Cellulose solvent-based pretreatment for enhanced second-generation biofuel production: a review

Abstract

Cellulose, in addition to hemicellulose and lignin, makes the major fraction of lignocellulosic biomass – the only sustainable feedstock to meet the long-term sustainable energy need of the world. Cellulose is soluble in a number of solvents, e.g., concentrated phosphoric acid (CPA), N-methylmorpholine-N-oxide (NMMO), and ionic liquids (ILs), and can be regenerated by an anti-solvent without major derivatization for various applications. For one, the regenerated and much less crystalline cellulose is highly reactive for its biological conversion to sugars, fuels, and chemicals mediated by enzymes and/or microbes. This ability can be used as a core pretreatment step for improved bioprocessing of lignocelluloses. In this comprehensive review, cellulose solvent-based lignocellulosic fractionation technologies for enhanced enzymatic hydrolysis to improve biofuel and renewable chemical production are reviewed. The first part is focused on the background information of lignocellulosic biomass, lignocellulosic derived biogas, biohydrogen, and ethanol as well as acetone, butanol, and ethanol (ABE) production, and enzymatic hydrolysis. In the second part, the conditions for pretreatments applying CPA, NMMO, and IL solvents, improvements in enzymatic hydrolysis rates and yields for solids resulting from application of these pretreatments, and the features of the lignocellulosic structure affecting the improved bioprocessing have been thoroughly reviewed.

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1 Different generations and types of biofuels

Recent concerns about climate change due to greenhouse gas emissions and energy crisis have prompted the need for transition from unsustainable fossil-derived energies to sustainable and renewable energies.¹

Development of sustainable and economically viable biorefinery processes for biofuel production needs to use renewable carbon sources.² Biofuels produced from food-based crops like sugar- and starch-based substrates, e.g., sugarcane and corn, are considered as first-generation biofuels.^3,4 Nevertheless, there is a food-versus-fuel debate in using feedstocks for first-generation fuels. Therefore, next-generation biofuels were introduced and are considered as essential for meeting the world's energy demand in the transportation sector.^5–7

Second-generation biofuels are produced from lignocellulosic biomass, which can reduce the carbon emission, increase energy efficiency, and reduce nations' energy dependency.^3,7–9 Non-food lignocellulosic substrates are abundant and potentially low-cost organic sources for renewable chemical and fuel production. Lignocellulosic wastes can be originated from industrial wastes (e.g., sawdust, paper mill discards, and food industry wastes), forestry wastes (i.e., hardwoods and softwoods), agricultural residues (e.g., straws, stover, and non-food seeds), domestic wastes (e.g., kitchen wastes, sewage, and waste papers), and municipal solid wastes.^10–12

Third-generation biofuels are produced from algae.^13,14 Biofuel production from algal species, including Botryococcus braunii, Chaetoceros calcitrans, several Chlorella species, Isochrysis galbana, Nannochloropsis, Schizochytrium limacinum, and Scenedesmus species, is a promising technology since algae are fast growing, compared to many terrestrial plants, with no soil needed, while they have high capturing ability for CO₂ and other greenhouse gases.¹⁵ Algae contain substantial amounts of carbohydrates and lipids (up to 70%), making them promising feedstocks for conversion to biofuels, e.g., by simple hydrolysis followed by fermentation or consolidated bioprocessing.¹⁶ A comprehensive overview of the composition, properties, and challenges of algal biomass for biofuel application was recently presented by Vassilev and Vassileva.¹⁷ Biodiesel, bioethanol, biohydrogen, and biogas were reported to be produced from micro- and macro-algae via different technologies.^13,14

Fourth-generation biofuels use engineered algae for biofuel production from oxygenic photosynthetic organisms.¹⁸ Gaseous biofuels, algal ethanol, algal butanol, four carbon alcohols, and algal biodiesel were reported to be possible to produce by using this technology.¹⁸

Nonetheless, the production cost of biofuels is extremely sensitive to the feedstock cost.¹⁹ Although algae do not need freshwater and can grow on wastewater streams (e.g., saline/brackish water/coastal seawater), harvesting and carbon supply are the major factors of algal biomass production cost.²⁰ Harvesting microalgae usually needs flocculation to aggregate small algal cells followed by filtration, centrifugation, and sedimentation to separate the algae from the liquid medium. Besides, advanced and cheaper technologies are required for the extraction of algal oil. Although the land use is low for algal cultivation, infrastructure requirements, and mixing and separation costs are still high. Moreover, the high cost of edible crops and land requirements to meet the demand make them unsustainable. Therefore, lignocellulosic biomass is the only sustainable and low-cost feedstock to meet the near future growing energy needs and mitigate environmental problems.^20,21

Regarding environmental impacts, all types of biofuels reduce greenhouse gas (GHG) emissions²² (Fig. 1). Life cycle assessment for biofuel production from different sources was performed and the net GHG emission for different fuels, e.g., fossil fuel, soya oil biodiesel, palm biodiesel, sugarcane ethanol, wheat ethanol, corn ethanol, corn stover ethanol, and algal biodiesel, was compared.^21,23 It was shown that among them corn stover derived ethanol released the lowest net GHG emission.

Fig. 1 A simplified diagram showing a neutral carbon cycle for biofuel production from plants.

Less (or negligible) competition to food, production of value-added byproducts, and energy security are among the advantages of second-generation biofuels. As shown schematically in Fig. 2, the main steps for second-generation biofuel and chemical production are usually substrate preparation, including size reduction and pretreatment, carbohydrate saccharification, fermentation, and product separation and purification.²⁴ The processing cost for second-generation ethanol is approximately two to three times higher than that for gasoline on an energy equivalent basis;²⁵ therefore, substantial attention has been recently focused on the improvement of process economy and technology development to make second-generation biofuels economically viable.

Fig. 2 Schematic of the production of various chemicals from lignocellulosic feedstocks (second-generation biofuels and chemicals).

1.1 Bioethanol

Ethanol, blended with gasoline or as a neat fuel in vehicles, is an attractive transportation fuel, giving a high octane number and heat of vaporization.²⁶ Currently, ethanol mainly produced by fermentation routes using sugar- and starch-based feedstocks, e.g., sugarcane and maize, is called first-generation ethanol²⁷ (Fig. 3). Following fermentation, ethanol is separated and purified from the fermentation broth via distillation and molecular sieves, respectively.²⁸ The industrial technology for the fermentation of glucose to ethanol is quite robust and high concentrations of ethanol (12–15%) can be achieved.²⁹ In the production of ethanol from starch, an extra step of liquefaction and saccharification by α-amylases and glucoamylases, respectively, is necessary for converting starch to sugar³⁰ (Fig. 3). Since the production capability of the first-generation ethanol is limited and is unsustainable on a large scale, second-generation ethanol was then introduced, which utilizes a variety of lignocelluloses as substrates.^27,31 (Fig. 3).

Fig. 3 Conversion of different feedstocks to ethanol via the fermentation route. The conversion from sugar- and starchy-based materials to ethanol is called first-generation and production from lignocelluloses is called second-generation.

1.2 Biobutanol

For gasoline blending, butanol, a four-carbon alcohol, is more desirable than ethanol due to its higher energy density, lower hygroscopicity, lower Reid vapor pressure, better blending ability, and use in conventional combustion engines without modification.³² Besides the fuel extender, biobutanol can be used as a feedstock for the synthesis of a variety of commercial products.^33,34 The fermentative route of production, e.g., by microorganisms that belong to the genus Clostridium, is more sustainably and environmentally attractive than the petrochemical route.³⁵ These microorganisms typically produce a mixture of different solvents, mainly including acetone, ethanol, and butanol; thus, the process is referred to as acetone–butanol–ethanol (ABE) fermentation.^36,37 However, the major challenge in the microbial production of butanol is the low butanol titer due to product inhibition.^38,39 Several strategies have been reported to address these issues⁴⁰ such as genetic and metabolic engineering of microorganisms⁴⁰ and promising integrated continuous culture technology with efficient product recovery techniques, e.g., using metal–organic frameworks,⁴¹ liquid–liquid extraction,^42–44 pervaporation technique,⁴⁵ and gas stripping.⁴⁶

Butanol can be synthesized via different metabolic and engineered pathways from different substrates. Starch/sugars can be converted to butanol via the clostridial route that includes glycolysis, pyruvate:ferredoxin oxidoreductase, thiolase, 3-hydroxybutyryl-CoA dehydrogenase, crotonase, butyryl-CoA dehydrogenase, and butyraldehyde/butanol dehydrogenase. The conversion of lignocellulosic feedstocks to biobutanol also follows the same route after being converted to C₅ and C₆ sugars in the preceding pretreatment and/or enzymatic saccharification steps. Lignocellulosic biobutanol production has received a lot of attention, and it has recently been the focus of vast studies.^47,48 However, the low butanol titers and yields and requirement of extra pretreatment and enzymatic saccharification steps are some of the challenges in butanol production from lignocellulosic biomass. Moreover, syngas or CO₂/H₂ can also be fermented to butanol via the clostridial pathway.⁴⁹ For starch and sugars, there is another non-fermentative pathway based on amino acid metabolism plus 2-keto acid decarboxylase and alcohol dehydrogenase.³⁶ Aerobic butane-utilizing bacteria use monooxygenase to oxidize butane to a butanol mixture (95% butanol, 5% iso-butanol).³⁶

1.3 Biodiesel

Biodiesel, a mixture of fatty acid methyl esters (FAMEs), can be produced by transesterification of vegetable oil or animal fat. It recently received much attention as a renewable source of energy.^50,51 However, these resources for biodiesel production do not meet the large-scale demands for transportation fuel and a sustainable renewable source is required.

Nonetheless, some microorganisms, called oleaginous microorganisms, can store intracellular lipids, usually referred to as single cell oil (SCO), especially triacylglycerols (TAGs).⁵¹ Microbial oil, as a raw feedstock for biodiesel production, is advantageous compared to vegetable oil because it has a short life cycle, requires less labor, is less affected by venue, season, and climate, and is easier to scale up.⁵² Different oleaginous microorganisms, including microalgae, yeasts, fungi, and bacteria, were reported to produce substantial amounts of SCO (e.g., 20–50% dry cell weight).^53–55 However, it is possible to increase lipid accumulation in oleaginous microorganisms via metabolic engineering technology, involving the enhancement of fatty acid synthesis approaches, enhancement of TAG synthesis approaches, regulation of related TAG biosynthesis bypass approaches, blocking of competing pathways, and multigene approaches.⁵⁶

A variety of carbon sources from lignocellulose-based carbohydrates and other low-cost industrial wastes, e.g., glycerol, food processing waste, and even wastewater, have been reported to be assimilated by oleaginous microorganisms to produce lipids.^57–61 Auxiliary nutrients such as phosphorous and nitrogen are available from the waste streams. However, lipid accumulation in oleaginous microorganisms is usually triggered by nutrient starvation, e.g., nitrogen or phosphorus, relative to the carbon source.⁶²

Lipid production from lignocellulosic biomass has attracted substantial attention in recent years and many research studies have focused on its commercialization; however, substantial process improvements and reduction in the production cost are required.^63–66

1.4 Biogas

Besides liquid biofuels, biomass with high organic content can be converted to another form of energy, biogas, via anaerobic digestion (AD). In this process, the organic matter is biologically decomposed by an assortment of microbes under oxygen-free conditions, producing biogas (about 50–75% CH₄ and 25–50% CO₂).^67,68 The AD process can be divided into four steps: (i) hydrolysis of proteins and lipids to amino acids and long-chain fatty acids and carbohydrates into sugars, (ii) conversion of hydrolysis products and monomers to volatile fatty acids (VFAs) and other minor products such as alcohol by acidogenic bacteria, (iii) conversion of VFAs to acetate, carbon dioxide, and/or hydrogen by acetogenic bacteria, and (iv) methane formation from other stage products by methanogenesis⁶⁹ (Fig. 4). Although methanogenesis is usually considered as the rate-limiting step in the AD process for a number of substrates, the hydrolysis step is believed to be the limiting step for lignocelluloses. Sawatdeenarunat et al.⁷⁰ classified the current technologies in the AD process of lignocellulosic biomass into anaerobic co-digestion, solid-state anaerobic digestion (SS-AD) (more than 15% TS) and using alternative biological pretreatment of feedstock for further biological conversion to sugars, e.g., by using rumen microorganisms.

Fig. 4 The main steps in the degradation of organic matter through the anaerobic digestion process.⁷¹

1.5 Biohydrogen

Biologically produced hydrogen, biohydrogen, is recently becoming of great interest as a renewable energy carrier, because hydrogen utilization for combustion, in fuel cells, and/or electricity production produces no carbon byproducts.⁷² Biological pathways for hydrogen production are primarily divided into photobiological processes and light independent methods.^73,74 Green algae from the genera Chlamydomonas, Scenedesmus, Lobochlamys, and Chlorella can reduce protons of water in the presence of light to produce mixed oxygen and hydrogen gases.⁷⁵ Some photosynthetic bacteria were also reported to produce hydrogen by the same mechanism of biophotolysis as that by green algae. Fermentative biohydrogen production, classified as photofermentation and dark fermentation, can be performed by a wide variety of microorganisms, e.g., strict anaerobes, facultative anaerobes, and aerobes kept under anoxic conditions.^73,76 Fermentative hydrogen production is more advantageous over the photosynthetic method since various organic feedstocks can be converted to hydrogen with high production rates and simple operations.⁷⁷ Several factors, including inoculum, i.e., mixed and pure cultures, substrates, reactor type, availability of nitrogen and phosphate micro-nutrients and metal ions, temperature, and pH, were reported to influence fermentative hydrogen production.^78,79 Because of the higher hydrogen evolution rate, dark fermentation hydrogen production is more commercially feasible than photofermentation. In dark fermentation, organic substrates like glucose are converted by facultative and obligate anaerobes to hydrogen, volatile fatty acids, and carbon dioxide operated at mesophilic, thermophilic, or hyperthermophilic temperatures in the absence of light.⁸⁰ The knowledge of biological pathways for dark fermentation hydrogen production is quite mature and is comprehensively presented in the literature.^{73,75,76,80–85} Here a brief discussion on the strategies to enhance biological hydrogen production and the feedstocks is presented.

Different carbon sources, e.g., agricultural residues, industrial waste, organic fraction of municipal waste, and pure sugars, were reported as feedstocks for biohydrogen production.^72,86,87 Lignocellulosic feedstocks are promising raw materials for biohydrogen production and recently have been the focus of a number of studies.⁷² Different approaches for bioconversion of lignocellulosic biomass to H₂, i.e., separate hydrolysis and fermentation, simultaneous saccharification and fermentation, and consolidated bioprocessing of lignocellulosic biomass to H₂, have been discussed by Cheng et al.⁷² and Ren et al.⁸⁸ Application of various pretreatment technologies for enhanced lignocellulosic bioconversion to biohydrogen has also been the topic of several studies.^89–93

While the theoretical hydrogen yield is 12 mole H₂ per mole of glucose, natural and genetically modified microorganisms can produce hydrogen at a maximum yield of 4 mol mol⁻¹ glucose when acetic acid is the only VFA product.⁸⁵ The strategies for biohydrogen production improvement include microbial culture immobilization, bioreactor modifications, optimization of operating parameters (i.e., temperature, pH, organic loading rate, hydrolytic retention time, and H₂ partial pressure), substrate type and inorganic nutrients, metabolic engineering of microbes, and cogeneration of biohydrogen and biomethane.^{73,78,81,94,95}

The inoculum for dark fermentation biohydrogen production can be either pure cultures or anaerobic microbial consortia. Mixed culture is generally preferable because of the ease of operation, no need for sterilization, and, especially for lignocelluloses, the presence of hydrolytic activities.⁹⁶ In such systems, methanogenesis activity can be easily eradicated by a heat shock or pH control, and the hydrogen-producing bacteria can sporulate.^74,97

Another noteworthy approach based on cell-free hydrogen production was originally proposed by Dr Jonathan Woodward at Oak Ridge National Laboratory,^98,99 and then has recently been revived by Ye et al.¹⁰⁰ and Zhang et al.¹⁰¹

2 Lignocellulosic biomass structure

Lignocelluloses typically contain lignin, carbohydrate polymers (∼75%; i.e., cellulose, hemicellulose, and pectin), acetate, proteins, salt, ash, and minerals.¹⁰²Table 1 summarizes the major composition (carbohydrates and lignin) of some lignocelluloses used for second-generation biofuel production. Being the nature's most abundant organic substance after cellulose, lignin comprises 28–30% of woody gymnosperm stems and 20–24% of woody angiosperms.¹⁰³ The lignin composition varies between hardwoods and softwoods. Lignin has a heterogeneous three dimensional β-O-4, β-5, β-1, β-β, 5–5, and 4-O-5 linked structure of phenylpropane units, e.g., p-hydroxycinnamyl, p-coumaryl, coniferyl, guaiacyl, syringyl, and sinapyl alcohol.^102,104 Lignin acts as a cement to hold the cell components together and provides the biomass integrity.¹⁰⁵

Table 1 Composition (based on % dry weight) of some widely used lignocelluloses for second-generation biofuel production^a

Biomass type	Substrate	Glucan	Xylan	Mannan	Galactan	Arabinan	Lignin			Ref.
							Total	Acid insoluble	Acid soluble
a The carbohydrate contents were measured by analyzing the sugars released during a concentrated sulfuric acid (72%) hydrolysis at 30 °C followed by a dilute acid treatment at 121 °C to cleave the carbohydrates to monomeric sugars. Acid-insoluble lignin was measured gravimetrically after subtracting the ash content of final acid-insoluble materials.^128
Hardwood		41.7	14.3	2.6	3.2	2.0	30.2			114
		45.2	20.3	4.2	—	—		21.0	3.3	115
		39.2	13.1	1.8	0.9	—	14.7			116
		40.3	16.9	1.7	0.6	0.3	20.3			117
		49.0	14.9	2.0	0.5	0.8		24.6	1.0	118
		43.6	20.3	1.5	1.2	—	26.3			119
Softwood		47.3	4.4	11.7	2.2	1.2		29.8	0.5	120
		38.2	8.5	11.3	4.3	—		29.5	4.9	121
		38.5	5.0	11.7	1.9	1.3	28.5			117
Agricultural residues and grasses		32.0	17.9	—	1.73	1.78	21.4			122
		37.4	22.4	—	0.51	6.2		13.2	1.9	123
		38.8	22.2	1.7	2.7	1.4	18.5			124
		40.87	20.82	—	—	1.53	24.81			125
		35.3	23.9	—	1.9	4		19.2	0.7	126
		41.33	17.96	0.85	1.26	1.96		16.4	1.78	127

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Cellulose, with over 10¹¹ metric tons production per year, is composed of linear chains of several hundreds to over tens of thousands of β-D-glucopyranose residues linked by β-1,4 glycosidic bonds with numerous inter- and intra-molecular hydrogen bonds.¹⁰⁶ It is a ubiquitous polysaccharide of the plant cell wall (Fig. 5), which makes it insoluble in water and common organic solvents.^104,107,108 Aggregation of cellulose chains forms nanofibrils and a 5–10 nm microfibril, hypothesized to be composed of 36 chains of cellulose, is used to define the next level of aggregation, which is observable via high magnification microscopy, e.g., electron microscopy, and atomic force microscopy^109–112 (Fig. 5). Cellulose is the dominant component of the primary cell wall (20–40% of cell wall dry matter).¹¹³ The research on cellulose revealed that native celluloses are crystalline and are composites of two forms, I_α (with one-chain triclinic structure) and I_β (a two-chain monoclinic structure), which coexist in all native forms.¹⁰⁷

Fig. 5 (A) Pictorial illustration of the lignocellulosic biomass framework (modified from Menon and Rao¹³⁷ with permission), (B) a simplified model showing the interaction of carbohydrate polymers present in the cell wall, modified from Himmel et al.,¹³⁸ (C) structure of a 36-chain model for the cellulose I_α or I_β elementary fibril (the reds show six true crystalline chains; greens are 12 subcrystalline chains with a small degree of disorder; the blues are 18 surface chains that are subcrystalline with a large degree of disorder, taken from ref. 111), and (D) a model of inter- and intra-chain hydrogen-bonding patterns in cellulose, taken from ref. 139 with permission.

