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32.5: Biofuels B - Cellulosic Ethanol

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    Search Fundamentals of Biochemistry


    In the last section, we explored how ethanol can be made from corn starch, an α(1,4) polymer of glucose with α(1,6) branches. Its production comes at a cost, however. Recent life cycle studies have shown that compared to fossil fuels, corn ethanol release as much but probably more CO2 than from fossil fuels. In addition, is it ethically justifiable to remove so much land from food production to produce bioethanol that, at present, is worse than fossil fuels from a climatic perspective?

    To address these issues, much work has been done to produce ethanol from cellulose, a β(1,4) polymer of glucose and the most abundant biomolecule in the world. Cellulose from trees, switch grasses, and "waste biomass" are prime sources of cellulose for the production of bioethanol. Waste biomass includes stover (field crop stalks and leaves), straw, wood chips and sawdust. From one ton of corn stover, about 113 gallons of ethanol can be made, close to the 124 gallons produced from corn. 

    Nature breaks down cellulose routinely using cellulases, enzymes found in bacteria, fungi, protozoans, plants and some animals. Ruminants and even termites obtain cellulases from microbes living within their guts. The fungal-mediated decay of dead trees requires microbial cellulases but think how slow that process is. This stems partly from the very strong β(1,4) glycosidic link connecting glucose monomers in the polymer, which in the absence of a catalyst and at neutral pH, has an estimated half-life of 5 million years. Fossilized plants have been found to have intact cellulose and chitin, a β(1,4) polymer of N-acetylglucosamine. The β(1,4) glycosidic link in cellulose is orders of magnitude more stable than the phosphodiester bond of nucleic acids and the amide link of proteins. They are, however, readily cleaved by glycosidases such as cellulases, which can increase the kcat/KM over the uncatalyzed rate by up to 1017 fold, even in the absence of active site metal ions to facilitate hydrolysis (reference).  

    Another reason for the slow decay of dead trees is the complex structure of the cell well, and in particular, the presence of a polymer called lignin, which stabilizes the cell wall and adds considerable barriers to the access of cellulose by added glycosidases. .  

    A final reason for cellulose's extreme stability is the "quaternary" structure of the β(1,4)-linked cellulose strands, which consists of densely packed and intertwined strands of cellulose, which limits solvent (in this case water) accessibility necessary for hydrolysis. In addition,  some exposed surface planes of the packed cellulose strands are hydrophobic. This might seem startling, given the polar nature of the glucose subunits of the polymer. Let's review it here now since our goal is present climate change from a biochemical perspective! Some of this material has been presented in previous chapters,  but we will reuse it here so this chapter section can stand alone.

    A review:  The Plant Cell Wall

    (See Chapter 7.3 for more details.) There are about 35 different types of plant cells, and each may have a different cell wall depending on the local needs of a given cell. Cells synthesize thin cell wall that extends and stay thin as the cell grows. Figure \(\PageIndex{1}\) shows the primary cell wall of plants. The primary cell wall contains cellulose microfibrils (no surprise) and two other polymers, pectin and hemicellulose. The middle lamella consisting of pectins, is somewhat analogous to the extracellular matrix.

    Figure \(\PageIndex{1}\): Primary cell wall of plants.

    After cell growth, the cell often synthesizes a secondary cell wall thicker than the first for extra rigidity. Since the enzymatic machinery for its synthesis is in the cytoplasm and the cell membrane, it is deposited between the cell membrane and the primary cell wall, as shown in the animated image in Figure \(\PageIndex{2}\).

    Figure \(\PageIndex{2}\): Primary and secondary cell wall of plants

    Figure \(\PageIndex{3}\) shows a structural representation of both the primary and secondary cell wall.

    NAC-MYB-based transcriptional regulation of secondary cell wall biosyn in land plantsIMAGE-01.svg
    Figure \(\PageIndex{3}\): structural representation of both the primary and secondary cell wall of plants. Nakano Yoshimi et al. Frontiers in Plant Science (6), 288 (2015) Creative Commons AttributionLicense (CC BY).

