any difference in the concentration of phenolics between S-deficient, S-rich and wild-type A. thaliana tissue samples. Therefore, it is unlikely that the observed increase in saccharification rate was due to the removal of inhibitors. The relative influence of phenolic molecules on enzyme activity becomes more pronounced as the ratio of phenols to protein increases at higher solid and lower protein loadings than used in this study. Although protein loadings are currently in the range of 2 to 10 mg/g lignocellulose solids, which represent up to a five-fold decrease in enzyme from only 5 years ago, an even greater reduction is needed for economically viable processes [13, 22].
It will be important to extend this study to other biomass feedstocks in the future. One of the factors that needs to be considered is the native lignin composition of different plant species, and possibly also the interaction of lignin with the polysaccharide components of the cell wall. It is likely that the dramatic increase in cell-wall degradability observed in Arabidopsis might be less apparent in plant species with high native S-lignin levels, such as hybrid poplar. However, significant increases in lignin extractability during the pulping process has been observed in S-lignin-enriched transgenic poplar , suggesting that the magnitude of S lignin increase may still contribute significantly to higher cell-wall degradability in this important biomass feedstock.
The available genetically modified A. thaliana plants with different lignin composition and structure provided an opportunity to evaluate the possible effect of lignin modification on cell-wall recalcitrance. Our study revealed that high levels of S-lignin have a positive effect on the effectiveness of LHW pretreatment and enzymatic hydrolysis, at least in Arabidopsis. This effect might result from the physicochemical changes of lignin brought about by more linear structure of S subunits. In the future, it will be important to determine how widespread is this phenomenon and to elucidate its underlying mechanism.
Generation of the A. thaliana fah1-2 and C4H:F5H lines has been described previously [6, 7]. The genetically modified and wild-type A. thaliana (Columbia) plants were grown side by side at 22°C under a 16-hour photoperiod. Mature stems were harvested by removing siliques and leaves.
Spezyme CP cellulase preparation from Trichoderma reesei containing exo-, endo- and β-glucosidase activities; batch number 3016295230) was provided by Genencor, Danisco Division (Palo Alto, CA, USA). Novozyme 188 (β-glucosidase from Aspergillus niger; catalogue number c6150) was purchased from Sigma Chemical Co. (St. Louis, MO, USA).
Lignin composition was determined using the derivatization followed by reductive cleavage (DFRC) method .
The composition of the plant samples was analyzed using standard National Renewable Energy Laboratory procedures . To calculate polysaccharide composition, monomer sugars were analyzed by high-performance liquid chromatography (HPLC) after acid hydrolysis of the samples. HPLC analysis of liquid samples was performed on a system consisting of a solvent-delivery system (9010 Gradient HPLC Pump, Varian/Agilent, Santa Clara CA, USA), an autosampler (717 Plus; Waters Corp., Milford, MA, USA), a carbohydrate analysis column (Aminex HPX-87H; Bio-Rad, Hercules, CA, USA); a refractive index detector (2414; Waters Corp.) a dual-wavelength absorbance detector (2487; Waters Corp.); and an integrator (HP3396G; Hewlett Packard, Santa Clara CA, USA). The mobile phase was 5 mmol/l H2SO4 filtered through a 0.2 μm nylon filter (Millipore Corp., Billerica, MA, USA) and degassed. The mobile phase flow rate was 0.6 ml/min and the column temperature was maintained at 60°C by a column heater (CH-30; Eppendorf) with a temperature controller (TC-50; Eppendorf, Hauppauge, New York, USA).
Cell-wall degradability analysis
The stem materials were ground for passage through a 20 mesh (841 μm) screen for cell-wall degradability analyses. Pretreatment was carried out by pressure cooking 50 mg samples in a metal tube containing 1.5 ml water at 200°C (30 seconds of heat-up time followed by a 10 minute hold). Each tube was placed in a fluidized sand bath (Tecam® SBL-1; Cole-Parmer, Vernon Hills, IL). The pressure within the tubes was held at the saturation vapor pressure of water to keep the water in a liquid state [9, 11–13]. The samples were cooled before the addition of 1.5 ml of 100 mmol/l citrate buffer pH 4.8, bringing the final volume to 3 ml (~2% solids (w/v)). For hot-water washing, samples were washed twice with 3 ml of 70°C water. After the second wash, no glucose was detected. The enzyme hydrolysis for all the conditions tested was based on initial solids loading and glucan concentration. Commercial cellulase (Spezyme CP) at 0.2 filter paper units (FPU)/ml or 50 FPU/g glucan (90 mg protein/g glucan) and β-glucosidase (Novozyme 188) at 0.35 cellobiase units (CBU)/ml or 105 CBU/g glucan (34 mg protein/g glucan) were added, and hydrolysis was carried out for different lengths of time at 50°C and pH 4.8 in an incubator shaker (New Brunswick Scientific, Edison, NJ, USA). The ratio of enzyme to solids was equivalent to 10 FPU/g total solids, 21 CBU/g total solids and 25 mg protein/g total solids. Enzyme hydrolysis of 50 mg untreated samples (also at ~2% solids (w/v), 50°C and pH 4.8) was carried out under similar experimental conditions.