Hemicellulose, a stereo-irregular polysaccharide, is a heterogeneous plant cell wall polymer composed of linear β(1,4)-D-glycan backbones branched with one monosaccharide and/or small oligosaccharides, with an approximate degree of polymerization of 200.^129–131 Unlike cellulose, hemicellulose has an amorphous, random, and branched structure, which is more susceptible to thermal, biological, and acid hydrolysis.^132–135 Xylan, mainly in the form of heteroxylan, is usually substituted with acetate and arabinose residues. It is the most abundant hemicellulose in nature, which dominantly contains β-D-xylopyranosyl residues linked by 1,4 glycosidic bonds.^102,104 The xylan content of the plant cell wall may vary depending on the biomass type, ranging between 15 and 35% of total dry weight.¹⁰² Hemicellulose interacts with cellulose and lignin and build a rigid network structure which is a barrier to enzyme-catalyzed deconstruction of cellulose.¹³⁶

Pectin (pectic polysaccharides) is a heterogeneous polysaccharide with a dominantly methyl esterified or de-esterified homogalacturonan (HG) backbone. Located in the cell wall and middle lamella of plants, pectin is the major component of the primary walls of several non-woody plant cells.^140,141 After cellulose, pectin acts as a major plant load-bearing component and plays a “glue” role in holding cell-wall components together.^{138,142–145}

3 Biomass recalcitrance and pretreatment

Lynd et al.¹⁴⁶ first defined the “biomass recalcitrance” as the natural resistance of lignocelluloses and their components to microbial and enzymatic deconstruction. Later, Himmel et al.¹³⁸ summarized the factors contributing to the biomass recalcitrance as “(i) epidermal tissue of plant body, especially cuticle and epicuticular waxes, (ii) the arrangement and density of the vascular bundles, (iii) the relative amount of sclerenchymatous (thick wall) tissue, (iv) the degree of lignification, (v) the structural heterogeneity and complexity of cell-wall constituents such as microfibrils and matrix polymers, (vi) the challenges for enzymes acting on an insoluble substrate, and (vii) the inhibitors to subsequent fermentations that exist naturally in cell walls or are generated during conversion processes”. Due to the biomass inherent recalcitrance, the release of fermentable sugars via appropriate enzymatic hydrolysis as well as microbial hydrolysis is the bottleneck of the industrial lignocellulosic biorefineries.^147,148

Therefore, an efficient pretreatment step is required to obtain renewable chemicals and fuels from lignocelluloses.¹⁴⁹ A suitable enzymatic or acid hydrolysis can then be applied to the pretreated substrates to convert them to fermentable sugars or the AD process can be applied to obtain biogas. There are many reviews in the literature on pretreatment methods to enhance the enzymatic digestibility of lignocellulosic feedstocks.^{11,69,136,150–157} Pretreatment is a “physical”, “chemical”, “physico-chemical”, or “biological” process, which can open up the lignocellulosic recalcitrance structure and make it amenable for subsequent enzymatic/microbial degradation. Physical pretreatments are divided into mechanical comminution and pyrolysis, whereas physicochemical pretreatments are steam explosion, ammonia fiber expansion (AFEX), and carbon dioxide explosion, and chemical pretreatments can be categorized into ozonolysis, acid hydrolysis, alkaline hydrolysis, oxidative delignification, and organosolv processes.^{11,158,638,639} The two most commonly used technologies for pretreatment of lignocelluloses are dilute acid and alkaline pretreatments.¹⁵⁹ Dilute acid and alkaline pretreatments mainly target hemicellulose and lignin fractions, respectively, in lignocellulosic biomass. Acids like HCl and H₂SO₄ and bases like sodium hydroxide and sodium carbonate are mostly employed, and the pretreatment temperature, time, and acid/base concentration are among the main factors determining the effectiveness of pretreatment. An additional process and/or additional chemicals are required for recovering and neutralizing the hydrolysates and removing the inhibitory compounds for downstream processes. Hydrothermal pretreatment with only hot water, which is performed by using saturated steam at temperatures and pressures below the water critical point (subcritical water) or supercritical water, has the advantages of a low amount of biological inhibitor production, minimal chemical cost, and relatively low cost of reactors compared with using acid or alkali solutions. A technology used for hydrothermal pretreatment, called steam explosion, is a pretreatment in which the lignocellulosic biomass is heated up by high-pressure steam (160–240 °C and pressures 0.7–4.8 MPa) followed by an explosive decompression. Hemicelluloses are mostly hydrolyzed in this pretreatment via the reaction called “autohydrolysis”.^160,161

For an advanced and low-cost pretreatment, several key criteria should be considered. It should be effective for a variety of lignocellulosic types with different characteristics. Significant sugar degradation products, formation of inhibitory byproducts for subsequent sugar fermentation, and production of waste residues should not occur during the pretreatment. Moreover, the pretreatment should need minimum heat and power requirement and reasonable size and moderate cost reactors.^150,153,162

An efficient biomass pretreatment strategy should, therefore, be capable of effectively disrupting and removing the linkages among cellulose, hemicellulose, and lignin present in the plant cell walls. Furthermore, reordering or removing highly ordered hydrogen bonds in cellulose fibers and subsequently increasing the porosity and surface area, resulting in cellulose accessibility to cellulase, are highly desirable traits of an effective pretreatment.^150,153,162

Recently, a new pretreatment category based on cellulose solvent lignocellulosic fractionation, meeting the desired criteria, was added to the traditional biomass pretreatments. A number of low toxicity and mostly environmentally friendly solvents, including N-methyl-morpholine-N-oxide (NMMO), ionic liquids (ILs), LiCl/N,N-dimethylacetamide (LiCl/DMAc), aqueous NaOH solution, alkali/urea and NaOH/thiourea aqueous solutions, tetra butyl ammonium fluoride/dimethyl sulfoxide system, metal complex solutions, concentrated phosphoric acid, and molten inorganic salt hydrates, have been introduced as cellulose solvents for regenerating cellulosic materials.^163–165 The cellulose solvents can be classified into (i) derivatizing, (ii) non-derivatizing, and (iii) aqueous and non-aqueous systems having the ability to eliminate the inter- and intra-molecular hydrogen bonds among cellulose molecules.¹⁶⁶ The cellulose can then be recovered using an anti-solvent such as water, ethanol, or acetone. The parallel arrangement of cellulose I, in most regenerated celluloses, is irreversibly converted into an anti-parallel orientation, cellulose II, which is much easier to hydrolyze using cellulases.¹⁶⁷ Cellulose II is thermodynamically more stable and has a more dense packing structure than cellulose I.¹⁶⁸ However, as examined by Wada et al.,¹⁶⁹ the hydrolysis of cellulose II (and especially its hydrate form) proceeds faster than the hydrolysis of cellulose I. Changes in polarity, crystallinity, and ultrastructure of cellulose I to cellulose II have been reported to be the factors responsible for cellulose II faster hydrolysis.¹⁶⁷

While some of the traditional pretreatments suffer from relatively low sugar yields, require severe reaction conditions (high temperature and/or high pressure), and result in the formation of fermentation inhibitory compounds, cellulose solvent-based pretreatments can be performed under relatively mild conditions (100–160 °C), resulting in an insignificant amount of inhibitors from degradation of cellulose and hemicelluloses.^170,171 Cellulose solvent-based fractionations are regarded as biomass-independent or feedstock agnostic pretreatments, which can break the recalcitrant structure of biomass by increasing the cellulose accessibility more than the traditional pretreatments.¹⁷² The recovery of non-fermentable co-products, e.g., pure and unaltered lignin, in these methods, adds revenue streams to the fermentation products.^173,174 The use of cellulose solvents over traditional solvent systems, which are typically (e.g., ethanol) volatile, for biomass pretreatment is promising for the future of lignocellulosic biorefineries.

This review paper has mainly focused on the most promising cellulose solvent-based pretreatment, i.e., concentrated phosphoric acid (CPA), N-methyl-morpholine-N-oxide (NMMO, or NMO), and ionic liquids (ILs). Although a few other reviews are available in the literature,^{172,175–186} this review is intended to be a comprehensive review, with the focus on recent research on cellulose solvent-based pretreatments to improve the reactivity of lignocelluloses for biogas, ethanol, and renewable chemical production. Furthermore, as the pretreatment is a preceding step to microbial conversion mediated by enzymes, the basic concepts and the limiting factors in the enzymatic hydrolysis of lignocelluloses are also briefly reviewed.

4 Hydrolysis of pretreated lignocellulosic substrates

The hydrolysis of lignocelluloses has long been done by dilute and concentrated acids, e.g., sulfuric acid.^187,188 The main drawback of acid hydrolysis is degradation of sugars and formation of byproducts that showed severe inhibition to fermentation microorganisms. High investments and maintenance cost, high utility and disposal costs, high energy consumption for acid recovery, and environmental impacts are among the major disadvantages of acid hydrolysis.¹⁸⁹ Hydrolysis of lignocellulosic materials by “enzymatic” processes has emerged as a prominent process for the production of monomeric sugars, e.g., for subsequent production of fuel ethanol.^190,191 Cellulases and hemicellulases are the two enzymes typically used for the depolymerization of lignocellulosic carbohydrates to fermentable sugars for second-generation biofuel production. Although a lot of efforts have been made to reduce the production costs, the enzymes are still expensive.^192,193

Cellulose can be hydrolyzed by three glycoside hydrolases: endo-1,4-β-D-glucanases (EG) (EC 3.2.1.4), which randomly hydrolyze internal β-1,4-glucosidic bonds in the cellulose microfibril; exo-1,4-β-D-glucanases or cellobiohydrolases I and II (CBH) (EC 3.2.1.91), which progressively convert cellulose into cellodextrins; and 1,4-β-D-glucosidases (EC 3.2.1.21), which hydrolyze cellobiose and cellodextrins to glucose.^{139,194–196} In a synergistic mixture, cellulases have higher combined activities than the sum of their individual activities.¹⁹⁷ Cellulases typically have two separate domains: a catalytic domain (CD) and a cellulose binding module (CBM), comprised of approximately 35 amino acids, linked by a flexible linker region.¹⁹⁸ Over the years, several kinetic models for lignocellulosic biomass hydrolysis by cellulase have been proposed and developed^199–201 to understand the mechanisms. For example, recently a comprehensive model was developed by Bansal et al.²⁰² that included the following steps: (i) adsorption of cellulases onto the substrate via the binding domain, (ii) direction of cellulases to a bond (located on the chain end or cleavable bond) susceptible to hydrolysis on the substrate surface, (iii) formation of the enzyme–substrate complex, (iv) hydrolysis of the β-glycosidic bond and simultaneous direction of the enzyme to the cellulose chain, (v) desorption of cellulases from the substrate or repetition of step iv or steps ii/iii if only the catalytic domain detaches from the chain, and (vi) hydrolysis of cellobiose to glucose by β-glucosidase (if available in the enzyme mixture). However, the exact mechanism of cellulose hydrolysis mediated by fungal cellulases is still unknown as the binding mechanism of the binding module to cellulose, catalytic action of cellulase, and stimulation of cellulose hydrolysis by CBMs are still not clearly understood.²⁰³

The catalytic domains in cellulase are connected to one or more CBMs by peptide linkers of varying length and structure.^139,204 CBMs, with high binding affinity, increase the interaction between cellulase and the cellulose surface and enhance enzyme penetration into the substrates.^139,205,206 Several synergistic proteins, e.g., plant expansins and expansin-like proteins such as swollenin,²⁰⁷ and auxiliary activity family 9 (formerly GH61) proteins,^208,209 are able to enhance the enzymatic hydrolysis of cellulose by cellulase in ways that are not yet clearly understood.²¹⁰

Hemicellulases refer to a diverse combination of enzymes that can synergistically hydrolyze hemicellulose from mixed sources and are divided into two major categories: depolymerases and debranching enzymes (accessory enzymes).^195,211 The former group is either endo-acting enzymes, that attack polymer chains internally, or exo-acting enzymes that act processively²¹² from the reducing or non-reducing terminals.²¹³ Depolymerases mainly include xylanases, mannanases, β-glucanases, and xyloglucanases, and debranching enzymes are α-glucuronidase, α-arabinofuranosidase, α-D-galactosidase, acetyl xylan esterase, and ferulic acid esterase.^211,214

Várnai et al.²¹⁵ reported that the synergistic action of xylanase and mannanase can improve the total hydrolysis of pretreated softwood. Synergism is defined as “the ratio of the rate or yield of product released by enzymes when used together to the sum of the rate or yield of these products when the enzymes are used separately in the same amounts as they were employed in the mixture”.²¹⁶ It depends on both the ratio of the enzymes involved and the characteristics of the enzymes and substrate.¹⁰² Synergism, as reviewed by Van Dyk and Pletschke,¹⁰² can be grouped into cellulase component interaction, as mentioned earlier, hemicellulase interaction, and combined enzymes on complex substrates.

For degradation of lignocelluloses, many aerobic bacteria and fungi, e.g., Acidothermus cellulolyticus, Trichoderma reesei, and Aspergillus niger, produce free enzymes. Nonetheless, some anaerobic bacteria from genera of Clostridium, Acetivibrio, Bacteriodes, and Ruminicoccus are capable of producing multi-enzyme extracellular protein complexes, called cellulosomes, which can degrade cellulose, hemicellulose, and pectin.^{102,139,198,205,217} The most important characteristic difference between cellulosomes and free enzyme is cohesion-containing scaffoldin(s) and dockerin-containing enzymes (hemicellulases, cellulases, and pectinases).^218,219 Besides, free non-cellulosomal enzymes usually contain a CBM that attach to the substrate. The structure and function of cellulosomes and their differences with free enzymes have been reviewed by Bayer et al.²²⁰

The enzymatic hydrolysis and fermentation can be conventionally performed by separate enzymatic hydrolysis and fermentation (SHF) or via an integrated process, i.e., simultaneous saccharification and fermentation (SSF), non-isothermal simultaneous saccharification and fermentation (NSSF), simultaneous saccharification, filtration, and fermentation (SSFF), or simultaneous saccharification and co-fermentation (SSCF)²²¹ (Fig. 6). Although a recent study reported a higher ethanol yield by SHF over SSF at very high solid concentration by using new preparations of a cellulolytic enzyme, Cellic® CTec2,²²² integrated approaches were developed to enhance the overall ethanol yield by reducing the inhibitory effect of sugar released during the hydrolysis process on enzymes.²²³ Another approach, called consolidated bioprocessing (CBP), can convert biomass to biofuel by using anaerobic bacteria capable of producing cellulosome enzymes with high activity and ferment the resulting sugars to, e.g., ethanol, in a single step.^192,203,219 Although CBP is a more effective process than the others, it is in the developing stage and further developments in metabolic and genetic engineering are required to meet the industrial requirements.

Fig. 6 Different strategies for hydrolysis and fermentation of lignocellulosic substrates (SHF: separate hydrolysis and fermentation; SSF: simultaneous saccharification and fermentation; CBP: consolidated bioprocessing; SHCF: separate hydrolysis and co-fermentation; and SSCF: simultaneous saccharification and co-fermentation).

Commercial enzymes are usually a cellulase mixture derived from fungi such as T. reesei supplemented with β-glucosidase and contain more than 80 proteins.^195,224 Novozymes is one of the companies that provide enzymes for process optimization and commercialization of cellulosic ethanol. In this regard, the company started a dedicated work in 2000, under a national renewable energy laboratory (NREL) subcontract funded by the United States Department of Energy (DOE), to reduce the cost of cellulases.²⁰³ In 2007, the company estimated 40–100 times higher cost for the hydrolytic cellulase enzyme than the cost of enzymes for starch hydrolysis to glucose on a per gallon ethanol basis.¹⁵⁷ The outcome of the work was Cellic CTec & Cellic HTec enzyme cocktails in March 2009 followed by an improved and cost-effective product, Cellic® CTec2, in February 2010, and the company reported a 35% lower enzyme price. The company then developed a new generation of enzyme, called Cellic® CTec3, with 1.5 times better performance than the previous best product in the market. Cellic® CTec3 has been shown to work across a variety of feedstocks with a consumption of approximately 50 kg of Cellic® CTec3 to produce 1 ton of ethanol (https://www.novozymes.com/). The cellulase assays usually measure the production of reducing sugars from high molecular weight cellulose,²²⁵ like Whatman 1 filter paper, as first developed by Ghose²²⁶ and later adopted and modified by NREL.²²⁷ The protein content of the enzymes is also of great interest, which is usually measured by the Bradford assay,²²⁸ Pierce BCA assay,²²⁹ and total crude protein by Kjeldahl nitrogen analysis.²³⁰

Equivalent glucose yield, proposed by the NREL, as % of theoretical yield (% cellulose or glucan digestibility) is usually calculated by using eqn (1):

where [Glucose] is the concentration of glucose (g L⁻¹), [Cellobiose] is the cellobiose concentration (g L⁻¹), [Biomass] is the biomass concentration on a dry basis at the beginning of the enzymatic hydrolysis (g L⁻¹), and f is the cellulose fraction in the biomass on a dry basis (g g⁻¹).²³¹

5 Obstacles in the enzymatic hydrolysis of lignocelluloses and the role of pretreatment

The enzymatic hydrolysis performance of lignocelluloses is affected by not only cellulolytic enzyme-related factors (discussed in Section 4) but also by the physical, chemical, and morphological characteristics of the lignocellulosic materials.^{151,232–234} Cellulose crystallinity, structure, degree of polymerization (DP), and accessibility, as well as hemicellulose and lignin contents are among the main structural and physicochemical features of cellulosic substrates that control the rate and extent of enzymatic hydrolysis.^{113,235–238} Among all the factors that control cellulose hydrolysis mediated by fungal enzymes and to an extent, by cellulolytic organisms such as Clostridium thermocellum and other microbes, cellulose accessibility to enzymes/microbes is believed to be the main factor affecting cellulose deconstruction.^{113,172,239,240} However, tracking only one factor governing biological conversion is practically impossible because an increase in cellulose accessibility in biomass is usually accompanied by hemicellulose and lignin removal and/or reduction in cellulose crystallinity.

5.1 Cellulose crystallinity and degree of polymerization (DP)

Cellulose microfibrils exist in different polymorphs, i.e., crystalline, paracrystalline (disordered), and amorphous structures. Amorphous cellulose is much easier to hydrolyze than crystalline cellulose.²⁴¹ One of the major obstacles for efficient hydrolysis of cellulose mediated by fungal enzymes is the cellulose crystalline structure since lignin- and hemicellulose-free substrates, e.g., cotton fibers, still show resistance to enzymatic degradation.²⁴² However, based on findings in the literature, the correlation between the cellulose crystallinity and enzymatic hydrolysis rate and yield is still debatable.^243–247Although cellulose accessibility and enzyme adsorption can be affected by cellulose crystallinity, lignin/hemicellulose contents and distribution, biomass porosity, and biomass particle size can also affect the accessibility.²⁴³ Besides, some reports have stated a constant crystallinity for cellulose during the course of hydrolysis;²⁴⁴ while others reported a decrease in cellulose crystallinity during hydrolysis.²⁴⁵

Reported by Hall et al.,²⁴⁶ at constant adsorbed enzyme concentration, crystallinity was found to be a more influencing factor for enzymatic hydrolysis rates than enzyme adsorption. Mittal et al.²⁴⁷ found a strong correlation between the initial rate of digestion (up to 24 hours) and the amorphous content for four cellulose samples with different degrees of polymerization and crystallinity indexes, which were subjected to aqueous sodium hydroxide and anhydrous liquid ammonia treatments. Besides, they reported a weak correlation of allomorph type with initial digestibility; however, a strong correlation with cellulose conversion was found at later hydrolysis times. Cui et al.²⁴⁸ prepared four types of cellulose allomorphs from α-cellulose and concluded that the amorphous content had a strong positive influence on cellulose digestibility. The allomorph digestibility was reported to be in the following order: cellulose III > cellulose II > cellulose I_α > cellulose I_β. In contrast, the crystalline polymorph of cellulose was reported to have a negligible influence on the conversion degree of non-dried and dried cellulose samples into glucose.²⁴⁹ Finally, cellulose crystallinity can affect the synergism among cellulase components and the cellulase processivity, which has a notable effect on the hydrolysis.²⁴¹

The crystallinity index measurements are highly dependent on the technique applied, i.e., Fourier transform infrared spectroscopy (FTIR), X-ray diffraction (XRD), Nuclear Magnetic Resonance (NMR) Spectroscopy, and Raman spectroscopy, and also the methods used for calculating the crystallinity index from the raw spectrographic data.^204,242 The cellulose crystallinity index (CrI) from XRD spectra has long been calculated by different calculation approaches.²⁴³ The most frequent and simple calculation technique is based on the peak height according to the empirical method of Segal et al.²⁵⁰ for native cellulose:

CrI (%) = [(I₀₀₂ − I_am)/I₀₀₂] × 100 (2)

where I₀₀₂ is the maximum intensity of the 002 lattice diffraction at 2θ = 22.4° and I_am is the diffraction intensity at 2θ = 18°. However, Segal's crystallinity method does not reflect the crystal sizes for a given polymorph, e.g., the two cellulose polymorphs, I_β and II, were calculated to have different CrIs despite having the same crystal sizes.²⁵¹