    The middle lamella, which contains pectins, lignins and some proteins, helps "glue together" the primary cell walls of surrounding plants.

    Primary Cell Wall:

    The main component of the primary plant wall is the homopolymer cellulose (40% -60% mass) in which the glucose monomers are linked β(1→4)-linked into strands that collect into microfibrils through hydrogen bond interactions. Two other groups of polymers, hemicellulose, and pectin, make up the plant cell wall.

    Hemicellulose can make up to 20-40% by the mass These polymers have β(1,4) backbones of glucose, mannose, or xylose (called xyloglucans, xylans, mannans, galactomannans, glucomannans, and galactoglucomanannans along with some β(1,3 and 1,4)-glucans. The most abundant hemicellulose in higher plants higher plants are the xyloglucans and have a cellulose backbone linked at O6 to α-D-xylose. Pectin consists of linked galacturonic acids forming homogalacturonans, rhamnogalacturonans, and rhamnogalacturonans II (RGII) [12] [13]. Homogalacturonans (α1→4) linked D-GalA make up more than 50% of the pectin. Figure \(\PageIndex{4}\) shows some of the structures.  The are generally branched, shorter than cellulose chains, and can often crystallize.  


    Figure \(\PageIndex{4}\): Variant of the cell wall components of a plant. Costa and Plazanet. Advances in Biological Chemistry 06(03):70-105. DOI: 10.4236/abc.2016.63008License CC BY 4.0

    Secondary Cell Wall

    The structure of the secondary cell wall depends on the function and environment of the cell. It contains cellulose fibers, hemicellulose, and a new polymer, lignin. It is abundant in xylem vessels and fiber cells of woody plants. It gives the plant extra stability and new functions, including the transport of fluids within the plant through channels. The proportion of cellulose in the secondary cell wall is higher than in the primary cell wall and is less hydrated than in the primary cell wall. Given the relative volume of the secondary and primary cell walls inferred from Fig 2, most of the tree-derived cellulose for bioethanol production comes from the secondary cell wall. Switch grasses, a perennial plant, are also valuable sources of cellulose (32–45% wt) and hemicellulose (21–31% wt) but also have significant amounts of lignin (12–28% wt). In summary, the secondary cell wall, formed after the cell stop growing, accounts for most of the carbohydrate biomass of plants. 

    Glycosidases, mostly α- and β-amylases, are needed to convert corn-derived starch into glucose for fermentation and ethanol production. Likewise, cellulases are needed to degrade cellulose into glucose for cellulosic-ethanol production. However, it is a much more complex process since most of the cellulose is in the secondary cell wall. The lignin barrier in the walls protects cellulose from accessibility to cellulases, even after chemical and thermal pre-processing. In addition, xylans, which can make up 30% of the mass of the secondary cell wall, also inhibit cellulose degradation. 

    A thermochemical process can convert cellulose to the synthetic gases CO and H2, which can be used as reactants to form ethanol. We'll discuss the biochemical process using pretreatment and enzymatic hydrolysis to make cellulosic ethanol. Lignin can be recovered and used to provide energy for the industrial-scale synthesis of cellulosic ethanol.  

    Let's explore the barriers posed by lignin and how they can be surmounted to facilitate access to cellulose and the liberation of glucose for cellulosic ethanol production.

    Lignin Structure and reactivity

    Lignins, which can make up to 25% of the biomass weight of secondary walls, are made from phenylalanine derivatives but more directly from cinnamic acid. This derives from is made from phenylalanine which is hydroxylated and converted through other steps to hydroxycinnamic alcohols called monolignols, as shown in Figure \(\PageIndex{5}\). Three typical monomers, p-coumaryl, coniferyl, and sinapyl alcohols, can polymerize into lignins, with their units in the polymer (P) named hydroxyphenyl, guaiacyl and syringly, respectively.