Stem cross-sections were adhered to a glass slide with epoxy adhesive (J-B Weld; J-B Weld Co., Sulphur Springs, TX, USA) and subjected to different treatments. Pretreatment was performed as described in the previous section. When applied, enzyme hydrolysis was carried out with a two-fold increase in enzyme loading to compensate for the particle size differences between the samples used for SEM and cell-wall degradability analysis. Subsequently, the stem cross-sections were fixed in two steps: first with a mixture of 2% paraformaldehyde and 2.5% glutaraldehyde in 0.1 mol/l cacodylate buffer, pH 7.4 for 1 hour and then with 1% OsO4 in 0.1 mol/l cacodylate buffer, pH 7.4 for 30 minutes. After critical point drying, the samples were sputter-coated with gold, and viewed under SEM (Nova nanoSEM; Fei Co., Hillsboro, OR, USA).
Protein and phenolic measurements
The protein content of the commercial enzyme preparations was determined using a commercial kit (Pierce BCA Protein Assay Kit; product number 23225; Thermo Scientific, Rockford, IL, USA). Phenolic compounds were assayed using Prussian blue . The diluted sample liquid (3 ml aliquots) was transferred to a 10 mm cuvette, then 200 μL of 0.008 mol/l K2Fe(CN)6 were added, followed by the immediate addition of 200 μL of 0.1 mol/l FeCl3 in 0.1 mol/l HCl. Absorbance was read at 700 nm after 5 min at room temperature, against a tannic acid standard.
We thank Junko Maeda, Xingya (Linda) Liu, Rick Hendrickson and Thomas Kreke for their technical assistance. We also thank Chia-Ping Huang at Purdue Life Science Microscopy Facility for assistance with SEM imaging. Finally, we are grateful for the enzymes provided by Genencor. This work was supported by USDA IFAFS contract #00-52104-9663 and DOE grant #DE-FG02-06ER64301.
- Li X, Weng JK, Chapple C: Improvement of biomass through lignin modification. Plant J 2008, 54: 569-581. 10.1111/j.1365-313X.2008.03457.xView Article
- Reddy MS, Chen F, Shadle G, Jackson L, Aljoe H, Dixon RA: Targeted down-regulation of cytochrome P450 enzymes for forage quality improvement in alfalfa ( Medicago sativa L. ). Proc Natl Acad Sci USA 2005, 102: 16573-16578. 10.1073/pnas.0505749102View Article
- Huntley SK, Ellis D, Gilbert M, Chapple C, Mansfield SD: Significant increases in pulping efficiency in C4H-F5H-transformed poplars: improved chemical savings and reduced environmental toxins. J Agric Food Chem 2003, 51: 6178-6183. 10.1021/jf034320oView Article
- Chen F, Dixon RA: Lignin modification improves fermentable sugar yields for biofuel production. Nat Biotechnol 2007, 25: 759-761. 10.1038/nbt1316View Article
- Meyer K, Cusumano JC, Somerville C, Chapple CC: Ferulate-5-hydroxylase from Arabidopsis thaliana defines a new family of cytochrome P450-dependent monooxygenases. Proc Natl Acad Sci USA 1996, 93: 6869-6874. 10.1073/pnas.93.14.6869View Article
- Meyer K, Shirley AM, Cusumano JC, Bell-Lelong DA, Chapple C: Lignin monomer composition is determined by the expression of a cytochrome P450-dependent monooxygenase in Arabidopsis. Proc Natl Acad Sci USA 1998, 95: 6619-6623. 10.1073/pnas.95.12.6619View Article
- Chapple CC, Vogt T, Ellis BE, Somerville CR: An Arabidopsis mutant defective in the general phenylpropanoid pathway. Plant Cell 1992, 4: 1413-1424. 10.1105/tpc.4.11.1413View Article
- Marita JM, Ralph J, Hatfield RD, Chapple C: NMR characterization of lignins in Arabidopsis altered in the activity of ferulate 5-hydroxylase. Proc Natl Acad Sci USA 1999, 96: 12328-12332. 10.1073/pnas.96.22.12328View Article