The degree of polymerization (DP) of cellulose is the number of glucose units in the cellulose molecule chain and varies between 6000 in the primary cell wall and up to 14 000 in the secondary cell wall.²⁵² The DP of cellulose is believed to contribute to the enzymatic hydrolysis of lignocelluloses since a long cellulose chain has more hydrogen bonds, while a shorter chain has more cellulose ends available to the exoglucanases.²⁵³

However, tracking changes in the DP of cellulose, especially for complex lignocelluloses, during the course of pretreatment cannot be easily assayed. A method developed by Zhang and Lynd²⁵⁴ is only applicable to pure cellulosic substrates. Besides, DP is not typically an independent factor influencing cellulose digestibility because altering DP is always accompanied by crystallinity changes.^148,255

5.2 Cellulose accessibility to cellulases

One of the primary barriers for cellulase enzymes in the hydrolysis of lignocellulose is their limited access to much of the cellulose confined in a highly packed structure.²⁵⁶ The presence of lignin significantly decreases the swelling/accessibility of cellulose resulting in low sugar yields at commercially viable low enzyme loading.¹²⁰ Arantes and Saddler²⁵⁷ found that the required protein loading to achieve efficient hydrolysis of lignocellulosic substrates, regardless of their source, structure, and type of pretreatment, had a strong linear dependency on the cellulose accessibility for each substrate. Biomass porosity is considered as the lignocellulosic interior surface area and exterior surface area and is largely determined by the particle size.²⁵⁸ The accessible pore sizes required for anaerobes and cellulase and hemicellulase enzymes were reported to be at least 0.2–20 μm and 40–60 nm in width, respectively, to allow sufficient penetration.²⁵³ Wiman et al.²⁵⁹ correlated the higher rate of enzymatic hydrolysis, in spite of the negative effect of lignin accumulation on the particle surface, with the increase in specific surface area. Rollin et al.²⁴⁰ also showed that increasing cellulose accessibility is more important than removing lignin in the enzymatic hydrolysis of pretreated substrates, while removing lignin increases the accessibility of hemicelluloses which in turn affects cellulose accessibility.²⁶⁰ Similar to cellulose crystallinity, a strong relationship was observed between the accessible cellulose surface and the degree of synergistic action of cellulase components, which is crucial to enhance the hydrolysis efficiency.²⁶¹

5.2.1 Cellulase adsorption. The rate-limiting step in enzymatic saccharification is the amount of protein adsorbed on the substrate during enzymatic hydrolysis. The rate of saccharification increases with increasing enzyme concentration up to a plateau, typically corresponding to the maximum capacity of the substrate to adsorb enzymes.^262,263 The decrease in hydrolysis rates with reaction is believed to be mainly due to reduced enzyme adsorption and accessibility to the substrate.²⁶⁴ The adsorption parameters (maximum adsorption capacity [σ] and equilibrium constant [K_d]) are usually determined by fitting the cellulase adsorption data to the Langmuir equation by non-linear regression:where [CE] is the amount of adsorbed enzyme in mg g⁻¹ substrate, [E_f] is the free enzyme concentration in mg mL⁻¹, σ is the maximum adsorption capacity in mg mg⁻¹ substrate, and K_d is the equilibrium constant in mg enzyme per mL.²⁶⁰

The concentration of free enzymes is measured either directly by analyzing the adsorbed protein on the substrate or calculated as the difference between the total amount of protein initially added and the amount left in aqueous solution at any time.^{262,265–267} The enzymes were reported to adsorb quickly in the initial stage and remain attached throughout hydrolysis.²⁶⁸ For instance, the equilibrium time for cellulase on pretreated sugarcane bagasse was approximately 120 min and was even shorter for Avicel (10 min), while β-glucosidase (from A. niger) was not significantly adsorbed.²⁶⁹

5.3 Hemicellulose content

Hemicelluloses, a physical barrier around cellulose, can retard the enzymatic hydrolysis by precluding the access of enzymes to cellulose (Section 5.2) and inhibiting the endoglucanase and cellobiohydrolase activity.^270,271 The presence of xylan is believed to limit the cellulose hydrolysability, as evident by slow digestion of delignified substrates compared to pure cellulose.^272,273 Although it is commonly found in the pulp and paper industry that xylan and other hemicelluloses adsorb on cellulose and enhance pulp strength, Kumar et al.^274,640 recently showed that hemicellulose adsorption and their strong association with cellulose during pretreatments can retard cellulose digestion significantly; however, supplementation of xylanase to cellulase was shown to relieve the inhibition. In another report, Wang et al.²⁷⁵ also reported that the re-adsorption of dissolved xylan, produced during the pretreatment, on cellulose can inhibit the cellulose hydrolysis by cellulases. The supplementation of cellulases by xylanase was suggested to hydrolyze the xylan adsorbed on cellulose and potentially improve the hydrolysis efficiency of lignocelluloses. As discussed earlier (Section 4), the supplementation of xylanase has also been reported to synergistically improve the performance of cellulases in the hydrolysis of lignocelluloses.^{216,276–278} Nonetheless, hemicellulase supplementation to cellulase not only enhances cellulose accessibility to cellulase by simultaneously removing structural/non-structural hemicelluloses but also depolymerizes shorter hemicellulose oligomers in the solution that have been shown to be strongly inhibitory to cellulases by Kumar and Wyman and others.^279–284 On the other hand, a negative effect of xylose accumulation on cellulase cocktails was also observed.²⁸⁵ Partial removal of hemicelluloses by concentrated NaOH was reported to be more effective than complete removal for poplar, and a maximum enzymatic hydrolysis of 94.6% was achieved.²⁸⁵ More information on the inhibitory effects of sugars and oligomers on the enzymatic hydrolysis is provided in Section 5.5.

5.4 Lignin content

In general, lignin plays a negative role in the biochemical processes for producing lignocellulosic biofuels.^286–289 Nonetheless, Nakagame et al.²⁹⁰ concluded that an increase in the carboxylic content of lignin resulted in a decrease in non-productive binding of cellulase and consequently an increase in hydrolysis yield. A slight enhancement in enzymatic hydrolysis was also reported by Wang et al.²⁹¹ by adding Kraft lignin to the enzymatic hydrolysates. Lai et al.²⁹² reported contrasting results for the effect of ethanol organosolv lignin on enzymatic hydrolysis. They found that the addition of 8 g L⁻¹ hardwood organosolv lignin significantly improved the enzymatic yield of organosolv pretreated sweetgum and loblolly pine, while addition of softwood organosolv lignin was shown to decrease the yields.

Lignin can retard the enzymatic hydrolysis of lignocelluloses via three mechanisms: (1) enzymes can be adsorbed on lignin through hydrophobic interactions, electrostatic interactions, and/or hydrogen-bonding interactions, (2) lignin in lignocellulosic materials acts as a surface barrier to block the accessible surface of carbohydrates through physical blockage on the surface and chemical blockage through the lignin–carbohydrate complex, and (3) enzyme deactivation by soluble lignin.^293,294

Öhgren et al.²⁷⁸ evaluated the effects of partial delignification of corn stover by acid-catalyzed or autocatalysis pretreatment to increase the enzymatic hydrolysis yield. Due to the delignification, a slight increase in glucose yield and a decrease in xylose yield due to hemicellulose loss were observed. Várnai et al.²⁷² concluded that the limitation in the enzymatic hydrolysis of spruce was mainly due to the presence of lignin, since the removal of lignin with chlorite delignification doubled the hydrolysis yield with near theoretical yield within 2 days. Nlewem et al.²⁹⁵ performed alkali, dilute acid, and hot water pretreatments on switchgrass and compared its enzymatic hydrolysability. Although it was not only due to delignification, the alkali pretreatment generally produced glucose at higher concentrations than the others, since it caused higher reduction in lignin content and lots of pores were formed by the pretreatment. In another study, fungal delignification of wet milled rice straw by Trichoderma viride in the presence of a surfactant for 30 days resulted in 74% of lignin removal and 56% of enzymatic saccharification.²⁹⁶

5.4.1 Adsorption of cellulases on lignin. Non-productive cellulase adsorption onto lignin is believed to be associated with the inhibitory effect of lignin on the enzymatic hydrolysis of lignocellulosic feedstocks.^297–299 Both raw softwood lignin and isolated lignin from steam pretreated softwood were reported to adsorb major commercial T. reesei cellulases (Celluclast) and inhibit the hydrolysis of Avicel.³⁰⁰ The composition and functional groups of lignin, e.g., syringyl/guaiacyl lignin ratio, carboxylic acid, aliphatic hydroxyl, and phenolic hydroxyl, were reported to affect the enzyme adsorption.³⁰¹ Lignin adsorbed the enzymes in the following order: cellobiohydrolases (CBHs) and xylanase > endoglucanase (EG) > β-glucosidase (BG). In contrast, Ko et al.³⁰² reported that β-glucosidase from T. reesei had the strongest adsorption onto lignin and only 2–18% of the initial β-glucosidase activity remained in the supernatant, while 50–60% of cellobiohydrolase and endoglucanase activities were recovered after incubation with lignin. However, they stated that β-glucosidase from A. niger exhibits less adsorption than that from T. reesei. Rahikainen et al.³⁰³ prepared lignin films from steam explosion pretreated and untreated spruce and wheat straw and compared their capacity to adsorb cellulases. The pretreated biomass film showed higher capacity to adsorb the major cellulase Cel7A of T. reesei than the untreated biomass. Yu et al.²⁹³ also showed that the lignin obtained from pretreated woods resulted in two to six times more cellulase adsorption than untreated woods. The degree of lignin condensation after pretreatment, which significantly increased especially for softwoods, has a critical impact on cellulase adsorption and enzymatic hydrolysis.²⁹³

5.4.2 Lignin-derived phenolic compounds. Lignin-derived phenolic compounds, e.g., vanillin, syringaldehyde, trans-cinnamic acid, and hydroxybenzoic acid, generally produced during pretreatment inhibit cellulase (endo- and exo-cellulases and β-glucosidase) as well as fermentative microorganisms.^304–307 The enzymes get deactivated and precipitated with vanillin, where a 10 mg mL⁻¹ vanillin concentration was reported to decrease cellulose conversion from 53% to 26%.³⁰⁴ The structure of the phenolic compounds, e.g., the presence of hydroxyl, carbonyl, and methoxy groups, can affect the inhibition. Li et al.³⁰⁸ reported that the aldehyde and phenolic hydroxyl groups of vanillin have inhibitory effects on cellulase. However, β-glucosidases from T. reesei and A. niger are less susceptible to inhibition and correspondingly require approximately 10 and 100 times higher concentrations of phenols for the same levels of inhibition as cellulase components.³⁰⁵ Oliva-Taravilla et al.³⁰⁹ showed that the addition of laccases was able to remove the phenolic compounds from steam-pretreated lignocellulosic materials; however, application of laccases reduced glucose yield during hydrolysis. They concluded that the proportion of lignin besides the composition of phenols is a key factor in the cellulase inhibition when the enzymatic hydrolysis is combined with laccase detoxification.

5.5 Formation of inhibitory byproducts

Besides hemicellulose and lignin-derived compounds, some inhibitory byproducts produced during pretreatment, e.g., furan aldehydes, weak acids, and hydrolysis-derived substances like soluble mono/oligomeric sugars (Section 5.3), hamper the performance of cellulases and fermentable organisms.^307,310,311 Furan aldehydes, i.e., furfural and 5-hydroxymethylfurfural (HMF), are formed by dehydration of pentose and hexose sugars, respectively^312,313 (Fig. 7). By the release of acetic acid during pretreatment, mainly by hydrolysis of the acetyl group, or by re-hydrolysis, furan aldehydes can be converted to weak acids such as levulinic acid and formic acid (eqn (4)).^314–316

Fig. 7 Formation of major inhibitory by-products from main carbohydrates present in lignocelluloses (modified from Reginatto et al.³¹⁷).

Formation of inhibitory byproducts during pretreatment is strongly dependent on feedstock and pretreatment type applied. For example, agricultural residues and hardwoods with higher amounts of acetylated xylan generate a higher concentration of acetic acid during pretreatment. Most of the pretreatments under severe conditions, such as a long reaction time and high temperature, result in the formation of inhibitory by-products. In acid-catalyzed thermochemical pretreatment processes, dehydration of pentose sugars and uronic acid results in inhibitory byproducts (Fig. 7). In addition, the splitting of the lignin's β-O-4 ether and other acid labile linkages forms phenolic and non-phenolic aromatic inhibitory compounds. However, the formation of carboxylic acids by the peeling-off reaction takes place under alkaline conditions.^307,311

Jing et al.³¹⁸ compared the inhibitory effect of the major lignocellulose degradation products on Spezyme®CP cellulase and found it be in the following order: lignin derivatives > furan derivatives > organic acids > ethanol. Arora et al.³¹⁹ reported a severe inhibition by formic acid (5 or 10 mg mL⁻¹) on the enzymatic hydrolysis of cellulose powder as well as dilute acid-pretreated poplar.

Xiao et al.³²⁰ quantitatively calculated the inhibitory effect of sugars on cellulase and β-glucosidase during the enzymatic hydrolysis of softwood substrates and showed a dramatic increase in the inhibition of both enzymes by increasing the glucose concentration. They also reported the significant inhibitory effect of mannose, xylose, and galactose during the hydrolysis on cellulase activity but not on β-glucosidase activity. Xylooligomers (XOs), especially at high concentrations, were reported to have a greater inhibitory effect than xylan and xylose, resulting in the decrease of the initial hydrolysis rate and final glucose yield of Avicel.^280,284 Addition of xylanase and β-xylosidase was recommended to reduce xylooligomer and xylan inhibition of enzymatic hydrolysis of pretreated corn stover.³²¹ In a recent study, Kumar and Wyman²⁷⁹ revealed that mannan polysaccharides and their enzymatically derived oligomers were more inhibitory to cellulase than XOs and cellobiose. They also showed that cellulase inhibition dramatically increased with mannan backbone substitution with galactose. However, the amount of mannan re-adsorption on cellulose after pretreatment was reported to be higher than that of glucomannan and galactomannan at the same concentrations.³²² In a recent study, the Cellic® CTec3 enzyme mixture was reported to be more resistant than Celluclast 1.5L cellulase to the inhibitory compounds produced during steam pretreatment of poplar and lodgepole pine.³²³ Furthermore, monomeric sugars were shown to have greater inhibitory effects than phenolics, depending upon their types, and oligomeric sugars.

It is notable that the discussed byproducts also have inhibitory effects on the bioconversion routes leading to biofuel and renewable chemical production. For example, the concentrations of furfural and HMF in the range of 0.5–1 g L⁻¹ and formic and acetic acids at more than 4 g L⁻¹ were reported to be toxic in batch lactic acid fermentation by Rhizopus oryzae.³²⁴ For a recombinant S. cerevisiae strain, initial furfural concentrations below 5 g L⁻¹ were reported to have a negligible effect on ethanolic fermentation in a xylose and glucose containing medium, while xylose consumption rates were affected at initial furfural concentrations of 10–15 g L⁻¹.³²⁵

6 Concentrated phosphoric acid pretreatment

Phosphoric acid (85%) was first recognized as a swelling agent to produce reactive cellulose from air dried cellulose by Walseth³²⁶ in the 1950s. Since then, phosphoric acid swollen cellulose (PASC) has been the subject of vast studies as a cellulose substrate for cellulase activity assays and preparation of microcrystalline cellulose.^327–329 Bellamy and Holub³³⁰ patented a process using CPA (80–85%) for decrystallization of cellulose to improve its hydrolysis. The process included the formation of a gel by mixing cotton and wood pulp with CPA at room temperature followed by acid removal from the cellulosic substrate by water washing. Zhang et al.,³³¹ however, observed cellulose dissolution behavior when the phosphoric acid concentration reached greater than 80.5%, the critical concentration value for the dissolution of Avicel. During the first stage of the dissolution, an esterification reaction between the hydroxyl group of cellulose and phosphoric acid occurs and cellulose phosphate (cellulose-O-PO₃H₂) is formed. In the second stage, a competitive hydrogen-bond reaction between the cellulose hydroxyl groups and the solvent molecules or hydrogen ions happens and regenerated cellulose and phosphoric acid without major substitution are recovered.^332,333 Meanwhile, cellulose hydrolysis remains minimum since the reaction temperature is kept low enough (30–70 °C) to retard the depolymerization and side reactions.³³⁴

Conte et al.³³⁵ by applying high- and low-field NMR confirmed that direct bonding between phosphoric acid and cellulose is formed. Zhang et al.³³³ particularly investigated the structural changes of microcrystalline cellulose (MCC) dissolution in 83% phosphoric acid (at temperatures of 30–70 °C) with X-ray diffraction, solid-state cross-polarization magic angle spinning ¹³C-NMR, and X-ray photoelectron spectroscopy (XPS). The XRD pattern demonstrated a decrease in χ_c (crystallinity index) with increasing temperature (from 30 to 70 °C) or time (from 2 to 6 h). χ_c was calculated according to the following equation,

χ_c=F_c/(F_a + F_c) × 100% (5)

where F_c and F_a are the area of the crystal (peak of cellulose I at 2θ = 22.8°) and non-crystal regions (peak at 2θ = 19.8°), respectively.

Besides, the crystallinity characteristic peaks for both cellulose I and II diminished or greatly decreased after cellulose regeneration from concentrated phosphoric acid (CPA). In the spectra of CP/MAS and ¹³C solid-state NMR, distinct peaks of C₄ verified a transition from crystalline to amorphous cellulose after CPA treatment.³³³ The XRD patterns of MCC treated with 85% CPA at 323 K also demonstrated that more cellulose I was converted to cellulose II by increasing the reaction time from 0 to 6 h.³³⁶ Jia et al.³³⁷ chemically modified MCC with phosphoric acid in order to enhance its processing for applications in gelling materials and emulsion stabilizers. Regenerated cellulose at some angles corresponding to crystallographic planes of cellulose II exhibited less crystallinity compared to intact MCC. Besides, the crystallinity index was reduced by 48% after regeneration.

The dissolution was also capable of fractionating lignocellulose components under the modest reaction conditions, and the cellulose can be regenerated by an organic solvent, e.g., ethanol and acetone, or water.^240,334,641 Addition of an antisolvent, e.g., acetone, causes the dissolved cellulose and hemicellulose to precipitate and partial dissolution of lignin in acetone also takes place. Besides, hemicellulose oligomers are fractionated from cellulose due to higher solubility in water and poor solubility in the water/acetone mixture.³³⁴ The regenerated amorphous cellulose, precipitated from the dissolved cellulose, demonstrated extremely high reactivity for enzymatic digestibility, suggesting the dissolution technique as a new approach for the pretreatment of lignocelluloses.³³¹ Recently, a new cellulose solvent- and organic solvent-based lignocellulosic fractionation (COSLIF) method using concentrated phosphoric acid, as a cellulose solvent, and an organic solvent (e.g., acetone or ethanol) for the solute precipitation, under modest reaction conditions was developed.³³⁴ This novel pretreatment was able to effectively disrupt the lignocellulosic structure of switchgrass,³⁴⁰ bamboo,³³⁸ common reed,³³⁹ and miscanthus and hybrid poplar.³⁴⁰Table 2 summarizes the results of glucan digestibility improvement after COSLIF, as well as the applied conditions, for different lignocelluloses. As can be seen in Table 2, COSLIF pretreatment is performed under mild conditions, e.g., temperatures of ca. 50–60 °C, atmospheric pressure, and short pretreatment time (∼1 h), using acetone, ethanol, and water as anti-solvents. Compared with other most commonly used pretreatment methods, such as dilute acid, alkali, and hydrothermal, the sugar yields for CPA pretreatment for a variety of hardwoods and agricultural residues are very high. For instance, over 90% glucan digestibility was achieved after 72 h hydrolysis even at low enzyme loadings. Moreover, some studies reported ethanol yield enhancement by the CPA pretreatment (Table 3).