    Figure \(\PageIndex{5}\): Monolignols and their polymers

    Lignols are activated phenolic compounds, which form phenoxide free radicals (catalyzed by enzymes called peroxidases), which can attack a second lignol to form covalent dimers. Reaction mechanisms for the dimerization of the MS sinapyl alcohol free radical are shown as an example in Figure \(\PageIndex{6}\).


    Figure \(\PageIndex{6}\): Dimers of lignols

    Now imagine this polymerization continuing through the formation of more phenolic free radicals and coupling at a myriad of sites to form a large covalent lignin polymer. Figure \(\PageIndex{7}\) shows one example of a larger lignin.


    Figure \(\PageIndex{7}\): A larger lignin. . By Smokefoot - Own work, CC BY-SA 3.0,

    Lignin strengthens the cell wall and further stabilizes the already unreactive cellulose fibers. Let's look at a specific example - using corn stover (CS) as a cellulose source - of how pretreatment of the biomass source with a chemical treatment followed by the addition of a bacterial strain Pandoraea sp. B-6 (B-6) isolated from long, narrow strips of bamboo (slips). Bamboo is a type of woody grass that grows rapidly. These bacteria produce two extracellular lignin-degrading enzymes, manganese peroxidase (MnP) and laccase (Lac). Laccase (Lac) is a multi-copper oxidase that uses O2 as an oxidizing agent in the degradation of the syringyl, guaiacyl and p-hydroxyphenyl monomers in lignins. MnP has similar properties. These and other enzymes can lead to the depolymerization of lignin and degradation of lignin-derived aromatic compounds

    The adddition of the B-6 bacteria (a source of MnP and Lac) to milled corn stover (CS) did not increase the rate of lignin degradation unless the corn stover was preincubated with a tetrahydrofuran–water (THF–H2O) with 0.5 wt% sulfuric acid and heated to 150 oC. This led to the erosion of the corn stover, allowing access to the bacterial enzyme. The untreated and pre-treated CS surface, along with a diagram showing access of Lac and MnP to the lignin, is shown in Figure \(\PageIndex{8}\).

    bacteria for improving the lignocellulose biorefinery processFig6.svg

    Figure \(\PageIndex{8}\): Untreated and pre-treated CS surface, and Lac and MnP interaction with lignins.  Zhuo, S., Yan, X., Liu, D. et al. Use of bacteria for improving the lignocellulose biorefinery process: importance of pre-erosion. Biotechnol Biofuels 11, 146 (2018).  Creative Commons Attribution 4.0 International License (

    In addition to restricting access of cellulase to cellulose, cellulase can also nonspecifically adsorb to lignin and its pretreated forms since the lignin derivatives present a more hydrophobic surface that promotes cellulase interactions. Some plant laccases are involved in lignin biosynthesis, whereas in bacteria and fungi, they may be involved in lignin degradation

    Of course, fungi, which are prime degraders of dead biomass in forests, are also sources of enzymes for lignin degradation. For example, species of white rot fungi produce manganese peroxidase (MnP), lignin peroxidase (LiP), versatile peroxidase (VP), and laccase (Lac). They work through forming reactive lignin-derived aromatic free radicals (similar to those produced in lignin synthesis), leading to breaking ether bonds, aromatic ring cleavage and removal of methoxy groups from the substrate in a process called delignification. Pretreatment of the biomass increases yields higher amounts of available cellulose. Fungi, however, grow slowly, and the rate of delignification is still low. In addition, they also have hydrolytic enzymes that decrease the yield of cellulose. That is why bacterial sources like B6 are sought for delignification.  

    As this is a biochemistry textbook, let's explore the structure and function of fungal laccase. The enzyme can bind a large variety of hydroxylated- and methyoxy-aromatic compounds as substrates, so its active site must be adaptable and likely dynamic. Structural analyses, in-silico docking experiments, and molecular dynamics simulations have been performed with the laccase (TvL) from the fungus Trametes versicolor.  