- Weil J, Sarikaya A, Rau SL, Goetz J, Ladisch C, Brewer M, Hendrickson R, Ladisch M: Pretreatment of corn fiber by pressure cooking in water. Appl Biochem Biotechnol 1998, 73: 1-17. 10.1007/BF02788829View Article
- Yu G, Yano S, Inoue H, Inoue S, Endo T, Sawayama S: Pretreatment of rice straw by a hot-compressed water process for enzymatic hydrolysis. Appl Biochem Biotechnol 2010, 160: 539-551. 10.1007/s12010-008-8420-zView Article
- Kim Y, Mosier NS, Ladisch MR: Enzymatic digestion of liquid hot water pretreated hybrid poplar. Biotechnol Prog 2009, 25: 340-348. 10.1002/btpr.137View Article
- Mosier N, Hendrickson R, Ho N, Sedlak M, Ladisch MR: Optimization of pH controlled liquid hot water pretreatment of corn stover. Bioresour Technol 2005, 96: 1986-1993. 10.1016/j.biortech.2005.01.013View Article
- Mosier N, Wyman C, Dale B, Elander R, Lee YY, Holtzapple M, Ladisch M: Features of promising technologies for pretreatment of lignocellulosic biomass. Bioresour Technol 2005, 96: 673-686. 10.1016/j.biortech.2004.06.025View Article
- Ximenes E, Kim Y, Mosier N, Dien B, Ladisch M: Inhibition of cellulases by phenols. Enzyme Microb Tech 2010, 46: 170-176. 10.1016/j.enzmictec.2009.11.001View Article
- Ximenes E, Kim Y, Mosier N, Dien B, Ladisch M: Deactivation of cellulases by phenols. Enzyme Microb Tech 2010. Accepted for publication on 3 Sept 2010
- Ralph J, Lapierre C, Marita JM, Kim H, Lu F, Hatfield RD, Ralph S, Chapple C, Franke R, Hemm MR, et al.: Elucidation of new structures in lignins of CAD- and COMT-deficient plants by NMR. Phytochemistry 2001, 57: 993-1003. 10.1016/S0031-9422(01)00109-1View Article
- Ralph J, Akiyama T, Kim H, Lu F, Schatz PF, Marita JM, Ralph SA, Reddy MS, Chen F, Dixon RA: Effects of coumarate 3-hydroxylase down-regulation on lignin structure. J Biol Chem 2006, 281: 8843-8853. 10.1074/jbc.M511598200View Article
- Franke R, Hemm MR, Denault JW, Ruegger MO, Humphreys JM, Chapple C: Changes in secondary metabolism and deposition of an unusual lignin in the ref8 mutant of Arabidopsis. Plant J 2002, 30: 47-59. 10.1046/j.1365-313X.2002.01267.xView Article
- Kristensen JB, Thygesen LG, Felby C, Jorgensen H, Elder T: Cell-wall structural changes in wheat straw pretreated for bioethanol production. Biotechnol Biofuels 2008, 1: 5. 10.1186/1754-6834-1-5View Article
- Selig MJ, Viamajala S, Decker SR, Tucker MP, Himmel ME, Vinzant TB: Deposition of lignin droplets produced during dilute acid pretreatment of maize stems retards enzymatic hydrolysis of cellulose. Biotechnol Prog 2007, 23: 1333-1339. 10.1021/bp0702018View Article
- Stewart JJ, Akiyama T, Chapple C, Ralph J, Mansfield SD: The effects on lignin structure of overexpression of ferulate 5-hydroxylase in hybrid poplar. Plant Physiol 2009, 150: 621-635. 10.1104/pp.109.137059View Article
- Sathitsuksanoh N, Zhu Z, Ho TJ, Bai MD, Zhang Y-HP: Bamboo saccharification through cellulose solvent-based biomass pretreatment followed by enzymatic hydrolysis at ultra-low cellulase loadings. Bioresour Technol 2010, 101: 4926-4929. 10.1016/j.biortech.2009.09.081View Article
- Lu FC, Ralph J: Derivatization followed by reductive cleavage (DFRC method), a new method for lignin analysis: Protocol for analysis of DFRC monomers. J Agric Food Chem 1997, 45: 2590-2592. 10.1021/jf970258hView Article
- Ruiz R, Ehrman T: NREL analytical procedure: LAP 002. Determination of carbohydrates in biomass by high performance liquid chromatography. National Renewable Energy Laboratory, Golden, CO; 1996.
- Budini R, Tonelli D, Girotti S: Analysis of total phenols using the Prussian Blue method. J Agric Food Chem 1980, 28: 1236-1238. 10.1021/jf60232a056View Article
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