Table 2 Glucan digestibility of various substrates prepared by cellulose solvent- (phosphoric acid) and organic solvent-based lignocellulosic fractionation (COSLIF) pretreatment

Substrate	COSLIF condition	Enzymatic hydrolysis	Glucan digestibility	Ref.
Sesbania grandiflora (L.) Pers.	H3PO4 (85%), 50 °C for 45 min, 95% (v/v) ethanol as an organic solvent	1 FPU cellulase from Sigma	86% glucose in 72 h	346
Achyranthes aspera and Sida acuta weed	70%, 75%, and 80% phosphoric acid (1.0 g/8.0 mL), and 60 °C for 1 h, and acetone as an organic solvent	30 FPU g^−1 dry biomass Celluclast 1.5 L and 60 U g^−1 dry biomass β-glucosidase	Up to 86.2% and 82.2% glucan conversion yields, respectively	347
Alamo switchgrass (Panicum virgatum)	85% H3PO4, 60 °C, 1 atm, for 45 min, 95% (v/v) ethanol as an organic solvent	Novozymes 50013, 15 and 3 FPU g^−1 glucan, supplemented with 10 IU g^−1 β-glucosidase	90% and 85%, respectively, in 72 h	240
Moso bamboo	85% H3PO4, 50 °C, 1 atm, for 60 min, 95% (v/v) ethanol as an organic solvent	Novozymes 50013 and β -glucosidase (Novozymes 50010), 1, 2, 5, and 15 FPU of cellulase per g glucan supplemented with 10 β-glucosidase IU g^−1	88.2%, 89.8%, 93.3%, and 94.9%, respectively, in 72 h	338
Common reed (Phragmites australis)	85% H3PO4, 50 °C, 1 atm, and 60 min, 95% (v/v) ethanol as an organic solvent	15, 10, and 5 FPU and 30 units of β-glucosidase per gram of glucan, (Novozymes 50013 and Novozyme 50010)	94%, 93%, and 90%, respectively, 24 h	339
Miscanthus and poplar	85% H3PO4, 50 °C, 1 atm, and 60 min, 95% (v/v) ethanol as an organic solvent	5 FPU of cellulase and 10 units of β-glucosidase per gram of glucan, (Novozymes 50013 and Novozyme 50010)	∼93% in 72 h	340
Microcrystalline cellulose	83% H3PO4 and ice-cold distilled water as an anti-solvent	15 FPU g^−1 cellulose and 60 IU β-glucosidase per g cellulose	100% cellulose conversion after 3 h	331
Avicel and α-cellulose	81.7% phosphoric acid at room temperature for a half-hour, and acetone as an organic solvent	15 FPU per g glucan of Genencor Spezyme®CP cellulase and 60 IU g^−1 glucan of Novozymes 188 β-glucosidase	100% conversion within 3 h	334
Corn stover and switchgrass	84% phosphoric acid at 50 °C for 45 min, and acetone as an organic solvent	15 FPU g^−1 glucan of Genencor Spezyme®CP cellulase and 60 IU g^−1 glucan of Novozymes 188 β-glucosidase	∼96–97% in 24 h	334,341
Hybrid poplar and douglas fir	85% phosphoric acid at 50 °C for 60 min, and acetone as an organic solvent	15 FPU g^−1 glucan of Genencor Spezyme®CP cellulase and 60 IU g^−1 glucan of Novozymes 188 β-glucosidase	∼97% and ∼75% in 24 h for hybrid poplar and douglas fir, respectively	334
Oriented strand board, chipboard, plywood, and wallpaper	85.9% phosphoric acid at 50 °C for 30 min, and acetone as an organic solvent	20 FPU cellulase (Sigma, C2730) and 50 IU β-glucosidase (Sigma, G0395) per gram of substrate	87.0–93.5% in 96 h	348
Hybrid poplar (P. tormentosa Carr.)	85% phosphoric acid and room temperature until complete dissolution, and water as solvent	50 FPU 1 : 1 blend of Celluclast 1.5 L and Novozyme 188/g substrate	92%, 72 h	349
Industrial hemp stalks	85.9% H3PO4 at 50 °C for 1 h, and organic solvent, acetone	15 FPU cellulase (Spezyme CP), and 60 IU β-glucosidase per gram of glucan	95.9%, 24 h	350
Bermudagrass, reed, and rapeseed	85% phosphoric acid at 50 °C for 60 min, and acetone as an organic solvent	25 FPU of Celluclast® 1.5 L per gram of cellulose	97.5–99.4% (24 h)	351
Eastern gamagrass (Trypsacum dactyloides) and switchgrass	The pretreatment method reported by Zhang et al.^334 and modified by Ge et al.^352	100 μL of Novozymes 188, or 600 μL of cellulase and 200 μL of Novozymes 1800 for high solid-loading	80.5–99.8% and 73.5–87.1%, for eastern gamagrass and switchgrass, respectively, 36 h	353
Giant reed, elephantgrass, and sugarcane clone	85% phosphoric acid at 50 °C for 60 min, and organic solvent, acetone	300 μL of cellulase (Sigma C2730) and 100 μL of Novozymes 188 (Sigma C6105)	Glucose yields from biomass: 0.306, 0.309, 0.331, 0.317, and 0.290 g g^−1 for giant miscanthus, giant reed, giant miscanthus (Q42641), elephantgrass, and sugarcane, respectively	352
Corn stover and Avicel	85%(w/w) phosphoric acid, 2% (w/v) solid loading, described by Zhang et al.^331	5 FPU g^−1 of glucan (Novozymes 50013) and 10 units of β-glucosidase (Novozymes 50010) per gram of glucan	∼90% (72 h) for corn stover and 100% (6 h) for Avicel	354

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Table 3 Ethanol production from pretreated lignocelluloses prepared by COSLIF pretreatment^a

Substrate	CPA pretreatment condition	Method and microorganism	Ethanol yield	Ref.
a SHF: separate enzymatic hydrolysis and fermentation; SSF: simultaneous saccharification and fermentation; NSSF: non-isothermal simultaneous saccharification and fermentation.
Dedicated energy crops and crop residues	The same as reported by Zhang et al.^331	SHF, three self-flocculating Saccharomyces cerevisiae strains: SPSC01, ATCC24859, ATCC4126	0.375 to 0.396 g g^−1 (SPSC01), 0.380 to 0.394 g g^−1 (ATCC24859), and 0.384 to 0.405 g ethanol per g (ATCC4126) glucose	352
Oil palm empty fruit bunches (OPEFB)	The same as reported by Zhang et al.^334	SSF, S. cerevisiae	89.4% of theoretical maximum ethanol yield	355
Aspen wood (Populus tremula)	Phosphoric acid (85%), 12.5% solid loading, 50 °C, 90 rpm, 30 min, and acetone as an organic solvent	NSSF, Mucor hiemalis	72.4% of theoretical maximum ethanol yield	356
Rice straw, elmwood, and pinewood	The same as of Rollin et al.^240	SHF, Mucor indicus	Over 78–92% ethanol yield based on glucose consumed	342
Tripsacum dactyloides	The same as of Zhang et al.^334	SHF, a self-flocculating yeast strain SPSC01	Up to 0.496 g ethanol per g glucose	353

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COSLIF was observed to follow a different mechanism than alkali or acid pretreatment with respect to changes in lignocellulosic components. Zhu et al.³⁴¹ compared glucan, hemicellulose, and lignin contents of the COSLIF and dilute acid (DA) pretreated corn stover. They reported that COSLIF removed more lignin compared to DA pretreatment. Siripong et al.³⁴⁷ reported removal of all xylan and ca. half of the acid-insoluble lignin from two wood species as a result of CPA (80%) pretreatment. Similarly, Rollin et al.²⁴⁰ reported a 67% and 34% hemicellulose and lignin removal, respectively, from switchgrass by CPA pretreatment. They reported a greater increase in cellulose susceptibility to hydrolysis in COSLIF pretreatment than soaking in aqueous ammonia (SAA, 10% w/w ammonia, 140 °C, 20 : 1 liquid/solid ratio, 14 h) for Alamo switchgrass (Fig. 8). However, the cellulose content remained almost constant after both the pretreatments. Another action of CPA pretreatment is to hydrolyze hemicellulose acetyl groups to acetic acid.^334,339 The remaining hemicellulose can be enzymatically depolymerized and used as a co-substrate for fermentation.^342,343

Fig. 8 Conceptual image of alteration in the lignocellulose structure as a result of cellulose solvent- (concentrated phosphoric acid) and organic solvent-based lignocellulose fractionation (COSLIF) and soaking in aqueous ammonia (SAA) pretreatments (taken from ref. 235 with permission).

There are few studies in the literature that showed biogas production improvement by CPA pretreatment. A study showed 40% improvement in the methane yield obtained after CPA pretreatment (85.7% CPA at 50 °C for 30 min) compared with that of the untreated oil palm empty fruit bunches.³⁴⁴ Conversely, CPA pretreatment did not improve the methane yield for berry and poplar woods.³⁴⁵ This is presumably due to the repelling interaction of anaerobic bacteria and biomass surface after CPA pretreatment. In addition, the pores generated following CPA pretreatment may not be large enough for anaerobic bacteria to penetrate into the biomass structure.

6.1 Criteria for efficient phosphoric acid pretreatment

A narrow range of phosphoric acid concentrations is required for the cellulose phase transition from swelling to dissolution to occur. Only phosphoric acid above its critical concentration is able to disrupt the lignocellulose recalcitrant structure.³³¹ The critical phosphoric acid concentration is in the range of 77–83 wt%, depending on substrate type and its moisture content. Fig. 9 shows the SEM images of pretreated cotton fibers with a range of o-phosphoric acid concentrations.³⁵⁷ As shown in this figure, amorphogenesis begins to develop at the surface of the cotton fibers when the acid concentration was increased to near its critical values of cellulose dissolution. At 74% acid concentration, splitting, roughening, fibrillation, and peeling/delamination were observed, indicating that amorphogenesis started at the surface of the cotton fibers. Amorphogenesis is developed by increasing acid concentration, and destruction and diminishing of fiber structure were observed at 76% and 78% acids, respectively.³⁵¹ Moxley et al.³⁵⁰ also found that a minimum phosphoric acid concentration of 81% is required to obtain a very rapid hydrolysis rate and high digestibility of hemp stalks. Jia et al.³³⁷ discovered minimum 77.8 wt% CPA for significant solubilization of MCC powder. Zhang et al.³³¹ showed that 77 wt% of CPA caused only cellulose swelling while ice-cold phosphoric acid (≥83%) completely dissolved MCC.

Fig. 9 SEM images of cotton linter pretreated with different concentrations of O-phosphoric acid (0–78% w/w). Pretreatment conditions were: ice-cold temperature, one hour with occasional mixing, and water as an antisolvent (taken from ref. 357).

The dissolution of (ligno)celluloses in CPA also depends on the reaction temperature and time. Cellulose dissolution by CPA usually occurs at modest reaction temperatures.³³⁴ Moxley et al.³⁵⁰ investigated the effect of 84.0% H₃PO₄ pretreatment at different reaction times (from 30 to 120 min) at 50 °C and pretreatment temperatures (from 40 to 60 °C) for 60 min on the enzymatic glucan digestibility of hemp stalks. Higher reaction temperatures and time resulted in faster fiber dissolution; however, significant hydrolysis of cellulose and hemicellulose or sugar degradation occurred under these conditions. In terms of enhanced MCC processing ability by CPA, however, a decreasing trend in solubility was observed by increasing the temperature from 5 to 75 °C.³³⁷ Sathitsuksanoh et al.³³⁹ optimized the COSLIF conditions for enhanced saccharification at decreased cellulase loadings by response surface methodology (RSM). The optimal conditions were 85% (w/v) CPA, 50 °C, and 60 min, regardless of the biomass moisture contents from 5–15% (w/w). These modest reaction conditions can minimize sugar degradation, inhibitor formation, and capital investment of industrial plants.

Addition of volatile organic solvents is used for regenerating amorphous cellulose and hemicellulose, dissolving the organic solvent lignin soluble fraction, and for the recycling and re-concentration of PA.³⁵⁸ Recently, replacement of acetone by ethanol was presented and widely used in a modified version of the COSLIF. This modification is advantageous because ethanol is more chemically stable than acetone for solid/liquid separation and less corrosive. Besides, a very high yield of acetone recovery (e.g., >99.99%) is required for having an economically viable COSLIF implementation, whereas lower ethanol recycling/recovery after pretreatment (e.g., 98–99%) is acceptable,^338–340 since the remaining ethanol can be separated in an ethanol distillation process. Moreover, a 40% decrease in organic solvent consumption was achieved in the replacement of acetone by ethanol.³³⁹

6.2 Why is phosphoric acid so effective in enhancing enzymatic hydrolysis?

Many studies reported that substrate accessibility to cellulase determines the susceptibility of lignocellulosic substrates to enzymatic hydrolysis.^{257,260,261,297,359–361} Cellulose accessibility to cellulase (CAC) is usually quantified by cellulase adsorption Langmuir kinetics, as discussed in Section 5.2.^261,362 Recently, a quantitative assay for CAC, based on the adsorption of a nonhydrolytic fusion protein containing CBM and GFP, was developed by Hong et al.³⁶³ and applied for pretreated substrate characterization. CAC (m2 g⁻¹ of cellulose) was calculated by multiplying a constant to maximum cellulase adsorption capacity obtained from the Langmuir equation (eqn (3)).³⁶³ For pretreated lignocellulosic biomass, total substrate accessibility to cellulase (TSAC) represented the cellulase adsorption capacity for the whole biomass and was calculated by adding CAC and non-cellulose accessibility to cellulase (NCAC).³⁴¹ TSAC was equal to CAC for protein thioredoxin-GFP-CBM (TGC) adsorption to biomass. Similarly, CAC and TSAC (m2 g⁻¹ biomass) can be calculated by TGC adsorption after BSA blocking of the lignin fraction. Therefore, NCAC (m2 g⁻¹ biomass) can be calculated as the difference between TSAC and CAC.³⁴¹ TSAC, CAC, and NCAC (m2 g⁻¹ biomass) measurements of intact lignocelluloses were reported to be approximately 1 m2 g⁻¹ biomass.^{240,339–341} Untreated Alamo switchgrass (Panicum virgatum), for example, had 1.27, 0.49, and 0.77 m² g⁻¹-biomass TSAC, CAC, and NCAC, respectively.³⁶⁴ SAA slightly improved all the accessibilities, while COSLIF resulted in a considerable increase of 9.6 and 8.0 for TSAC and CAC (m2 g⁻¹ biomass), respectively.²⁴⁰ Similarly, an almost 2-fold increase in the accessibilities was observed for COSLIF-treated corn stover compared to DA pretreatment.³⁴¹ TSAC (m2 g⁻¹-biomass) of miscanthus and poplar also increased after COSLIF pretreatment but more radically from 0.18 to 20.7 and 0.23 to 18.2, respectively.³⁴⁰ Common reed followed the same pattern as miscanthus and poplar after the pretreatment.³³⁹

Breaking or even restructuring the highly ordered intra- and inter-molecular hydrogen-bond network of crystalline cellulose is believed to enhance its depolymerization rate.^106,337,365 The evidence of breaking hydrogen-bonding networks in cellulose fibers of switchgrass after COSLIF was confirmed by CP/MAS ¹³C-NMR and FTIR.³⁶⁵ Other analytical techniques, e.g., microscopy and X-ray diffraction, also showed the disruption of the hydrogen-bond network of cellulose for MCC regenerated from CPA.³³⁷ John et al.³⁶⁶ investigated the structures of native and regenerated celluloses by X-ray methods and proposed the same lattice plane location of the inter-molecular hydrogen bonds between adjacent cellulose molecules. The empty space between adjacent cellulose chains could be occupied by the hydrogen ion from phosphoric acid; therefore, inter-molecular hydrogen bond formation is destroyed during the regeneration process.³⁴⁹

Recently, computer simulations have been employed to study the biomass recalcitrance at the molecular level that otherwise cannot be analyzed with available experimental techniques.^367,368 Molecular dynamics simulation (MDS) and quantum chemical calculations, e.g., density functional theory (DFT) methods, are the tools of molecular simulation. These techniques have been used for the simulation of lignin biosynthesis and degradation,^369,370 cellulose insolubility,³⁷¹ and recently for the simulation of the effect of ammonia pretreatment on cellulose I_β.¹⁰⁶ Although models of secondary plant cell walls incorporating cellulose, xylan, water, and lignin by MD simulations were generated,³⁷² the molecular simulation studies on lignocelluloses are scarce. This is due to the complex lignocellulose biomass structure and also the intricate relationship between enzymes, chemicals, and biomass. Molecular simulation for lignocelluloses is still in its early stage of development and needs further investigation to fill the gap of advancing analytical methods in pretreatment.

6.3 Summary and future perspectives of phosphoric acid pretreatment

Taking all into consideration, COSLIF was successful with a number of agricultural residues and hardwoods^342,373 and demonstrated the advantages of high glucan digestibility even at low cellulase loadings, high hydrolysis rates, modest reaction conditions, higher revenues from co-products (acetic acid, lignin, and hemicelluloses), and less inhibitor formation. Besides, the remaining CPA on treated biomass did not show inhibitory effects for enzymatic hydrolysis or fermentation processes. However, it is still in its early stage of development and its commercialization is a far promising priority that needs pervasive consideration. Although there are only a few studies in the literature, CPA pretreatment does not seem to be very effective in improving biogas production from lignocelluloses. Substantial reduction in the use of chemicals (both CPA and organic solvent) is required in order to have an economically competitive process. Improvement of ethanol production process economy was suggested by the production of two major value-added byproducts, i.e., unaltered and purified lignins by the COSLIF and byproducts from fermentation.³⁴² Although CPA pretreatment seems to be very promising given the high end-product and by-products yields, a detailed techno-economic analysis of CPA pretreatment is required in order to study the feasibility of this pretreatment for a large-scale operation.

7 N-Methylmorpholine-N-oxide (NMMO) pretreatment

N-Methylmorpholine-N-oxide (NMMO or NMO) is categorized as a family of cyclic, aliphatic, tertiary amine oxides.^374,375 Tertiary amine oxide systems were first patented by Graenacher and Sallmann³⁷⁶ in 1939 to dissolve cellulose for enhanced chemical processing. However, Johnson,³⁷⁷ for the first time in 1969, introduced a cyclic mono(N-methylamine-N-oxide) compound to interact with inter-molecular hydrogen bonding networks and dissolve cellulose, wool, silk, hair, and feather, which are insoluble in commonly used solvents. Since the late 1970s, research on the dissolution of cellulose in NMMO was initiated when McCorsley and Varga³⁷⁸ produced a highly concentrated, yet economical, cellulose solution by dissolving cellulose in a NMMO–water system. At that time, research on NMMO–cellulose tertiary systems was mainly focused on producing regenerated cellulose fibers that have applications in textiles and nonwovens, the lyocell process, strengthening paper films, and paper coatings.^{374,377,379–381} However, this technology has been recently introduced as a pretreatment method of lignocelluloses, e.g., for the improvement of either second-generation bioethanol^{115,382–388} or biogas production.^388–397

Having a strong N–O dipole, which acts as either an ionic or donative and single bond, NMMO is capable of disrupting the hydrogen-bond networks of cellulose and building new hydrogen bonds between the polymer and the solvent^375,379,398 (Fig. 10). Cellulose dissolution in NMMO leads to a tertiary phase of the cellulose–NMMO–water system.^379,399 Hydration with 1–1.2 water molecules per NMMO (water content 13.3–17 wt%) significantly improves its interaction with a solute and boosts its solvation ability, while increasing the water content to 19–24% and 25–30% results in heterogeneous swelling by forming balloons and ballooning, respectively.⁴⁰⁰ Higher water contents (above 35%) make fibers swell homogeneously and precipitate, because in the tertiary system water is further preferred to form hydrogen bonds with NMMO than cellulose.^375,379,400 Ballooning and swelling modes of cellulose dissolution are more efficient for biogas production, while for ethanol production pretreatment with 85% NMMO leads to better lignocellulose bioprocessing.³⁸⁴Fig. 11 shows a microscopic image of wood fiber swollen by ballooning in NMMO solution, where the three zones of the membrane of the balloons, the inside of the balloons, and the nonswollen crystalline regions are easily identified.

Fig. 10 The mechanism of cellulose dissolution in NMMO, adapted from Wang et al.¹⁶³ with permission.

Fig. 11 Wood fiber swollen by ballooning in a 78 wt% NMMO solution in water, taken from ref. 404.

Lignocelluloses are directly dissolved in the solvent at moderate temperatures (90–130 °C) under atmospheric pressure for 20 min to 5 h with negligible derivatization. Cellulose is subsequently regenerated by adding water as an anti-solvent to the slurry. The regenerated cellulose (cellulose precipitated from the NMMO solution) is converted from cellulose I to cellulose II structure, which is much more reactive for cellulase adsorption and subsequent hydrolysis.^401,402 The solvent is washed away from the regenerated solids with distilled boiling water, and the excess water can be easily vaporized due to the low vapor pressure of NMMO, allowing approximately 99% of NMMO recycling.⁴⁰³

7.1 Effect of NMMO pretreatment on the superstructure of lignocelluloses

The enhancement in the digestibility of regenerated lignocellulosic biomass by NMMO pretreatment is mainly due to reduced cellulose crystallinity. The crystalline structure of regenerated lignocellulose from the NMMO solution as well as the untreated one are usually expressed by the Total Crystallinity Index (TCI) and Lateral Order Index (LOI) using FTIR.⁴⁰⁵ The FTIR spectra of lignocelluloses can also give some valuable information on the structure and the variation in characteristic bands by the pretreatment. Table 4 summarizes the characteristics of bands, their corresponding functional groups, and assignments to the major biomass constituents.