    The enzyme has four copper ions in a T1 Cu site and a tri-nuclear Cu cluster (T2 Cu, T3α Cu and T3β Cu) at a T2/T3 site. As the mechanism involves free radical intermediates with O2 as an oxidant and substrate, 4 electrons are passed in single electron steps to the T1 Cu, then to the other three coppers, and finally to O2 to form two water molecules as products. The amino acid side chain ligands for the four copper ions are shown for white rot fungi laccase from Trametes Versicolor in Figure \(\PageIndex{9}\).


    Figure \(\PageIndex{9}\): T1 Cu (top left) and the trinuclear Cu cluster (T2, T3α and T3β) and their ligands for Trametes Versicolor laccase (TvL, pdb:  1GYC)

    Fungal laccases are extracellular proteins with about 550 amino acids arranged in three cupredoxin-like, beta-barrel domains. The T1 Cu is close to the surface and is found in domain 3, while the other copper ions are buried at the interface to domains 1 and 3. Figure \(\PageIndex{10}\) shows an interactive iCn3D model of Laccase from the Fungus Trametes Versicolor (1GYC)

    Laccase from the Fungus Trametes Versicolor (1GYC).png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{10}\): Laccase from the Fungus Trametes Versicolor (1GYC). (Copyright; author via source).
    Click the image for a popup or use this external link:

    Domain 1 is green, domain 2 magenta and domain 3, which contains the single T1 Cu, orange. The protein is glycosylated, as shown in the blue glycan cube cartoons representing N-acetylglucosmine. Key substrate binding and catalytic side chains are shown in sticks and labeled; Asp 206 is a critical residue involved in substrate binding. 

    The binding interactions of TvL with a wide variety of aromatic substrates are shown schematically in Figure \(\PageIndex{11}\). 

    A structural-chemical explanation of fungal laccase activityFig2.svg

    Figure \(\PageIndex{11}\):  Binding modes of representative compounds for TvL.  Mehra, R., Muschiol, J., Meyer, A.S. et al. A structural-chemical explanation of fungal laccase activity. Sci Rep 8, 17285 (2018).  Creative Commons Attribution 4.0 International License.

    Most substrates interact with the highly conserved His-458 (blue color, ligand for the Ti Cu)and Asp-206 (orange color)  residues and form hydrogen bonds, salt bridges, or π-π stacking interactions with them. Asn-264 (blue) and Phe-265 (green) form important hydrogen bonding and π-π stacking interactions with substrates. The green color highlights nonpolar side chains. Ligands bind near the Ti Cu in domain 3 to initiate electron transfer. The active site of TvL must be dynamic to bind the various-sized ligands shown above. Molecular dynamics simulations, shown in Figure \(\PageIndex{12}\), support this.

    A structural-chemical explanation of fungal laccase activityFig8Act.svg

    Figure \(\PageIndex{12}\):  Display of molecular dynamics simulations showing the loop regions of TvL (magenta colored) and another laccase, CuL (yellow colored), and high levels of fluctuations. Mehra, R., ibid

    Breaking down Cellulose

    We just explained how the lignin barrier could be degraded so that cellulase can access cellulose. As we described above, that also poses a difficult challenge given the stability of inaccessibility of the glucosidic bonds in cellulose.   The inaccessibility of "naked" cellulose fibers stems partly from the tight binding of cellulose strands into crystal lattices. Multiple crystal forms of cellulose, called polymorphs, can form. Plant cellulose has two predominant polymorphs, cellulose Iβ and Iα. Their structures are shown below in Figure \(\PageIndex{13}\). 


    Figure \(\PageIndex{13}\): Natural and synthetic cellulose polymorphs. Christina M. Payne et al. Chem. Rev. 2015, 115, 3, 1308–1448 (2015), Open access through a Creative Commons public use license.