Table 4 Characteristics of bands from the FTIR spectra of lignocelluloses, from ref. 121

Wavenumber (cm^−1)	Functional group	Assignment
3175	–OH stretching (inter-molecular hydrogen-bonds)	Cellulose II
2900	C–H stretching	Cellulose
1740	C=O stretching (acetyl or carboxylic acid)	Hemicellulose and lignin
1510, 1610	C=C stretching (aromatic ring)	Lignin
1465	C–H3 (bending)	Lignin
1420, 1430	C–H2 (bending)	Cellulose
1375	C–H (bending)	Cellulose
1335	–OH (bending)	Cellulose
1315	C–H2 (wagging)	Cellulose
1158	C–O–C (stretching)	Cellulose

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Purwandari et al.³⁹⁶ reported that the TCI (the absorbance ratio A1427/A898 calculated from FTIR spectra) of oil palm empty fruit bunch (OPEFB) reduced by up to 78% following the pretreatment in 85% NMMO at 120 °C for 3 h. In addition, ballooning and swelling modes of NMMO result in a lower TCI at 120 °C than at 90 °C. This finding is in contrast to ballooning and swelling modes of NMMO pretreated cotton that result in lower crystallinity indexes at 90 °C than 120 °C.³⁸⁴ However, compared to the untreated cotton, crystallinity indexes decrease slightly for different modes of dissolution, ballooning, and swelling.¹²¹ Besides, the intra-molecular hydrogen-bonding OH stretching at about 3350 cm^−l (FTIR spectra) in pretreated cotton is broadened and shifted to a higher wave number,³⁸⁴ which is an indication of transforming cellulose I to cellulose II.^406,407 This finding is in accordance with another report on NMMO pretreatment of straw³⁸⁹ and also confirms that the pretreatment reduced the structural lignin content. NMMO pretreatment of bagasse at 130 °C for 1 h transformed the crystalline structure into an amorphous form, since the TCI and LOI decreased from 1.39 and 1.44 to 1.18 and 1.10, respectively.³⁸⁵ The LOI, a criterion for the estimation of the amorphous to crystalline portion of the structure, considerably decreased from 2.68 to 0.88 when the straw fraction of manure was pretreated for 5 h at 120 °C using 85% NMMO and decreased more with increase in pretreatment time to 15 h.³⁸⁹ Moreover, the LOI and TCI of rice straw pretreated with 85 wt% NMMO for 5 h at 120 °C decreased from 0.46 to 0.40 and 1.69 to 1.62, respectively.³⁸⁷ Likewise, Khodaverdi et al.⁴⁰⁸ reported that NMMO (85%) treatment of cotton linter at 120 °C for 2 h resulted in a TCI and LOI decrease from 7.1 and 2.7 for untreated cotton to 3.3 and 1.1, respectively.

The FTIR analysis also indicated that lignin and acetyl groups from the hemicellulose backbone were partially removed by the pretreatment, while the cellulose content increased. Liu et al.³⁶² qualitatively studied the abundance and distribution of lignin and cellulose in NMMO-pretreated pine flour using the FTIR technique. Diminishing of the absorbance peaks at 1270 cm⁻¹ and 1596 cm⁻¹, corresponding to lignin,⁴⁰⁹ indicated a reduction in lignin content on the surface of pine flour after NMMO pretreatment.³⁶² Furthermore, crystallinity measurement of the biomass by X-ray diffraction confirmed a linear correlation (R² = 0.91) between cellulose crystallinity and initial hydrolysis rates of the pine flour samples. Virtanen and Maunu⁴¹⁰ investigated the dissolution process of softwood pulp fibers in NMMO at 110 °C for 15, 30, and 90 min by employing different NMR spectroscopic methods: solid state cross polarization magic angle spinning (CP-MAS), ¹³C and ¹⁵N spectroscopies, and ¹H high resolution MAS NMR spectroscopy. The cellulose crystallinity of the NMMO pretreatment sample for 90 min decreased by 15%, and the C₄ signal appeared different from the untreated pulp, while it remained almost constant for the first 30 min of treatment with a broadening C₄ signal.

7.2 Changes in composition and microstructure during NMMO pretreatment

In general, carbohydrate contents of lignocelluloses do not undergo significant changes and high solid recoveries are achieved after NMMO pretreatments.^{115,394,396,397,411} This is an advantage of NMMO pretreatment over conventional pretreatment methods, since carbohydrate loss is a major problem in most chemical, physicochemical, and biological pretreatments.^136,187 However, a longer pretreatment time and/or temperature lead to partial removal of acid-insoluble lignin and xylan (or mannan in softwoods) and enrichment of the glucan constituent.^{382,383,386,387,389,393,411} Furthermore, structural studies confirmed the liberation of acetic acid from the acetyl groups of biomass during NMMO pretreatment, especially at longer pretreatment times and higher temperatures.¹¹⁵ The ash content was also reported to decrease from 5.4% up to 1.3% as a result of NMMO pretreatment of OPEFB,³⁹⁶ while no considerable change was reported for rice straw.³⁸⁷

7.2.1 Cellulose accessibility to cellulases. The porosity or specific surface area (SSA) of exposed cellulose is considered as another key feature of pretreated lignocellulosic substrates that influence the hydrolysis of cellulose by cellulases. In other words, cellulose accessibility is directly associated with the rates and extents of enzymatic deconstruction of lignocelluloses.⁴¹¹ Simons' Staining (SS) is a potentially useful semi-quantitative technique for specific surface area measurement of lignocellulosic substrates,⁴¹³ which was first introduced in 1950 to evaluate mechanical damage of pulp fibers during beating.⁴¹⁴ The SS method is based on dying substrates with direct blue 1 (DB) and then direct orange 15 (DO) to quantify smaller and larger pore sizes, respectively.⁴¹³ It has the advantages of measurement of interior and exterior surface area at even the wet state and being relatively fast and simple over other accessible surface area measurement techniques.⁴¹⁵ The total adsorbed dye amount, which represents the number of overall pores, considerably increased up to 1.5- and 2.2-fold for barley straw and forest residues, respectively, after NMMO pretreatment at 90 °C for 3–30 h.³⁹⁴ Moreover, the more the pretreatment time, the more the overall dye adsorbed. This finding was also confirmed by Teghammar et al.⁴¹⁶ for rice and triticale straw. Over 74% and 86% increase in total dye adsorption was observed for rice and triticale straw, respectively, after 15 h NMMO pretreatment at 130 °C. The biomass displays the same pattern in dye adsorption as in enzymes adsorption,⁴¹⁶ which is directly related to the enzyme accessibility of the substrate.²⁶³ An increase in enzyme adsorption by 100, 140, and 290% for triticale straw and 11, 50, and 240% for rice straw was observed after 1, 3, and 15 h of NMMO pretreatment, respectively.⁴¹⁶

Cellulose accessibility for NMMO-treated substrates was then evaluated by comparing the maximum adsorption capacity (by Langmuir adsorption isotherm) of pretreated samples and enzyme lignin (EnzL),³⁶² prepared by complete hydrolysis of carbohydrates in the pretreated biomass with excessive cellulase loadings.²⁶¹ The maximum adsorption capacity of cellulase onto pine flour samples as well as cellulose accessibility considerably increased with increasing NMMO pretreatment time from 30 to 120 min at 120 °C. Moreover, a nearly good linear correlation between cellulose accessibility and overall glucan conversion rate was also reported for pine flour.³⁶²

The other rapid specific surface area assessment technique is the water retention value (WRV) or water swelling capacity, which has been used to quantify the swelling potential of paper pulps.⁴¹⁷ WRV is the ability of water adsorption or the swelling capacity of the substrate and reflects the accessibility of the substrate to subsequent hydrolysis by enzymes.⁴¹² Besides, since substrate swelling and water adsorption occur mainly in the amorphous regions, the WRV can be used as a criterion to assess changes in the crystalline structure after pretreatment.⁴¹⁸ The water swelling capacity of birch hardwood after pretreatment with 85% NMMO at 130 °C for 3 h substantially increased by 46.6–119.9% depending on the applied drying method.³⁸³ The WRV of triticale straw also slightly increased by 10%, 10%, and 20% at NMMO pretreatment times of 1, 3, and 15 hours, respectively, and a smaller increase of 10% for rice straw was realized.⁴¹⁶ However, significant reduction in the WRV of cellulose was reported by NMMO pretreatment in dissolution mode at either 90 or 120 °C.³⁸⁴ This behavior was observed less at lower concentrations of NMMO (than 85%), but it still had lower WRV values compared to the untreated one.³⁸⁴

7.3 Advantages and disadvantages of NMMO pretreatment

NMMO is able to dissolve up to 15 wt% of cellulose⁴¹⁹ with no/less chemical modification under relatively mild conditions (low/moderate temperatures and atmospheric pressures). High bioprocess efficiency, high solvent recovery, and formation of low carbohydrate degradation and inhibitory products are also among the favorable characteristics of NMMO pretreatment. Tables 5 and 6 summarize an overview of treatment conditions along with improvements in saccharification/fermentation and biogas production from different lignocelluloses after NMMO pretreatment. These tables show that NMMO pretreatment is conducted under relatively mild conditions, i.e., temperature 90–130 °C for a few hours, using ∼85% NMMO. As can be seen in these tables, NMMO pretreatment causes significant improvement in ethanol, biogas, and enzymatic hydrolysis yields for different types of lignocelluloses including hardwoods, softwoods, agricultural residues, and other cellulosic substrates. By applying NMMO pretreatment, ethanol can be produced by S. cerevisiae, M. indicus, and Z. mobilis via different strategies, e.g., SSF, SHF, and NSSF (Table 5). The pretreatment resulted in up to 100% conversion of cellulose in enzymatic hydrolysis and 93.3% ethanol yields of theoretical maximum for rice straw (Table 5). An improvement of about 100% in the methane yield was also reported after NMMO pretreatment of cotton linter.³⁹² At pilot scale, maximum hydrolysis sugar yields of 195 and 175 mg sugar per g wood for spruce and birch wood chips, respectively, in NSSF with Mucor indicus were also achieved.³⁸⁶

Table 5 Improvement in glucan conversion/ethanol production yield from different lignocelluloses pretreated with NMMO

Substrate	NMMO condition	Method and fermentation microorganism	Glucan conversion and/or ethanol yield	Ref.
a Non-isothermal simultaneous saccharification and fermentation.
Spruce and oak	90, 110, and 130 °C, 1–3 h	NSSF^a, Saccharomyces cerevisiae	Up to 85.4% and 89% improvement in ethanol yield for spruce and oak, respectively	115
Rice straw	85 wt% NMMO, 120 °C, 1, 3, and 5 h, 5% loading	SSF, S. cerevisiae	Hydrolysis yield of glucan 96%, 93.3% of theoretical maximum ethanol yield	387
Cotton linter	90 and 120 °C, 0.5–15 h using 85%, 79%, and 73% NMO	SSF, S. cerevisiae	Improvement of up to 100% yield in enzymatic hydrolysis and 83.75% ethanol yield	384
Spruce and birch	85% NMMO, 130 °C, 1–5 h	Bench-scale and airlift cultivations, Mucor indicus	Maximum ethanol yields of 195 and 175 mg g^−1 wood for spruce and birch, respectively	386
Birch	85% NMMO, 130 °C, 3 h	SHF, S. cerevisiae,	Maximum 76.8% ethanol of theoretical yield, 9-fold increase in ethanol yield compared to untreated	383
Wheat straw	85% NMMO 120 °C for 1–5 h	Anaerobic cultivations, M. indicus	Up to 92.1% of theoretical maximum ethanol yield	382
Sugarcane bagasse	NMMO monohydrate, 130 °C, 1 h	SSF, Zymomonas mobilis	Approximately 0.15 g ethanol per g bagasse (86% of the theoretical maximum ethanol yield)	385

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Table 6 Improvement in biogas production from different lignocelluloses pretreated with NMMO

Substrate	Pretreatment condition	(Improvement in) methane yield	Ref.
a Volatile solid.
Oil palm empty fruit bunch (OPEFB)	90 and 120 °C, 1, 3, and 5 h, 85%, 79%, and 73% NMMO	Methane yield up to 0.408 Nm^3 kg^−1-VS, improvement by 167% compared to untreated	396
	73, 79, and 85% NNMO, 90 and 120 °C, 1, 3, and 7 h	Maximum 0.408 Nm^3 CH4 per kg-VS	395
Softwood spruce, rice straw, and triticale straw	130 °C, 1–15 h, 85% NMMO	Up to 245, 157, and 203 Nml CH4 per g raw material, respectively, 400–1200% improvement compared to the raw materials	397
Forest residues	120 °C, 3, 7, and 15 h, 75% and 85% NMMO	Up to 0.17 Nm^3 kg^−1-VS^a methane yield (83% of theoretical maximum yield)	393
Straw fraction of cattle and horse manure	5 h and 15 h, 120 °C, 85% NMMO	Maximum methane yield increase by 53% and 51% for cattle and horse manure, respectively, after 15 h pretreatment	389
Barley straw and forest residues	85% NMMO, 3–30 h, 90 °C	0.23 and 0.15 Nm^3 CH4 per kg-VS from barley straw and forest residues, respectively; corresponding to 88% and 83% of the theoretical maximum yields	394
Blended-fiber waste textiles	85% w/w NMMO, 120 °C, 2 h	Up to 62.18% of theoretical maximum methane yield (after 6 days)	391
Forest residues	NMMO concentrations of 75% and 85%, 120 and 90 °C, 3 and 15 h	Maximum 141% increase in methane production (75% NMMO at 120 °C for 15 h)	388
Jeans textiles	85% NMMO, 120 °C, 3 h	Two-stage semi-continuous process, 400 mL methane per g-VS per day	390
Cotton linter	85% NMMO, 5% w/w solid loading, 120 °C, 3 h	Approximately 100% methane yield (% of maximum theoretical) for 5 g L^−1 cellulose concentration after 30 days	392

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The solvent is recycled by treating the solution with ion-exchange resins to remove contaminants and subsequently dewatering the solvent.⁴²⁰ Due to the low vapor pressure of NMMO, excess water can be easily vaporized from the recycled solvent leaving the monohydrate form of NMMO.⁴⁰³ However, the water evaporation unit demands high-energy input which has considerable negative effects on the economy of the whole process.^421,422 Besides, in order to have an economically feasible process of bioethanol and biogas production by NMMO pretreatment of lignocelluloses, more than 99 percent of NMMO recovery is required.^421,422 Some side reactions and/or NMMO ring cleavage can occur in cellulose–NMMO solutions, especially at elevated process temperatures,^423,424 which hamper efficient solvent recovery. A study showed that a smaller amount of reducing sugars was liberated from NMMO-pretreated sugarcane bagasse at 130 °C rather than 100 °C, possibly due to NMMO or cellulose degradation.³⁸⁵ In some studies, recycled NMMO showed the same performance in hydrolysis improvement and biogas production of pretreated sugarcane bagasse and barley straw, respectively,^385,394 as compared with fresh NMMO. However, in contrast, forest residues with high lignin and bark content resulted in 55% reduction in methane yield after pretreatment with recycled NMMO in comparison with those pretreated with the fresh NMMO.³⁹⁴

The remaining NMMO in the regenerated solids may prove to have inhibitory effects on fermenting organisms and/or hydrolytic enzymes. NMMO concentrations of 5 and 100 g L⁻¹ have been shown to reduce the enzymatic hydrolysis yields by 12% and 76%, respectively, after 12 h of hydrolysis for cotton linter.³⁸⁴ Although NMMO decreased the glucose uptake rate by S. cerevisiae a little, it had negligible impact on the final ethanol yield even at a concentration of 100 g L⁻¹.^115,384 However, the ethanol yield and productivity decreased at concentrations above 2% NMMO for M. indicus, while the total production of metabolites was not significantly changed.³⁸⁶ This was because some glucose shunted from the ethanol to the glycerol pathway as the glycerol yield and production increased in proportion to NMMO concentration. Recently, He et al.⁴²⁵ introduced a NMMO-tolerant cellulase-producing strain from a newly isolated Galactomyces sp. CCZU11-1. The results showed that up to 25% (w/v) NMMO had no significant effect on the saccharification of NMMO-pretreated sugarcane bagasse prepared at 130 °C for 1 h or fermentation by S. cerevisiae. On the other hand, NMMO remaining in the pretreated substrate at concentrations higher than 0.002% was reported to considerably decrease the methane yield.³⁹³

Once the solvent was washed away from the substrate, NMMO leaving the process ends up in the wastewater stream. Nevertheless, it is not of great concern, since NMMO is an environmentally friendly solvent.^426,427

Techno-economic analysis of NMMO pretreatment of spruce for bioethanol and biogas⁴²¹ and forest residues for biogas production elucidated high process energy efficiency.⁴²² In the case of bioethanol production, a biogas plant in parallel to valorize pentoses can improve the process economy.⁴²¹ This is because most ethanol-producing organisms cannot assimilate pentoses efficiently.³⁴³ When forest residues were co-digested with two-thirds of organic fraction of municipal solid waste in order to avoid nitrogen deficiency, the process of biogas production was evaluated to be financially feasible at 15% internal rate of return or higher for a minimum plant capacity of 50 000 tons per year.⁴²² Generally, large amounts of water need to be vaporized in order to efficiently recover NMMO, and this is among the barriers for its commercialization.⁴²¹

8 Ionic liquid pretreatment

8.1 Ionic liquids: historical evolution and general properties

Ionic liquids (ILs) are usually defined as large organic salts, composed entirely of an organic cation and an organic or inorganic anion, which exist in liquid form at or below 100 °C.⁴²⁸ The field of ILs was first discovered in 1914 by Walden,⁴²⁹ who synthesized and characterized ethyl-ammonium nitrate ([EtNH₃][NO₃]) by neutralizing ethylamine with concentrated nitric acid. Organic based chloroaluminates ILs were first developed by Hurley et al.⁴³⁰ in 1951. A new class of ILs with melting point lower than ambient temperature based on 1-alkyl-3-methylimidazolium cation, called room-temperature ionic liquids (RTILs) and considered as the first generation ILs, has emerged since 1982 after the study by Wilkes et al.⁴³¹ The replacement of the moisture-sensitive anion in the first generation ILs by the tetrafluoroborate ion ([BF₄]⁻) and other anions resulted in more water-stable ILs in 1992, known as second-generation ILs.⁴³² Third generation ILs, known as “task-specific” ionic liquids (TSIL), which covalently incorporate either anions or cations or both as functional groups, were introduced by Davis⁴³³ in 2004.

ILs have negligible vapor pressures, high viscosity, and reasonable thermal and chemical stability, compared with typical organic solvents.^434,435 These properties can be changed and controlled by selection of cations and anions developed for a special application. This is why ILs are usually defined by the term “designer solvents”.⁴³⁶ ILs, due to their unique properties, have received significant attention for vast applications in chemical and biochemical industries.^{428,435,437–441}

8.2 Solvation in ILs

A simulation and vibration spectroscopy study of water–IL mixtures suggested the concentration dependent solubility of water in ILs. At low concentrations, the dissolution mechanism of water is molecular dispersion, while water aggregation takes place at higher concentrations. Dissolution of benzene in [DMIM](1,3-dimethylimidazolium)[PF₆], however, makes an expansion in the IL structure, while the long-range charge ordering pattern in the IL still exists.⁴⁴² One of the promising solvation features of ILs is their ability to dissolve monosaccharides, which are barely soluble in common solvents, except water.^443,444 Like benzene, a simulation understanding of glucose dissolution in the ionic liquid [DMIM][Cl] has been established.^445,446 The nature of the solute–solvent interaction in the system is mainly the hydrogen bond with a high chloride content of the IL. Youngs et al.⁴⁴⁶ suggested that the dominant coordinate of glucose dissolution in excess IL is the formation of three hydrogen-bonds between the OH groups of glucose and three anions, and an OH⋯Cl⋯HO bridge between the last two OH groups and the fourth chloride. The RTILs that contain dicyanamide anions were also reported to dissolve significant amounts of glucose, sucrose, lactose, and cyclodextrin.⁴⁴⁷ Other monosaccharides, including arabinose, fructose, mannose, and xylose, seem to have partial to high solubility in different ILs.⁴⁴⁸ Surprisingly, not only monosaccharides but also oligosaccharides and even polysaccharides are soluble in ILs. α-Cyclodextrin and starch, for example, were shown to have 30% and 10% solubility, respectively, in [BMIM](1-butyl-3-methylimidazolium)[Cl].⁴⁴⁸ Unlike dissolving saccharides in classic solvents, e.g., DMF and DMSO, the derivatization of native carbohydrates in ILs is of great importance since it is a green process.⁴⁴³ However, the aim of most studies on the carbohydrate–IL interaction is to produce non-derivatized cellulose, which has demonstrated vast applications in fiber and composite fiber production,⁴⁴⁹ as monoliths and films,⁴⁴³ and more recently, lignocellulosic biomass pretreatment.¹⁷³

8.2.1 Dissolution of cellulose in ILs. The first attempt to dissolve cellulose in ILs dates back to 1934 when Graenacher⁴⁵⁰ first utilized heated N-ethylpyridinium chloride in the presence of N-containing bases. Although many studies consider Graenacher's patent as the pioneer in IL dissolution of cellulose, recently, Sun et al.⁴⁵¹ claimed that the dissolution was stipulated by the addition of nitrogen-containing bases and not by the IL alone. Besides, the co-solvents used were volatile and the IL itself had a relatively high melting point (T_m; 120 °C) over conventional ILs. More recently, Swatloski et al.⁴⁵² investigated cellulose dissolution in ILs based on 1-butyl-3-methylimidazolium cations by publishing a highly cited paper in 2002. They further analyzed cellulose and cellulose oligomers in 1-butyl-3-methyl-imidazolium chloride IL solution using high-resolution ¹³C NMR.⁴⁵³ The ¹³C NMR data indicated that β-(1→4)-linked glucose oligomers were disordered, with conformational behavior being parallel to the one observed in water.

The selection of cations and specially anions in ILs plays a crucial role in cellulose dissolution.¹⁰⁸ Since the cellulose–IL bond, in nature, is the hydrogen-bond,⁴⁴⁶ it seems that anions with more hydrogen-bond-acceptor capability, e.g., OAc⁻, HCOO⁻, (MeO)₂PO₂⁻, and Cl⁻, are the suitable candidates for the solubility, while ILs with low-basicity anions, such as dicyanamide-based ILs, are not that efficient in dissolving cellulose.¹⁰⁸ ILs containing ‘noncoordinating’ anions, including [BF₄]⁻ or [PF₆]⁻, on the other hand, display no cellulose solubility.⁴⁵² Unlike anions, cations in ILs play an unclear, but effective role, in the cellulose dissolution.¹⁰⁸Table 7 lists the structure of some well-known cations of ILs for cellulose dissolution. Li et al.⁴⁵⁴ performed a simulation study and concluded that ILs with unsaturated heterocyclic cations can dissolve cellulose, whereas ILs with saturated ring cations can hardly dissolve cellulose. The reason for that was reported to be related to the structure factor and dynamic effect of the cations. Zhang et al.⁴⁵⁵ synthesized and used 1-allyl-3-methylimidazolium chloride as a non-derivatizing solvent for molecular dissolution of cellulose at room temperature. Although intra- and inter-molecular hydrogen-bond disruption were mainly due to the formation of a chloride hydrogen-bond network, it was suggested that the small polarized cation, [AMIM]⁺, also helped the attack on oxygen atoms of cellulose hydroxyl in this case.^455,456 The ¹³C NMR spectrum of MCC dissolved in [AMIM][Cl] clearly resolved the six signals of carbon atoms of unmodified anhydroglucose similar to cellulose dissolved in sodium hydroxide solution or [BMIM][Cl].⁴⁵⁵ However, the dissolution mechanism was not dominated by hydrogen bond formation between cellulose and chloride. Thus, first, it is important to compare cellulose solubility in chloride alkali metal salts. Chloride in LiCl, for example, perfectly interacts with cellulose hydroxyl groups and dissolves cellulose in the presence of N,N-dimethylacetamide (DMAc).⁴⁵⁵ Nevertheless, other chloride salts, e.g., sodium, potassium, barium, and calcium chloride, are unable to dissolve cellulose.⁴⁵⁵ Although concentrated zinc chloride is able to dissolve cellulose, its solvation behavior is due to the formation of the zinc–cellulose complex.⁴⁵⁷ Second, a unique anion with solubility potential, when combining with all range of cations, was not found. Vitz et al.⁴⁵⁸ conducted a thorough study on cellulose dissolution in imidazolium-based ILs with particularly bromide and chloride anions. An odd-even effect for imidazolium chloride ILs with more cellulose solubility in even-numbered alkyl chains of cations compared to the odd-numbered was observed, whereas, this pattern was not generalized for imidazolium-based ILs containing bromide anions. 1-Ethyl-3-methylimidazolium diethyl phosphate, however, demonstrated the maximum solubility of cellulose among all the imidazolium-based ILs. Although the role of cation is not yet well-clarified, it was reported that the size and polarizability and attached functional groups of the cation, e.g., hydroxyl end-group, or basic oxygen atoms affected its solubility.¹⁰⁸

Table 7 Structure of cations of well-known ILs used in the dissolution of lignocellulosic feedstocks

Cation structure	Name
	R = CH3: 1,3-dimethylimidazolium
	R = C2H5: 1-ethyl-3-methylimidazolium
	R = C3H7: 1-propyl-3-methylimidazolium
	R = C4H9: 1-butyl-3-methylimidazolium
	R = C5H11: 1-pentyl-3-methylimidazolium
	R = C6H13: 1-hexyl-3-methylimidazolium
	R = C7H15: 1-octyl-3-methylimidazolium
	R = C8H17: 1-nonyl-3-methylimidazolium
	R = C9H19: 1-decyl-3-methylimidazolium
	1-Cyano-3-methylimidazolium
	1-Allyl-3-methylimidazolium
	1-Benzyl-3-methylimidazolium
	1-(3-Methoxybenzyl)-3-methylimidazolium
	1-(3,6-Dioxahexyl)-3-methylimidazolium
	1-Ethyl-3-(3,6-dioxaheptyl)imidazolium
	1-(3,6,9-Trioxanonyl)-3-methylimidazolium
	1-Ethyl-3-(3,6,9-trioxadecyl)-imidazolium
	1-Butyl-3-(3,6,9-trioxadecyl)-imidazolium
	1-Ethyl-3-(4,8,12-trioxatridecyl)-imidazolium
	3,3-Ethane-1,2-diylbis(1-methyl-1H-imidazole-3-ium)
	1-Butyl-3-methylpyridinium
	1-Butyl-1-methylpyrrolidinium
	1-Ehyl-3-(3,6,9,12,15,18,21-heptaoxadococyl)-imidazolium
	1-(3,6-Dioxaheptyl)-3-(3,6,9-trioxadecyl)-imidazolium
	N-Benzyl-N,N-dimethylammonium
	Tetrabutylphosphonium
	Trihexyltetradecylphosphonium
	R = CH3: 8-metyl-1,8-diazabicyclo[5.4.0]undec-7-enium
	R = C4H9: 8-butyl-1,8-diazabicyclo[5.4.0]undec-7-enium
	R = C8H17: 8-octyl-1,8-diazabicyclo[5.4.0]undec-7-enium
	N,N-Dimethylathanolammonium
	Choline bis[(trifluoromethane)sulfonyl]imide
	1-Butyl-3-methylpyridinium
	1,8-Diazabicyclo[5.4.0]undec-7-enium
	N,N,N-Triethyl-3,6,9-trioxadecylammonium

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Sets of TSILs, or so-called tailor-made ILs, were also designed and characterized for the dissolution and depolymerization of cellulose under mild conditions.⁴⁵⁹ Thermal heating especially by microwave or sonication, degree of polymerization of cellulose, and the IL viscosity are among the non-IL-intrinsic effective factors in cellulose dissolution.^{108,460–463}

8.2.2 Dissolution and regeneration of lignocellulosic biomass in ILs. Kilpeläinen et al.⁴⁶⁴ demonstrated the capability of imidazolium-based ILs in dissolving hardwoods and softwoods under mild conditions. Xie et al.⁴⁶⁵ also reported the preparation of wool keratin/cellulose blended materials by dissolution and regeneration using [BMIM][Cl]. Fort et al.⁴⁶⁶ processed and analyzed the dissolution of woods of different hardness in [BMIM][Cl]. They reported the partial dissolution of untreated wood and celluloses with purities, physical properties, and processing characteristics comparable to those of pure cellulose samples subjected to similar treatment, which can be easily recovered from the resulting solutions by the addition of a variety of precipitating solvents. Li et al.⁴⁶⁷ investigated the factors affecting the dissolution of three wood species and regeneration in [AMIM][Cl]. Wood density, pulverization intensity, and the nature of the regeneration anti-solvents were reported as the main factors affecting the overall process. Generally, the ILs' anion and cation (cf. Section 8.2.1 for cellulose), viscosity, solvation properties, melting point and thermal decomposition, biomass particle size and type and loading, temperature and time of treatment, and microwave heating and sonication are among the important factors governing the dissolution of lignocellulosic biomass in ILs, which were recently reviewed by Badgujar and Bhanage.⁴⁶⁸ Freire et al.⁴⁶⁹ determined a set of thermophysical properties, i.e., density, viscosity, and refractive index, and isobaric thermal expansivity and heat capacities, for eight imidazolium-based ILs, as the important intrinsic IL parameters in the lignocellulose dissolution, and also the impact of anion type was investigated. Among the studied ILs, [EMIM][CH₃CO₂] was reported as the best candidate for lignocellulose dissolution, since it has shown to have a low viscosity and density. As the solvent properties of ILs, Doherty et al.⁴⁷⁰ concluded, by comparison of Kamlet–Taft α, β, and π* solvent polarity parameters of three RTILs, (i.e., [EMIM][OAc], [BMIM][OAc], and [BMIM][MeSO₄]) that the β parameter is an excellent predictor of pretreatment efficacy. Regarding the properties of lignin, Li et al.⁴⁷¹ achieved rapid dissolution of bagasse and southern yellow pine in [EMIM][OAc] by using a dissolution temperature above the glass transition of lignin.
8.2.2.1 Role of solvent in the regeneration of cellulose from IL solutions. Hauru et al.⁴⁷² characterized the Kamlet–Taft (KT) values of [EMIM][OAc], [TMGH][EtCO₂], and [TMGH][OAc], and NMMO at several water contents and temperatures to investigate the role of the solvent in cellulose regeneration from the IL solution. The regeneration of cellulose was reported to start at threshold values of approximately β < 0.8 (β − α < 0.35). Shi et al.⁴⁷³ investigated the pretreatment of switchgrass with different [EMIM][OAc] and water concentrations (50–80%) at 160 °C and concluded a strong dependency of the chemical composition and crystallinity of the pretreated biomass as well as the corresponding lignin dissolution and depolymerization on the IL concentration. They found the hydrogen-bond basicity of the [EMIM][OAc]–water mixture to be a suitable indicator of predicting the cellulose dissolution, lignin depolymerization, and sugar yields. Besides, their molecular simulation indicated that water acts as a co- and anti-solvent in cellulose dissolution at below and above 50% [EMIM][OAc] concentration, respectively. The role of anti-solvent, e.g., ethanol, water, and acetone, in cellulose regeneration from a cellulose/[BMIM][OAc] mixture was studied by molecular simulation.⁴⁷⁴ Structural analysis based on radial distribution function revealed that among the three studied solvents, water was the most effective solvent at breaking the cellulose–[Ac]⁻ H-bonds, leading to the subsequent formation of cellulose–cellulose H-bonds, and was demonstrated to be the best solvent for cellulose regeneration. Another molecular dynamics study was conducted to investigate the interaction of [EMIM][OAc], a cellulose oligomer, and water as an antisolvent.⁴⁷⁵Fig. 12 shows the proposed intermediate formed during the regeneration of a cellulose oligomer from the IL solution by using water, based on the simulation.

Fig. 12 Intermediate structure of celluloses regenerated from an IL in the presence of water as an anti-solvent proposed by Liu et al.⁴⁷⁵ using MD simulation, picture adapted from Liu et al.⁴⁷⁵

8.2.2.2 Biomass loading. Some studies on IL pretreatment have focused on high biomass loading in the pretreatment, instead of typical approximate 5.0 wt%, since it is a crucial factor for process economy. Besides, a minimum amount of consumed IL and waste generation happen at high biomass loading.⁴⁷⁶ Cruz et al.⁴⁷⁷ investigated the effects of switchgrass loading on [EMIM][OAc] pretreatment in terms of viscosity, cellulose crystallinity, chemical composition, saccharification kinetics, and sugar yield. The IL pretreatment caused reduction in biomass recalcitrance for 3, 10, 20, 30, 40, and 50 wt% biomass loading and a “solid” like behavior was observed when the biomass loading increased. Moreover, the IL pretreatment caused the transformation of the cellulose crystalline structure from I to II for 3, 10, 20 and 30 wt% samples, while a mostly amorphous structure was found for 40 and 50 wt% samples. Likewise, Wu et al.⁴⁷⁸ reported the feasibility of [EMIM][OAc] pretreatment of corn stover at 125 °C for 1 h at 50 wt% biomass loadings in dramatically reducing the recalcitrance of the biomass. In another study, da Silva et al.⁴⁷⁹ used a twin-screw extruder with high shear force to pretreat sugarcane bagasse at high solid loadings in [AMIM][Cl]. They obtained the maximum glucan digestibility of 90% after 24 h of enzymatic saccharification of the pretreated substrate at a loading as high as 25 wt% at 140 °C for 8 min. The pretreatment decreased the crystallinity significantly and increased the specific surface area (SSA) by more than 100-fold. At a higher biomass loading of 50 wt%, still 76.4% glucose yield was obtained. Li et al.⁴⁸⁰ obtained 99.8% fermentable sugars from switchgrass by [EMIM][OAc] pretreatment at 15% (w/w) biomass loading during a 600- and 60-fold process scale-up for the pretreatment and enzymatic hydrolysis, respectively. Ninomiya et al.⁴⁸¹ investigated the cholinum IL pretreatment as a function of IL/biomass weight ratio of bamboo. They obtained a critical IL/biomass ratio of 3 g g⁻¹ to obtain a cellulose saccharification of 80%, in a solid-state pretreatment.

8.2.3 Dissolution of lignin in ILs. The mechanism of lignin dissolution and regeneration in [AMIM][Cl] has been investigated by density functional theory (DFT), atoms in molecules (AIM) theory, natural bond orbital (NBO) analysis, and Wiberg bond index (WBI) by Ji et al.⁴⁸² The theoretical results showed that lignin mainly reacted with [AMIM][Cl] via H bonds, and it can be precipitated by adding water, since the absolute value of the interaction energy of AmimCl–nH₂O (n = 1, 2, and 3) is greater than that of AmimCl–LigOH. Further analyses of the regenerated lignin by FTIR, TG, and SEM revealed that no chemical reaction occurred for lignin during the dissolution and regeneration process. Wang et al.⁴⁸³ investigated the lignin dissolution in dialkylimidazolium-based IL–water mixtures at 60 °C. They found the maximum lignin solubility at 70 wt% IL, which was consistent with the Hansen theory, in which the IL type is important in the solubility. Accordingly, 1-butyl-3-methylimidazolium and methanesulfonate showed the maximum solubility of lignin among the examined ILs with the same anions and cations, respectively. Diop et al.⁴⁸⁴ invented new ILs for the dissolution of lignin and concluded that the lignin solubility decreased with increasing the length of the grafted carbon chain.

The capability of ILs in dissolving lignin can be employed in the delignification of lignocelluloses. Fu et al.⁴⁸⁵ chose [EMIM][OAc] amongst six ILs as the best candidate for the selective extraction of lignin to improve the enzymatic hydrolysis of triticale and wheat straw at various temperatures (70–150 °C) and time intervals (0.5–24 h). Lee et al.⁴⁸⁶ also reported the enhancement in cellulose digestibility caused by partial delignification of wood flour by [EMIM][CH₃COO]. They reported the maximum digestibility of cellulose to be 95% for triticale straw pretreated at 150 °C for 90 min. Wen et al.⁴⁸⁷ used [EMIM][OAc] under varying IL pretreatment conditions (i.e., 110–170 °C and 1–16 h) to isolate poplar alkaline lignin. Chemical transformation monitoring of the isolated lignin via elemental analysis, 2D-HSQC spectra, quantitative ¹³C-NMR spectra, ³¹P NMR, and GPC analyses revealed a decrease of aliphatic OH, mainly as a result of cleavage of the β-O-4′ linkage happened at high temperatures, and an increase in phenolic hydroxyl groups in lignin, attributed to the dehydration reaction during the pretreatment. The same study confirmed the β-O-4′ linkage broken with the dehydration and demethoxylation reactions during kraft lignin dissolution.⁴⁸⁸ 2D NMR bond abundance data and size exclusion chromatography (SEC) results also revealed that lignin was depolymerized during [EMIM][OAc] pretreatment at 120 and 160 °C of wheat straw, miscanthus, and Loblolly pine,⁴⁸⁹ and lignin with different molecular mass was released in different stages of the pretreatment. Brandt et al.⁴⁹⁰ obtained the same result in lignin characteristics isolated from miscanthus after extraction with the protic ionic liquid 1-butylimidazolium hydrogen sulfate ([HC₄im][HSO₄]). Their ¹³C-NMR, ¹H–¹³C HSQC NMR, ³¹P-NMR, Py-GC-MS, GPC, and elemental analyses showed that the lignin-hemicellulose linkages break and more than 80% depolymerization of lignin through the cleavage of glycosidic, ester, and β-O-4 ether bonds occurs during the early stage of the pretreatment. As the pretreatment proceeded, repolymerization of lignin happened, which was evidenced by the increased lignin molecular weight determined by GPC, and the increased phenolic hydroxyl group content and C/H ratio in the lignin prepared at the later stage. In another study, Varanasi et al.⁴⁹¹ pretreated Panicum virgatum and Eucalyptus globulus with [EMIM][OAc] at different temperatures and studied compositional changes in lignin. Preferential breakdown of S-lignin in both eucalyptus and switchgrass at high pretreatment temperature (160 °C) and breakdown of G-lignin for eucalyptus and no preferential breakdown of either S- or G-lignin in switchgrass were observed at lower pretreatment temperatures (120 °C), which may be linked to its hydrogen-bond accepting capacities at these temperatures. Accordingly, they suggested the mechanism similarity of the IL pretreatment to alkali pretreatment at lower temperatures and to acid pretreatment at higher temperatures. S-G-H type lignin was obtained from bamboo by [AMIM][Cl] treatment, where partial degradation of lignin and hemicellulose was observed.⁴⁹²

Thermochemical analysis is also used for the characterization of lignin extracted with ILs. The depolymerization and breakdown of lignin pretreated with [EMIM][OAc] at 120 °C and 160 °C for 1, 3, 6, and 12 h in model biomass compounds and bioenergy feedstocks, by thermogravimetric analysis (TGA), and differential scanning calorimetry (DSC) was reported.⁴⁹³ Lignin dissolution in cholinium ILs resulted in a higher maximal decomposition temperature (T_m) and a higher glass transition temperature (T_g) of kraft lignin.⁴⁸⁸ Moghaddam et al.⁴⁹⁴ compared the physicochemical properties of lignin isolated from sugarcane bagasse pretreated with acidified aqueous ethylene glycol (EG) and ILs, and soda lignin from NaOH pretreatment of bagasse. Accordingly, depolymerization of thermally stable IL and EG lignins occurred at higher temperatures compared to soda lignin. Moreover, unlike soda lignin, IL and EG lignins contained less/no carbohydrates, with slightly lower hydrogen and higher oxygen contents.⁴⁹⁴

George et al.⁴⁹⁵ investigated the effects of imidazolium-based IL cation and anion combinations on the macromolecular structure of three lignins, i.e., organosolv, alkali, and alkali low sulphonate. The results showed a significant reduction in molecular mass and a remarkable structural change of the lignins, primarily influenced by the anion, with anion influence in the reduction in the order sulfates > lactate > acetate > chlorides > phosphates, meanwhile cleavage of different linkages within the lignins was caused by different anions. However, at least 40% of the original large-lignin molecules, from each of the lignins studied, were observed to remain intact. On the other hand, extraction of lignin, with relatively uniform molecular weight without significant structural changes, from bagasse using an ionic liquid mixture [EMIM][ABS] at atmospheric pressure and elevated temperatures (170–190 °C) with a maximum yield of 93% was reported by Tan et al.⁴⁹⁶ They also concluded that ILs with better phase separation properties would be desirable for higher lignin extraction.