    They both form hydrogen bonds within a layer, with the main differences resulting from interlayer stacking. There are no hydrogen bonds between layers. You might find that surprising at first glance until you remember that all the OH groups in the lowest energy chair form of the glucose are equatorial, which allows intralayer hydrogen bonding. The interactions between layers predominantly arise from Van der Waals interactions, specifically induced dipole-induced dipole interactions. The hydrophobic planes, arising from axial H atoms projecting above and above each planar layer of the cellulose fibers, can be readily seen in Figure \(\PageIndex{14}\).


    Figure \(\PageIndex{14}\):  Hydrophobic planes arising from axial H atoms projecting above and above each planar layer of the cellulose fibers. Akira Isogai et al. Progress in Polymer Science, 86 (2018), Creative Commons license

    Now we can explore the structure of cellulases and how they bind to and cleaves cellulose. 

    Cellulases, which cleave β(1,4) glycosidic bonds in cellulose, are members of a family of enzymes that go by many names, including glycosidases, or more recently, glycoside hydrolases (GH). The Carbohydrate Active Enzymes (CAZypedia) has over 128 glycoside hydrolase (GH) family web pages with enzymes that form hemiacetals on the cleavage of glycosidic bonds. The fungal cellulases that work on cellulose are found in GH families 5, 6, 7, 12 and 45.  

    There are many types of secreted or cell-surface cellulases, including endoglucanases, exoglucanases (example is cellobiohydrolases (CBHs), and β -glucosidase (BG) ). We will focus on cellobiohydrolases (CBHs), the most studied one, which cleaves a 2-glucose unit (cellobiose) from either end of cellulose as it proceeds (processes) along the chain. Fungal and bacterial CBHs can work on crystalline cellulose as well. The resulting cellobiose is further cleaved by β-glucosidases. Ruminants and even termites obtain cellulases from microbes living within their guts.   The enzyme has a "tunnel" between two surface loops which interacts with and processively cleaves cellulose.  

    As mentioned above, fungi are the major degraders of biomass, and are critical in the carbon cycle. Some fungi(brown-rot) use the Fenton reaction (Chapter 13.3) to produce the very reactive hydroxyl free radical (.OH) which causes biomass degradation. Filamentous fungi (like white and soft rot like T. reesei ) use enzymes.   The T. reesei cellulase called cellobiose hydrolase 1 is more recently named TrCel7A as it is in the GH 7A family.

    Cellulase mechanism.

    We have previously described the mechanisms of polysaccharide synthesis (Chapter 20.3), so we will discuss in less detail the mechanism of the very similar reaction of cellulose degradation by cellulase. Two general mechanisms are possible, one leading to the retention of configuration at the resulting hemiacetal end of the cellobiose and another that inverts the configuration. These mechanisms are shown in  Figure \(\PageIndex{15}\) for glucose in alpha-linkage at the anomeric carbon (not the beta-linkage found in cellulose).


     Figure \(\PageIndex{15}\): Two Primary Catalytic Mechanisms of GHs.  After Payne et al. Chem. Rev. 2015, 115, 3, 1308–1448.

    Scheme (A) shows the reaction that inverts the configuration. Water acts as a nucleophile in a SN2 type of reaction, with catalytic assistance by two proximal carboxylic acid side chains acting as general acids and bases. This results in an inversion of the stereochemistry at the anomeric carbon.

    Scheme (B) proceeds with the retention of configuration as two different nucleophilic attacks occur. In the first, an active site carboxylate forms a covalent acetal intermediate with the anomeric carbon. The carboxylate hence acts as a nucleophilic catalyst. Water, acting as a nucleophile, then attacks to form the hemiacetal with the expulsion of the carboxylate leaving group. As we discussed in Chapter 20.3 (section on glycosyl transferases), other variants of these mechanisms would include a SN1 reaction or one with an oxocarbenium-like transition state.  

    The CBH1 (family 7) has a long tunnel for binding cellulose. The CBH1 (TrCel7A) cellulose catalytic site spans at least 9 glucose monomers (n-7, n-6,...,n-1,n+1, n+2) with cleavage typically of a cellobiose from the reducing end (between n-1 and n+1). The structure of the TrCel7A glycoside hydrolase (cellobiose hydrolase) with a small bound cellulose is shown in  Figure \(\PageIndex{16}\).