8.3 Effects of IL pretreatment on the cellulose structure

A majority of studies on the characterization of IL-treated lignocelluloses have focused on the transition of the cellulose crystalline structure and the surface morphology of biomass (e.g., Fig. 13 for macroscopic morphological changes). Zhang et al.⁴⁹⁷ studied the changes in the cellulose crystalline structure of three different feedstocks, switchgrass, corn stover, and rice husk, pretreated with [BMIM][OAc] at temperatures of 50–130 °C for 6 h by XRD. Increasing the treatment time led to a drop in biomass CrI, which was due to the swelling of crystalline cellulose and transition of cellulose I to cellulose II. Cheng et al.^498,499 pretreated Avicel cellulose, switchgrass, pine, and eucalyptus with [EMIM][OAc] at 120 °C and 160 °C for 1, 3, 6, and 12 h, and investigated the structural transformation and crystalline structure of cellulose. Although for Avicel the transformation to cellulose II occurred under all processing conditions, higher temperatures and times were required for the same transformation process for the other feedstocks, and only an expanded cellulose I lattice was observed under the mild conditions applied. Comparable with these results, XRD analysis showed a decrease in CrI from 39.2% to ∼0.09% and 28.6% to ∼0.03% for switchgrass and agave bagasse, respectively, after [EMIM][OAc] pretreatment at 120 °C for 3 h.⁵⁰⁰ The regenerated cellulose from rice husk resulted from [EMIM][DEP] pretreatment at 100 °C for 10 h (1.5% (w/v) loading) showed the highest decrease in the crystallinity index from 46.0 to 32.0, amongst the different ILs used.⁵⁰¹

Fig. 13 The macroscopic effects of [EMIM][OAC] pretreatment on spruce softwood (picture taken from Shafiei et al.⁵⁰⁷)

The morphological characterization of the wood cell wall treated with 1-ethylpyridinium bromide ([EtPy][Br]) and [EMIM][Cl] was studied by Kanbayashi and Miyafuji.^502,503 The analyses of three hardwood by light microscopy and SEM revealed that treatment with [EMIM][Cl] at 120 °C for 72 h caused significant swelling of all the woods. However, depending on the wood species, various behavior and different morphological changes in pits have occurred mainly due to their chemical component and the microfibril angle.⁵⁰⁴ Similarly, treatment of Japanese cedar with [EtPy][Br] caused the cell wall swelling and elimination of warts, while it did not change pit membranes and the cellulose crystalline structure.⁵⁰³ Additionally, Raman microscopic analysis showed that chemical changes in the cell walls were different for different cell wall layers in that lignin in the compound middle lamella and the cell corner was resistant to interaction with [EtPy][Br]. Singh et al.⁵⁰⁴ used auto-fluorescent mapping to visualize cellulose and lignin in switchgrass stems for determining the mechanism of biomass dissolution during 1-n-ethyl-3-methylimidazolium acetate pretreatment. Swelling of the secondary cell wall followed by complete dissolution of biomass within 3 h at 120 °C, and subsequent lignin removal by adding an anti-solvent was observed. The surface roughness of switchgrass, pine, and eucalyptus samples pretreated with [EMIM][OAc] at 120 °C for 1, 3, 6, and 12 h showed that switchgrass possessed much rougher internal surfaces than eucalyptus and pine.⁴⁹⁹ Zhang et al.⁵⁰⁵ monitored the swelling and dissolution behavior of poplar during [EMIM][OAc] pretreatment by employing confocal Raman microscopy. They concluded that the dissolution of the biomass was divided into two parts: slow penetration of the IL, which determined the process reaction rate, and rapid dissolution of lignin and carbohydrates. Therefore, enhancement of the penetration capacity of the IL, which was suggested to depend upon the properties of the IL, was crucial for improving the pretreatment efficiency. Confocal Raman microscopy and confocal fluorescence microscopy were also used to analyze the changes in different cell types including tracheids, sclerenchyma cells, and parenchyma cells of corn stover during [EMIM][OAc] pretreatment.⁵⁰⁶ A direct correlation was then observed between changes in the morphologies and chemical composition and swelling occurred mainly in the secondary plant cell walls.

8.4 Acid-catalyzed hydrolysis in ionic liquids

Different acids, e.g., mineral acids, Brönsted acids,^508,509 solid protic-acid resin,⁵¹⁰ and even amino acids,⁵¹¹ have been functionalized or co-utilized to enhance the effect of the IL pretreatment on enzymatic hydrolysis. The research conducted on using acidic ionic liquid solutions for the pretreatment of lignocelluloses can be divided into three parts. The use of acids for direct depolymerization of polymeric carbohydrates in the presence of ILs,^512–514 the use of an acid in ILs as a boosting pretreatment agent which can enhance the effectiveness of pretreatment in enzymatic hydrolysis,^510,515,516 and using acid-functionalized ILs for either direct hydrolysis or enhanced enzymatic hydrolysis of lignocelluloses.^{508,517–521} Although the use of homogeneous acid catalysts has its drawbacks, acid catalysts are currently used in IL pretreatment. One of the main reasons is the economic viability of the IL used for pretreatment. For example, even though [BMIM][Cl] costs ca. 1/60^th of [EMIM][OAc], it is not a highly efficient solvent for pretreatment; however, the addition of an acid catalyst can boost the performance of cheaper ILs for pretreatment.⁵¹⁰ Another reason is that acid hydrolysis of carbohydrates in such systems occurs at lower temperatures than in the aqueous phase.⁵¹³ Besides, because of the presence of lignin in the solid phase, sugar-lignin fractionation is easily achieved in such systems compared with aqueous phase reactions.⁵¹³

Development in the acidic IL pretreatment was first focused on the direct conversion of lignocelluloses into monomeric sugars. Li et al.⁵¹² developed a method for direct hydrolysis of cellulose in [BMIM][Cl] at 100 °C under atmospheric pressure catalyzed by mineral acids. The maximum glucose and total reducing sugar (TRS) yield of 43% and 77%, respectively, was obtained at 0.11 sulfuric acid/cellulose mass ratio for 540 min reaction time. Likewise, a maximum TRS yield of 65% was obtained from corn stover pretreated with [AMIM][Cl] at 100 °C for 90 min in the presence of 2.0 mmol HCl per gram lignocellulosic substrate.⁵¹⁴ Pretreatment of three wood species including eucalyptus, pine, and spruce thermomechanical pulp was performed at 120 °C for 3 h in [AMIM][Cl] followed by dilute hydrochloric acid hydrolysis for 5 h.⁵¹³ This IL-based acid pretreatment resulted in near-complete conversion of the woods' cellulose and hemicellulose at acid concentrations of 1.4–1.5 mole of HCl per g wood. However, at higher acid concentrations, the presence of several degradation compounds, such as 5-hydroxymethylfurfural (HMF), furan-2-carboxylic acid, catechol, methylcatechol, methylguaiacol, acetoguaiacone, and acetol, were detected in recycled ILs.

Although the dissolution step was conducted at low-water content, the hydrolysis step required much more water. Consequently, the high processing cost for sugar separation is a major barrier for industrialization of this process. da Costa Lopes and Bogel-Łukasik⁵²² comprehensively reviewed the challenges and possibilities of direct IL acid-catalyzed conversion of cellulose and lignocellulosic biomass.

A majority of studies on acidic IL pretreatment, however, have recently focused on acid co-solvent IL pretreatment for enhanced enzymatic hydrolysis. Partial saccharification of carbohydrates is inevitable in such systems; however, a 2- to 12-fold higher glucose conversion rate was reported from combined acid-IL pretreatment of pine than the single pretreatment of acid or IL.⁵²³ Besides, the sole use of ILs in the pretreatment usually requires a high temperature and longer reaction time.⁵¹⁵ Moreover, using an IL solution containing significant amounts of water and acid solution can reduce the expensive IL usage, in spite of significantly reducing the solubility of lignocelluloses in most ILs at above 1% water concentration. The action of acids in ILs is a catalytic role in the hydrolysis of ether linkages between adjacent glucose in the cellulose chain, consequently reducing the length of the cellulose chain⁵¹⁰ (Fig. 14). Zhang et al.⁵¹⁵ developed an optimized sugarcane bagasse pretreatment process using aqueous [BMIM][Cl] containing 1.2% HCl in the presence of 10–30% water at 130 °C for 30 min. Accordingly, a glucan digestibility of 94–100% was obtained after 72 h of enzymatic hydrolysis using HCl, in the pretreatment medium, as a more effective catalyst than H₂SO₄ and FeCl₃. Hydrochloric acid was also reported to be the most effective catalyst, amongst seven other inorganic acids studied, with [MMIM][DMP] in the pretreatment of corn stover at 110 °C for 2 h.⁵²⁴ Under these conditions, a maximum TRS yield of 92.7% was obtained after 96 h enzymatic hydrolysis.

Fig. 14 A comparison between lignocellulosic biomass pretreatment with conventional ILs and acid-based ILs (modified from ref. 510 with permission).

The first attempt for the dissolution and hydrolysis of cellulose (DP ≈ 450) in Brönsted acidic ionic liquids 1-(1-propylsulfonic)-3-methylimidazolium chloride and 1-(1-butylsulfonic)-3-methylimidazolium chloride at moderate reaction temperatures was reported by Amarasekara and Owereh⁵²⁵ in 2009. A maximum TRS yield of 62% was obtained after 1 h of preheating at 70 °C followed by 30 min heating after adding 2.0 moles equivalent of water per glucose unit. Then, they discovered a more effective dilute aqueous solution of 1-(1-propylsulfonic)-3-methylimidazolium chloride and p-toluenesulfonic acid to be a better catalyst than aqueous sulfuric acid with the same H⁺ ion concentration for the degradation of cellulose at moderate temperatures and pressures.⁵²⁶ Amarasekara and Shanbhag⁵²⁷ dissolved switchgrass biomass in 1-(alkylsulfonic)-3-methylimidazolium Brönsted acidic ILs by heating at 70 °C for 2 h (0.22 g water per g switchgrass) and obtained maximum 58.1% and 15.3% TRS and glucose yields, respectively. Li et al.⁵¹⁷ used six kinds of SO₃H-functionalized ILs based on 1-methylimidazole, 1-vinylimidazole, and triethylamine to promote the hydrolysis of MCC in [BMIM][Cl]. The acidic ILs resulted in over 83% TRS yield at 100 °C with the maximum yield of 99% for triethyl-(3-sulfo-propyl)-ammonium hydrogen sulfate. Zhuo et al.⁵¹⁹ synthesized and used six acidic ILs based on 2-phenyl-2-imidazoline for the hydrolysis of cellulose in [BMIM][Cl]. A maximum TRS yield of 85.1% was obtained by using 1-propyl sulfonic acid-2-phenyl imidazoline hydrogen sulfate, functionalized with HSO₄⁻ and Cl⁻ instead of H₂PO₄⁻, at 100 °C for 60 min and a dosage of 0.2 g water per g cellulose. The hydrolysis activity was reported to be directly related to the activity of the catalyst and also the possibility of further degradation of the resulting carbohydrates in acidic IL to HMF.⁵²⁸ Tao et al.⁵²⁹ investigated the effect of acidity and structure of fifteen SO₃H-functionalized ILs on the MCC hydrolysis and selectivity for HMF, furfural, levulinic acid (LA), and TRS yields. A maximum MCC conversion of 91.2% and selectivities for HMF, furfural, and LA of 45.7%, 26.2%, and 10.5%, respectively, were achieved in MnCl₂-containing ILs. The efficiency of Brönsted acidic ILs for the conversion of hardwood hemicellulose to pentose sugars at 160 °C was reported to be related to acid strength in the following order: [C₃SO₃HMIM][HSO₄] > [C₃SO₃HMIM][PTS] > [C₃SO₃HMIM][Cl] > [BMIM][Cl].⁵⁰⁹ Besides the activity of catalysts, solution pH is also another factor, which was investigated by Zhang et al.⁵²⁰ using different acid-catalyzed imidazolium IL solutions (80% in water) at 130 °C for 30 min for sugarcane bagasse. The pretreatment effectiveness was reported to be similar by using [BMIM][CH₃SO₃], [BMIM][CH₃SO₄], and [EMIM][Cl], at the same solution pH. Besides, by decreasing solution pH from 6.0 to 0.4, an increase in bagasse delignification, xylan removal, and consequently glucan digestibility was reported. Apart from Brönsted acidic ionic liquids, Muhammad et al.⁵¹⁸ synthesized and used an amino acid-based ionic liquid, namely 1-ethyl-3-methylimidazolium glycinate, which was capable of effectively dissolving bamboo and changed its cellulose from type I to type II.

8.5 Enhancement in enzymatic digestibility of IL-pretreated lignocelluloses

The key and widely studied role of the IL pretreatment is to enhance the enzymatic digestibility of lignocelluloses.^530–534Table 8 reviews some reports on improving the enzymatic hydrolysis of different lignocelluloses due to IL pretreatment. ILs with [BMIM] and [EMIM] cations and [Cl], [OAc], and [CH₃COO] anions have been vastly used for the pretreatment of different lignocellulosic substrates. The pretreatment conditions applied were temperature 90–160 °C, reaction times ranging from several minutes to a few hours, and a biomass loading of ca. 2–15% depending on the biomass and IL types. As reviewed in the table, enzymatic hydrolysis of different (ligno)celluloses improved significantly as a result of IL pretreatment. From the reported data, maximum 100% digestibility was reported for MCC by using [BMIM][Cl] at 90 °C for 20 min. For lignocelluloses, the TRS yield was sometimes reported and in these cases the xylose yield was also significantly improved due to IL pretreatment. For Typha capensis, for instance, a maximum reducing sugar yield of 82.4 g/100 g biomass was obtained by [BMIM][OAc] pretreatment. A high xylose yield of 87% was also obtained from switchgrass pretreated with [EMIM][Lys] at 140 °C for 1 h.

Table 8 Improvement in enzymatic digestibility of different lignocelluloses pretreated with ionic liquids (ILs)

Ionic liquid	Substrate	Pretreatment conditions	Enzymatic digestibility	Ref.
[BMIM][Cl]	Avicel	130, 140, or 150 °C for 10, 30, 60, 120, or 180 min, 5% biomass (w/w) loading	Maximum ∼80% cellulose conversion to glucose after 24 h of enzymatic hydrolysis	552
[EMIM][CH3COO]	α-Cellulose	110 °C for 40 min, 2% (w/v) cellulose loading	61% yield of glucose at 76 h	553
	Medium fibers of cellulose		69% yield of glucose at 76 h
	Long fibers of cellulose		75% yield of glucose at 76 h
	Microcrystalline cellulose		71% yield of glucose at 76 h
[EMIM][MeO(H)PO2]	α-Cellulose		67% yield of glucose at 76 h
	Medium fibers of cellulose		86% yield of glucose at 76 h
	Long fibers of cellulose		88% yield of glucose at 76 h
	Microcrystalline cellulose		75% yield of glucose at 76 h
[BMIM][OAc]	Typha capensis (TC)	110 °C for 6 h, 5.0 g IL per 0.26 g TC	Maximum reducing sugar yield of 82.4 g/100 g	554
[EMIM][OAc]	Cellulose isolated from sugarcane bagasse	90 °C for 6 h, 33.3 g IL per g cellulose	95.2% glucose yield	555
Cholinium amino acids ILs	Rice straw	90 °C for 5 h	Maximum glucose and xylose yields of 84.0% and 42.1%, respectively	556
Cholinium lysine IL ([Ch][Lys] IL)-water mixtures	Rice straw	20% [Ch][Lys]-water mixture at 90 °C for 1 h	Maximum sugar yields of 81% for glucose and 48% for xylose	557
[EMIM][OAc]	Sugarcane bagasse	150 °C, 90 min and 5% bagasse in IL	83% and 21% glucan and xylan digestibility, respectively	558
[EMIM][OAc]	Energy cane bagasse	120 °C for 30 min, 5% (w/w) biomass loading	87.0% and 64.3% glucan and xylan digestibility, respectively	125
[EMIM][OAc]	Pinus radiate compression wood	120 °C for 3 h	93% glucan digestibility, 65% xylan digestibility, and 39% mannan digestibility after 24 h	559
1-Hexylpyridinium chloride	Avicel and bagasse	80 °C or 100 °C, 5% (w/w) loading	Over 95% conversion to glucose after 24 h for Avicel, and 1–3-fold higher conversion than untreated biomass for bagasse	560
1-Butylimidazolium hydrogen sulfate	Miscanthus giganteus	120 °C for 15 min up to 24 h, 10% (w/v) biomass loading	Recovery of up to 90% of the glucan as fermentable glucose and up to 39% saccharification yield for hemicellulose	561
[EMIM][OAc]–DMSO solutions	Eucalyptus	Ratios of 4 : 1, 3 : 2, 2 : 3 and 1 : 4 (v/v) [EMIM][OAc]-to-DMSO, 15% (w/v) biomass loading, at temperatures ranging from 80 to 140 °C	95% of glucose theoretical maximum yield and up to 65% xylose yield	562
Chloride, acetate, and formate based IL	MCC	90 °C for 20 min	100% digestibility by using [BMIM][Cl]	563
[EMIM][OAc]	Rice husk	100 °C for 10 h, 1.5% (w/v) biomass loading	42.1% reducing sugar yield	501
[EMIM][OAc]	Agave bagasse (AGB) and switchgrass (SWG)	120 and 160 °C for 3 h and 15% biomass loading	Increase in TRS by 100% for SWG and by 183% for AGB	500
[EMIM][OAc]	Bagasse	Optimum conditions: 145 °C, 15 min and 14 wt% solid loading	69.7% of RS yield	564
[EMIM][OAc]	Sugarcane bagasse	8 min at 140 °C, 25 wt% biomass loading	Glucose yields of more than 90% after 24 h of enzymatic saccharification and maximum xylose yield of ca. 85%	479
[EMIM][OAc]	Switchgrass	160 °C and 3 h, 15% biomass loading	Glucose and xylose yields of 94.8% and 62.2%, respectively	480
1-Butyl-3-methylimidazolium methyl sulfate and 1-butyl-3-methylimidazolium hydrogen sulfate	Miscanthus giganteus, pine (Pinus sylvestris), and willow (Salix viminalis)	120 °C and 2 h, 10% (w/v) biomass loading	Up to 90% of the glucose and 25% of the hemicellulose by the combined ionic liquid pretreatment and the enzymatic hydrolysis	565
[EMIM][OAc]	Eucalyptus globulus	120 °C for 3 h, 9.7 g IL and 0.3 g biomass	37 and 30% glucose and xylose yields, respectively, after 4 h enzymatic hydrolysis	566
1-Ethyl-3-methylimidazolium acetate	Wheat straw	Temperature (130–170 °C), time (0.5–5.5 h) and ionic liquid concentration (0–100%), biomass loading 5% (w/w)	71.4% sugars recovery under optimum conditions of 158 °C, IL concentration, 49.5% (w/w), and 3.6 h	567
[EMIM][OAc]	Triticale straw	150 °C for 90 min, 1.5 g straw to 48.5 g of water-IL mixture	81% fermentable sugar yield	568
[EMIM][OAc] and [BMIM][Cl]	Cotton cellulose	Microwave irradiation or 110 °C for 30 min, 2% (w/v) biomass loading	At least 12-fold and by 50-fold enhancement in enzymatic hydrolysis at 110 °C and microwave irradiation, respectively	569
[BMIM][Cl]	Sugarcane bagasse	Temperatures (110–160 °C) and times (30–180 min); 0.25 g bagasse in 5 g IL (≤5% impurities and 2% moisture)	Optimum conditions: 150 °C for 90 min complete (100%) and rapid (3 h) glucan saccharification up to 70% xylan solubilization	570
[BMIM][Cl]	Cotton	130 °C for 20 min, 5% w/w	At least 4-fold enhancement on cellulose saccharification conversion	571
[EMIM][CH3COO]	Wood flour	Various temperatures, 5% w/w biomass loading	>90% conversion of cellulose	486
Choline acetate (ChOAc)	Bagasse	IL/ultrasound-assisted pretreatment (60 min at 24 kHz and a power of 35 W), 0.25 g bagasse in 5 g IL	Cellulose and hemicellulose saccharification percentages 80% and 72%, respectively, in situ saccharification for 48 h	572
Choline formate (ChFor), choline acetate (ChOAc), and choline propionate (ChPro)	Kenaf powders	Microwave heating or 110 °C for 20 min, 5% w/w biomass loading	20% cellulose conversion for regular heating and 60–90% for microwave heating	573
[BMIM][Cl]	Sweet sorghum bagasse	110 °C for 1 h, 10% w/w biomass loading	Approximately 40% conversion of cellulose after 60 h enzymatic hydrolysis	574
[BMIM][Cl]	Populus tomentosa Carr.	130 °C, 0.5 g of the substrate and 9.5 g of the IL	92% glucose yield after 72 h enzymatic hydrolysis	349
[EMIM][Cl] and [EMIM][OAc]	Pine wood	80, 100, or 120 °C for 3 h with stirring, 0.35 g wood and 7.0 g IL	Glucan conversions ranging from 23% to 84% with [EMIM][OAc] being more effective than [EMIM][Cl]	575
[BMIM][Cl]	Oil palm frond (OPF)	Temperatures less than 100 °C and times less than 1 h, and maximum loadings of 10%	100% glucose recovery with pretreatment conditions: 80 °C, 15 min, and 10% solid loading	576
[EMIM][OAc]	Panicum virgatum (switchgrass)	90 °C for 5 h, 10% (w/w) biomass loading	Glucose yield: 31%; xylose yield: 29%	577
[EMIM][Lys]			Glucose yield: 70%; xylose yield: 68%
[Ch][Lys]			Glucose yield: 42%; xylose yield: 58%
[Ch][OAc]			Glucose yield: 27%; xylose yield: 23%
[EMIM][OAc]		140 °C for 1 h, 10% (w/w) biomass loading	Glucose yield: 65%; xylose yield: 86%
[EMIM][Lys]			Glucose yield:59%;xylose yield: 87%
[Ch][Lys]			Glucose yield: 61%; xylose yield: 82%
[Ch][OAc]			Glucose yield: 55%; xylose yield: 79%
[EMIM][OAc]	Poplar and switchgrass	120 °C for 30 min, 5% (w/w) biomass loading	∼70% and 46% glucan conversion after 24 h enzymatic hydrolysis for poplar and switchgrass, respectively	578
[EMIM][MeO(H)PO2] and [EMIM][CH3COO]	Cotton cellulose	45 and 25 °C for 20 min, 2%, w/v biomass loading	Glucose yield after 24 h of enzymatic hydrolysis: 58.5% and 45.4%	579
[EMIM][OAc]	Spruce and oak sawdust	110 °C for 40 min, 2% w/v biomass loading	Up to 7 times increase in enzymatic saccharification compared with the untreated substrate	580
[AMIM][Cl]	Cotton-based waste textiles were	90, 110, and 130 °C until 2% (w/w) biomass was dissolved	7 times higher yield of fermentable sugars than untreated fabrics	581
[EMIM][OAc]	Oil palm empty fruit bunch (OPEFB)	130 °C, 2 h, 5% (w/w) biomass loading	95.5% enzymatic digestibility of glucan	582
[BMIM][Cl]			54.8% enzymatic digestibility of glucan
MTBS			22.0% enzymatic digestibility of glucan
[EMIM][DEP]			48.9% enzymatic digestibility of glucan
[EMIM][DEP]	Wheat straw	130 °C for 30 min, 4% (w/w) biomass loading	54.8% reducing sugar yield after being enzymatically hydrolyzed for 12 h	583
ChOAc	Bamboo powder	110 °C for 60 min, and ultrasonic pretreatment in the same IL at 25 °C for 60 min, 0.5 g bamboo in 5 g IL	55% and 92% cellulose saccharification for regular heating and ultrasonic pretreatment, respectively	584
[EMIM][OAc]	Beechwood chips	115 °C for 1.5 h, 500 mg or 1 g of the wood in IL to obtain a mass of 10 g	Cellulose conversion of 90.2 wt% for a hydrolysis time of 72 h	585