    Figure \(\PageIndex{16}\):  Crystal structure of the first GH7 CBH and EG.  Payne et al. Chem. Rev. 2015, 115, 3, 1308–1448. Open access through Creative Commons public use license.

    The ligand from the TrCel7A Michaelis complex (PDB code 4C4C (441)) is shown in all panels. (A) CBH TrCel7A CD (PDB code 1CEL (172)) view from side, exhibiting the β sandwich structure that is characteristic of GH7 enzymes. TrCel7A was the first GH7 structure solved and is the best-characterized member of GH7. (B) TrCel7A view from bottom showing the more closed substrate binding "tunnel". (C) EG F. oxysporum Cel7B (PDB code 1OVW (174)) view from side. (D) FoCel7B view from the bottom showing the more open binding "groove". (E) TrCel7A Michaelis complex (PDB code 4C4C (441)) shows the standard numbering of the substrate binding sites (catalytic residues shown in green for reference). A cellulose chain enters from the −7 site. Hydrolysis occurs between the −1 and +1 sites. The +1/+2 sites are termed the "product sites".

    Active site carboxylates (E212, D214, and E217) are shown near the -1/+1 cleavage site in Figure \(\PageIndex{17}\). Glu 217 is covalently attached to the -1 glucose, supporting the retaining mechanism illustrated in Fig 15 above.


    Figure \(\PageIndex{17}\): Michaelis complex and glycosyl-enzyme intermediate of TrCel7A. Payne et al. ibid.

    Panel (A) shows the TrCel7A Michaelis complex (PDB code 4C4C (441)). 

    Panel (B) shows a TrCel7A glycosyl-enzyme intermediate (PDB code 4C4D (441)) with a covalent bond between the nucleophile and the broken cellooligomer chain. There is an approximate 30° rotation of the E217 nucleophile during glycosylation.

    Figure \(\PageIndex{18}\) shows a more detailed view of the first step (glycosylation f Glu 217) for the Hypocrea jecorina GH Family 7 cellobiohydrolase Cel7A

    The Mechanism of Cellulose Hydrolysis by a Two-Step Retaining mechFig2.svg

    Figure \(\PageIndex{18}\): Figure 2. Glycosylation step for Hypocrea jecorina GH Family 7 cellobiohydrolase Cel7A.   Knott, Brandon C. et al. - J. Am. Chem. Soc.329 (2013) Open access article published under an ACS AuthorChoice License

    Panel (a) shows a  snapshot of the reactant the conformation from a representative AS trajectory (with the substrate in green and catalytic residues in yellow) for the glycosylation step. The proton resides on the acid residue, Glu217.

    Panel (b) shows a representative snapshot of the transition state. The −1 glucopyranose ring now adopts a different conformation.

    Panel (c) shows the product of the glycosylation reaction.

    Panel (d) shows a schematic view of the overall glycosylation reaction with the collective variables identified by LM colored at the transition state. The best three-component RC identified by LM includes the forming/breaking bonds involving the anomeric carbon, the breaking bond between Glu217 and its proton, and the orientation of the nucleophile Glu212. 

    Figure \(\PageIndex{19}\) shows the corresponding deglycosylation (of Glu 217) step.

    The Mechanism of Cellulose Hydrolysis by a Two-Step Retaining mechFig4.svg

    Figure \(\PageIndex{19}\): Figure 4. Deglycosylation step results. Knott, Brandon C. et al, ibid

    Panel (a) shows a snapshot of the reactant conformation from a representative AS trajectory (with the substrate in green and catalytic residues in yellow) for the deglycosylation step. The covalent glycosyl–enzyme bond is intact, and the cellobiose product is in primed GEI mode. 

    Panel (b) shows a representative snapshot of the transition state. Note the distorted conformation of the −1 sugar, as the nucleophilic water molecule is ripped apart.

    Panel (c) shows a snapshot of the product in which the glycosyl-enzyme bond has been broken, and the catalytic residues have been regenerated.

    Panel (d) shows a schematic view of the overall deglycosylation reaction with the collective variables identified by LM colored at the transition state. The best three-component RC identified by LM includes the forming/breaking bonds involving the anomeric carbon, the forming/breaking bonds involving the transferring proton, and the orientation of the C3 hydroxyl of the +1 sugar.

    Binding of cellulase to cellulose fibers and lignin

    Many glycoside hydrolases contain distinct carbohydrate binding domains/modules (CBD/CBM) and catalytic domains (CD). For example, TrCel7A can be cleaved by the protease papain into a 56K domain with catalytic activity on small substrates but not large cellulose one and a smaller 10K (C terminal) domain that itself is glycosylated and which binds to the hydrophobic surface of cellulose crystals.  

    Many GHs, in addition, have linkers connecting the catalytic domain (CD) and the carbohydrate module (CBM), which add different functions to the enzymes. The linkers vary in size and amino acid sequence. Linkers in fungi tend to be long and N- and O-glycosylated, affecting binding/catalysis. The linkers can also be intrinsically disordered, which adds dynamic complexity to their effects.

    The actual cellulose binding site on cellulase has been determined by solution NMR using a synthetic 36 amino acids protein fragment from the C-terminal domain of Trichoderma reesei Cel7A (the "carbohydrate binding module or CBM"). The amino acids involved in the binding of cellohexaose (6-mer) were determined by perturbation of the 2D NMR structure on binding cellohexaose. As we mentioned above, cellulase also binds lignin, decreasing their catalytic efficiency towards cellulase. Results of NMR binding studies of the TrCel7A carbohydrate binding module with cellohexaose and lignins from Japanese cedar (C-MWL) and Eucalyptus globulus (E-MWL) are shown in Figure \(\PageIndex{20}\).

    NMR Analysis binding lignin cellulosetocellulaseFig8.svg

    Figure \(\PageIndex{20}\): Comparison of interaction property between cellohexaose and MWLs.  Tokunaga, Y., Nagata, T., Suetomi, T. et al. NMR Analysis on Molecular Interaction of Lignin with Amino Acid Residues of Carbohydrate-Binding Module from Trichoderma reesei Cel7A. Sci Rep 9, 1977 (2019).  Creative Commons Attribution 4.0 International License.

    Panel (a) shows cellohexaose specifically bound to the flat plane surface and cleft.   The flat plane surface is defined by a triplet tyrosine (Y5, Y31, Y32) and H4, G6, Q7, I11, L28, N29, Q34, L36.

    Panel (b) shows both MWLs bound to multiple binding sites, some of which are included in the flat plane surface and cleft even in low concentrations of titrant. These non-specific binding sites are labeled green.  

    Figure \(\PageIndex{21}\) shows an interactive iCn3D model of the C-terminal cellulose-binding module of cellobiohydrolase I from Trichoderma reesei (2CBH).

    C-terminal domain of cellobiohydrolase I from Trichoderma reesei (2CBH).png

    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{21}\): C-terminal cellulose-binding module of cellobiohydrolase I from Trichoderma reesei (2CBH). (Copyright; author via source). Click the image for a popup or use this external link:

    This synthetic carbohydrate binding module (CBM) from the C-terminal domain of cellobiohydrolase I consists of 36 residues. NMR was used to determine the structure of the CBM.  2D NMR was used to determine the amino acids interacting with cellulose. The interacting side chains are shown as sticks underneath the molecular surface (gray). The side chains are colored according to hydrophobicity, with green followed by yellow being most hydrophobic. The numbering system refers to the 36 amino acid synthetic peptide, not the native protein. This model clearly shows this domain binds to the hydrophobic face of the cellulose microcrystal.

    The TrCel7A CBM has serine and threonine side chains that are glycosylated and affect binding. The interaction of the CBM with the nonpolar cellulose surface is shown in Figure \(\PageIndex{22}\).


    Figure \(\PageIndex{22}\): Glycosylated TrCel7A CBM on the hydrophobic surface of cellulose. Payne et al., ibid.

    Note that the aromatic groups of the triplet tyrosines, Y5, Y31, and Y32 (not labeled), are coplanar with the cellulose surface.

    As mentioned above, linkers that connect the C-terminal carbohydrate binding module (CBM) and the catalytic domain (CD) can be of different lengths and sequences and are also N- and O-glycosylated.  Figure \(\PageIndex{23}\) shows the interactions of the glycosylated linkers with the cellulose fibers.


    Figure \(\PageIndex{23}\): Molecular snapshots of TrCel7A and TrCel6A wherein the linker binds to the cellulose surface from microsecond-long MD simulations. Payne et al., ibid.

    These computational predictions of cellulose linkers enhancing binding of CBMs to the cellulose surface were corroborated experimentally via binding isotherm measurements. N-glycosylation and O-glycosylation are shown in blue and yellow. The glycans attached to the enzyme significantly enhance the binding of cellulase to the cellulose fibers. Payne et al. Chem. Rev. 2015, 115, 3, 1308–1448. Open access through Creative Commons public use license

    A pictorial view of the hydrolytic cleavage of cellobiose from cellulose fibers is shown in Figure \(\PageIndex{24}\).  




    Figure \(\PageIndex{24}\): Complete processive cycle of a GH7 CBH. TrCel7A is shown with its CD, linker, and CBM in gray "cartoon" representation. Payne et al., ibid.

    N-glycosylation and O-glycosylation are shown in blue and yellow, respectively. The cellulose surface is green, and the released cellobiose product magenta. Following the CBM and CD adsorption to the substrate and initial chain threading, TrCel7A processively cleaves cellobiose from a cellulose chain end. The "Processive Cycle" includes chain processivity, hydrolysis, and product expulsion (Figure 35). This processive cycle repeatedly occurs until the enzyme desorbs from the cellulose surface.

    Figure \(\PageIndex{25}\) shows an interactive iCn3D model of cellulose bound  to cellobiohydrolase I from Trichoderma reesei (7CEL)


    Cellulose chain is bound in the 50 A long tunnel of cellobiohydrolase I from Trichoderma reesei (7CEL).png


    NIH_NCBI_iCn3D_Banner.svg Figure \(\PageIndex{25}\): Cellulose bound to cellobiohydrolase I from Trichoderma reesei (7CEL). (Copyright; author via source). Click the image for a popup or use this external link:


    Key Points - Beta version from Chat.openai
    1. Cellulosic ethanol is a biofuel that is produced from the cellulose, hemicellulose, and lignin in plant material.
    2. Unlike first-generation biofuels like corn and sugar cane ethanol, which are produced from sugars and starches, cellulosic ethanol can be produced from a wide range of plant material, including agricultural waste, wood chips, and switchgrass.
    3. Cellulosic ethanol is considered a "second-generation" biofuel because it addresses many of the limitations of first-generation biofuels, including the competition with food crops for land and resources.
    4. Cellulosic ethanol production involves a two-step process: first, the plant material is broken down into sugars through a process known as pretreatment, and then the sugars are fermented to produce ethanol.
    5. The most common pretreatment methods include acid hydrolysis, ammonia fiber expansion, and steam explosion.
    6. Cellulosic ethanol has a higher energy balance than first-generation biofuels and lower greenhouse gas emissions, making it a more sustainable option for biofuel production.
    7. However, the technology for cellulosic ethanol production is still in the early stages of development, and the cost of production remains high.
    8. Research is ongoing to improve the efficiency and cost-effectiveness of cellulosic ethanol production and to find ways to make it a viable alternative to fossil fuels.

    This page titled 32.5: Biofuels B - Cellulosic Ethanol is shared under a not declared license and was authored, remixed, and/or curated by Henry Jakubowski.

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