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8.6 Challenges with in situ enzymatic hydrolysis of lignocelluloses in aqueous-IL media

Unlike water and buffers, which are capable of dissolving enzymes without unfolding their active structure, the biocatalysis activity in organic solvents may be hampered by a variety of factors. Most cellulases and other hydrolytic enzymes are deactivated in the presence of ILs, even at low concentrations.^535,536 A comprehensive review on the enzymatic hydrolysis of lignocelluloses in the presence of ionic liquids, or the so-called in situ or one-pot pretreatment and hydrolysis, has been recently published by Wahlström and Suurnäkki.⁵³⁷ This review was mainly focused on the ways to keep the cellulase enzyme active for enzymatic hydrolysis in the presence of ILs. The hydrolytic enzyme stabilization techniques include enzyme immobilization, e.g., by encapsulation⁵³⁸ or thermostabilization.⁵³⁹ Besides, the discovery and development of IL-tolerant enzymes,^540,541e.g., enzymes isolated from thermophilic and halophilic microbes,^542,543 are of great importance in this regard. A review on various cellulase stabilization techniques for the single-step process and the design of enzyme compatible biomass-dissolving ILs was recently published by Elgharbawy et al.⁵⁴⁴ The recent trends in IL-tolerant enzymes and microorganisms were also critically reviewed by Portillo and Saadeddin⁵⁴⁵ and Xu et al.⁵⁴⁶

It is notable here to mention that the residual ILs in the enzymatic hydrolysates inhibit the growth and productivity of microorganisms in downstream and fermentation processes.^547,548 The residual [EMIM][OAc] in the hydrolysates (higher than 0.1%) was reported to inhibit the growth and ethanol production by S. cerevisiae, suggested due to a potential synergistic effect between this particular combination of anions and cations.⁵⁴⁹ The water-wash step results in a significant sugar loss and generation of large amounts of wastewater. To address this issue, Xu et al.⁵⁵⁰ recently developed a one-pot conversion process via using dilute bio-based ILs to produce high-titer cellulosic ethanol. Moreover, a novel CBP process was developed for ethanol production using IL pretreatment by cellulase-displaying yeast and approximately 90% ethanol yield was reported.⁵⁵¹

Another challenge in enzymatic in situ saccharification of lignocelluloses is the recovery of sugars produced during the hydrolysis in IL media. Chromatographic techniques⁵⁸⁶ and membrane-based methods⁵⁸⁷ have been suggested for the separation and recovery of sugars and ILs from biomass hydrolysates. This challenge also applies to the case of acid-catalyzed hydrolysis in ILs,⁵⁸⁸ and it is considered a major challenge and a drawback in using ILs as pretreatment agents. However, in most cases, the recovery of sugars and the recycling of ILs occur simultaneously in a single process.

8.7 Recovery and reuse of ionic liquids

Due to the current high price of ILs for an economically viable pretreatment process, efficient recovery and recycling of ILs is vital.^589–592 Besides, the wastage of ILs can cause environmental issues associated with slow degradation and toxicity to downstream processes^589,590 Mai et al.⁵⁹³ reviewed different methods for the recovery of ILs in detail. Here, we discuss briefly the methods of IL recovery with application in the IL pretreatment of lignocelluloses.

The most widely used method for the recovery and recycling of ILs from IL-anti-solvent–lignocellulose systems is distillation.^{580,590,594–596} The method consists of evaporating anti-solvent (e.g., water and alcohol) after removing precipitated lignocellulose from the pretreatment media. Since a large quantity of precipitating solvent is required to prevent gel phase formation, the evaporation step needs a lot of energy and often presents environmental problems. When water is used for the precipitation, the situation is even worse, because of its high specific heat capacity and the high solubility of produced biomass compounds, e.g., monomeric and small oligomeric carbohydrates.^590,596 Approximately, 85–90% recovery of [EMIM][OAc] was reported in an IL–water system via distillation by Qui et al.⁵⁹⁴

The ability of ILs to form aqueous biphasic systems with a kosmotropic anion, e.g., phosphate, carbonate, or sulfate, was first reported by Gutowski et al.,⁵⁹⁷ which can be utilized to recycle hydrophilic ILs from aqueous solutions. The upper IL-rich phase can be easily recovered by simple decantation or a magnetic field. However, further separation is required in order to extract water and remaining monosaccharide hydrolysates from the IL solution.⁵⁹⁸ The recovery of ILs in these systems depends on the salt type and concentration as well as the IL cation and anion type.⁵⁹⁹ A recovery of over 95.0% for [BMIM][OAc] in K₃PO₄-containing systems (pH 12–13) was reported.⁵⁹⁹

Apart from these traditional separation methods, recently, the so-called green processes were developed based on chromatography (resin and alumina column chromatography)^598,600 and electrodialysis^601–603 for the IL recovery.

Unfortunately, the recovered ILs do not sometimes show the same performance in the pretreatment as their virgin forms.^594,604 Qiu et al.⁵⁹⁴ reported a decrease in recycled [EMIM][OAc], by evaporation, performance in pretreatment of energy cane bagasse. This phenomenon could be attributed to the accumulation of IL degradation products in the pretreatment process which affect the recycling efficiency and properties of ILs. Besides, the recycled ILs may contain carbohydrate monomers and oligomers and biomass decomposition products. It is most likely for recycled ILs to have both of the mentioned impurities that negatively affect their performance in pretreatment.¹⁷⁵ On the other hand, Auxenfans et al.⁵⁸⁰ reported the same ability and similar efficiency of recycled [EMIM][OAc], via distillation by rotary evaporation, to the fresh IL in enzymatic saccharification performance for pretreatment of industrial wood sawdust. In another study, they again reported the similar performance of two recycled imidazolium-based ILs in maintaining their efficiency to pretreat cellulose.⁵⁷⁹

8.8 IL pretreatment of lignocelluloses for enhanced biogas and renewable chemical production

Different pretreatment methods for enhanced biogas production from lignocelluloses have been comprehensively reviewed by Zheng et al.⁶⁹ Mancini et al.⁶⁰⁵ also reviewed the solvent pretreatments of lignocellulosic materials to enhance biogas production. Nonetheless, there are few studies in the literature on the enhancement of biogas production from lignocelluloses by using IL pretreatment. Gao et al.⁶⁰⁶ pretreated water hyacinth, rice straw, mango leaves, and spruce with [C_nMIM][Cl] (n = 2, 4, and 6) under different conditions and evaluated the effect of the pretreatment on biogas production. The maximum enhancement of biogas production was obtained by pretreatment with [BMIM][Cl] at 120 °C for 2 h for water hyacinth followed by spruce, while maximum methane production from rice straw and mango leaves, i.e., 233 and 125 mL g⁻¹ carbohydrates, was obtained for pretreatment at 140 °C for 2 h and 140 °C for 8 h, respectively. They also obtained an increase in biogas yield and methane concentration by 16.3–97.6% and 13.2–28.3%, respectively, from water hyacinth by [BMIM][Cl] and DMSO co-solvent pretreatment at 120 °C for 120 min.⁶⁰⁷ Li and Xu⁶⁰⁸ reported severe toxicity of imidazolium-based ILs in anaerobic digestion of grass (1 : 10 ratio). However, they reported a low toxicity and high recyclability potential for [BMIM][OAc] in the methane production of 221 mL g⁻¹-VS from grass.

Most of the studies on using IL pretreatment for the production of renewable products from lignocelluloses have focused on the hydrolysis of pretreated biomass into sugar-rich hydrolysates (cf. Section 8.5), which are then used by microorganisms for carbon and energy sources, e.g., for microbial lipid production. For example, Gong et al.⁶⁰⁹ prepared corn stover solids by [EMIM][OAc]–N-methylpyrrolidone (NMP) pretreatment at 140 °C and converted the pretreated solids to microbial lipids using Cryptococcus curvatus via a simultaneous saccharification and enhanced lipid production process. They obtained maximum 112 mg g⁻¹ pretreated biomass lipid yield with efficient co-utilization of cellulose and hemicellulose. Xie et al.⁶¹⁰ also used 20% (mole fraction) [EMIM][OAc] in NMP at 140 °C for 60 min to pretreat corn stover followed by enzymatic hydrolysis for the cultivation of Rhodosporidium toruloides Y4 for lipid production. The oleaginous microorganism utilized both C₆ and C₅ sugars in the hydrolyzates and produced a moderate 15.2 g L⁻¹ biomass yield and 36.4% lipid yield. Bokinsky et al.⁶¹¹ reported the synthesis of biofuels, i.e., fatty acid ethyl esters, butanol, and pinene, from [EMIM][OAc] pretreated switchgrass using engineered Escherichia coli which can express cellulase, xylanase, β-glucosidase, and xylobiosidase enzymes. They reported that the IL pretreatment made the biomass completely susceptible to hydrolysis.

On the use of ILs for renewable chemical production from lignocelluloses, Huang et al.⁶¹² evaluated the effects of residual [EMIM][Cl], [EMIM][DEP], and [EMIM][OAc] on the lipid production by oleaginous yeast Rhodosporidium toruloides AS 2.1389. By adjusting pH to 6.0 in the presence of 30 mM ILs, minor inhibition effects were reported, while the presence of 60 mM ILs caused a significant inhibition on the yeast. Liu et al.⁶¹³ also reported that the residual ILs in the hydrolysate of rice straw inhibited the growth and lipid accumulation by Geotrichum fermentans. The inhibition was induced by both the anion and cation of the ILs used, and the side chain of the cation showed a clear inhibition.

Varanasi et al.⁶¹⁴ focused on the production of lignin-based renewable chemicals from different lignocelluloses by selective breakdown of lignin using [EMIIM][OAc] pretreatment at 120 and 160 °C for 6 h. The generated chemicals, (i.e., phenols, guaiacols, syringols, eugenol, and catechols), their oxidized products (i.e., vanillin, vanillic acid, and syringaldehyde), and their easily derivatized hydrocarbons (i.e., benzene, toluene, xylene, styrene, biphenyls, and cyclohexane) were produced from lignin by tuning the process conditions. The production of levulinic acid directly during the IL pretreatment was reported in some studies.^615,616 Muranaka et al.⁶¹⁶ successfully converted 60.7% (72.9 mol%) of cellulose into levulinic acid by using [EMIM][Br] and [EMIM][P] at 80–120 °C for 1–6 h under stirring. Sun et al.⁶¹⁵ used a series of heteropolyacid (HPA) ILs to catalyze one-pot depolymerization of cellulose into glucose and subsequent levulinic acid (up to a 60% yield) in a water–methyl isobutyl ketone (MIBK) biphasic system. Xiao et al.⁶¹⁷ optimized the catalytic conversion of cellulose into HMF by AlCl₃ in DMSO–[BMIM][Cl] mixtures. They obtained a maximum HMF yield of 54.9% from cellulose at 150 °C after 9 h in a mixed solvent of DMSO–[BMIM][Cl] (10 wt%).

8.9 Techno-economic analysis of ionic liquid pretreatment

For economic evaluation of IL pretreatment of lignocelluloses, Sen et al.⁶¹⁸ identified the IL cost as the major cost driver in IL pretreatment. They suggested lower IL consumption and/or effective separation strategies to improve the economy of IL pretreatment. Konda et al.⁶¹⁹ compared the cost drivers and economic potential of two variants of IL pretreatment for ethanol production: first based on complete removal of the IL prior to hydrolysis and second based on a one-pot process. At a high biomass loading of 50%, both routes were reported to be economically viable with a minimum ethanol selling price of $6.3 per gal. With more reduced water and acid/base consumption in the first and second routes, respectively, improved pretreatment efficiency, and by lignin valorization, the minimum ethanol selling price could be reduced to $3.2 for the former route and $2.8 for the latter route. Baral and Shah⁶²⁰ conducted a techno-economic comparison study on [EMIM][OAc] and sulfuric acid pretreatment of corn stover, poplar, and switchgrass, and estimated a sugar production cost of 2.7, 3.2, and 3.0 $ per kg, respectively. They further reported that optimistic considerations of at least 97% IL recovery, less than $1 per kg IL cost, and >90% heat recovery are required to have an economically competitive IL pretreatment. To improve the process economy, George et al.⁶²¹ attempted to lower the cost of IL, which is one of the main impediments to IL utilization in the pretreatment step, by designing a number of low-cost and stable protic ILs based upon the [HSO₄] anion with promising potential in the pretreatment. The most effective solvent, triethylammonium hydrogen sulfate IL, demonstrated approximately 75% as effective as [C₂C₁IM][OAc] for switchgrass. A set of new low-cost RTILs based on butylammonium prepared by reacting carboxylic acids with aliphatic amine cations under ambient conditions was also synthesized and characterized by de Andrade Neto et al.⁶²² for lignocellulose hydrolysis applications. Among them, n-butylammonium acetate favored the subsequent acid hydrolysis of corn fiber. Oleskowicz-Popiel et al.⁶²³ also reported the acidolysis of IL pretreated lignocelluloses as a more economically viable route than using costly enzymes for saccharification. They further calculated that the minimum ethanol selling price could be reduced to $4.00 per gal, when the performance of the hydrolysis, extraction, and sugar recovery is improved. Socha et al.⁶²⁴ synthesized some tertiary amine-based ILs from aromatic aldehydes derived from vanillin, p-anisaldehyde, and furfural, and confirmed their effectiveness in switchgrass pretreatment. Their approach of producing renewable ILs from lignocelluloses in a so-called “closed-loop” can be a solution for the drawback of expensive ILs in the pretreatment.

8.10 Present status and future prospects of IL pretreatment

Taking all these points into consideration, ILs, as powerful non-derivatizing cellulose solvents, have been recently subjected to vast studies for lignocellulose dissolution and regeneration. The promising features of using ILs for biomass pretreatment are negligible vapor pressure, thermal stability, non-flammability, and high polarity, and being “green” solvents in many cases.^625–627 They are capable of fractionating a variety of lignocelluloses, reordering or restructuring the hydrogen bonds in the cellulose network, decreasing cellulose crystallinity, and increasing cellulose accessibility to cellulases. The pretreatment requires low equipment costs with low energy consumption. The regenerated materials are more susceptible to enzymatic hydrolysis than the untreated form, with comparable or even superior yields of fermentable sugars, than the conventional pretreatments.^628,629 By adjusting the ILs' anion and cation, different ILs are synthesized to tune their properties. Mora-Pale et al.⁶³⁰ stated that lignin released during RTIL pretreatment of lignocelluloses is likely to be far more “pristine” than Kraft lignin, and has general applications in phenol-formaldehyde replacements, conversion into liquid fuels following hydrogenative depolymerization, or possibly into specific low molecular weight chemicals.

A process diagram for bioconversion of lignocelluloses to ethanol and biogas using IL pretreatment is proposed in Fig. 15. The process can be applied for the production of other fermentative chemicals via a sugar platform as well. Besides, this process is advantageous in the production of other byproducts, which can improve the overall process economy.

Fig. 15 A process diagram for IL pretreatment of lignocelluloses for ethanol or biogas production applying two routes: (1) one-pot/in situ hydrolysis and (2) separated enzymatic hydrolysis and fermentation, with IL and solvent recycling.

However, IL pretreatment of lignocelluloses is facing several technological and economic challenges. Although some efforts have been made to design low-cost ILs^621,631 (as discussed in Section 8.9), still a major obstacle in implication of many ILs on a large scale is their high price. Efficient recovery and recycling of ILs are crucial in order to reduce the inhibitory effects of ILs on subsequent enzymatic hydrolysis and fermentation. Besides, the residual ILs in waste streams can cause environmental problems depending on their degradability.⁶³² Although research is under way using amino acids to synthesize biodegradable ILs, most ILs for biomass processing are not easily biodegradable. However, a cost-effective technology, despite the ease of recycling via distillation, is needed to make the process competitive to conventional pretreatment strategies. This problem is not only for IL recovery, but also for the anti-solvent used in the regeneration process. Although many ILs, e.g., [EMIM][OAc] and [AMIM][Cl], were reported to be excellent solvents for cellulose dissolution,⁶³² the selection of ILs for biomass pretreatment should compromise between solubilizing power and compatibility with enzymes and/or organisms. In the case of acidic ILs, the rate of formation of degradation products should be manipulated by the side chains of the cation. Not all, but some ILs are corrosive, toxic, and hygroscopic, which should be utilized with care.

9 Comparison of IL, CPA, and NMMO pretreatments

A few studies compared the effectiveness of the IL, CPA, and NMMO pretreatments in improving enzymatic hydrolysis yield and fuel/chemical production.^{165,386,571,633–637} Wheat straw was pretreated with CPA (85%, 50 °C for 1.5 h), NMMO (130 °C for 2 h), and IL ([AMIM][Cl], 110 °C for 1 h) and the results showed that the most prominent difference in the chemical composition of wheat straw was >90% solubilization of xylan due to CPA pretreatment, while for the other two treatments, xylan was solubilized <10%.⁶³³ Phosphoric acid acts as a Brønsted acid catalyst and generally promotes the hydrolysis of glycosidic bonds in cellulose/hemicellulose, which leads to considerably higher solubilization of xylan. Moreover, cellulose hydrolysis followed the order of CPA > NMMO > [AMIM][Cl]. Similarly, CPA was more successful in pretreatment of corn stover than IL, whereas, compared to 96% glucan digestibility of corn stover pretreated with CPA, a 55% glucan digestibility was obtained after pretreatment with [BMIM][Cl]. A tradeoff between cellulose disruption and the inhibitory effects of the presence of residual lignin and residual cellulose solvent on the pretreated biomass was reported as the main reason for incomplete hydrolysis for IL pretreatment. However, cellulose digestibilities of 100% and 92% were obtained with CPA and [BMIM][Cl], respectively, for Avicel cellulose as a substrate.⁶³⁴

In a study, high glucose yields were obtained for rice straw following pretreatment with NMMO and [BMIM][OAc], and the obtained yields were comparable for both pretreatments.³⁸¹ The crystallinity index and total crystallinity of the substrate were quite similar for both pretreatments. Similarly, the glucan conversion (after 72 h) for Populus tomentosa pretreated with IL ([BMIM][Cl] at 130 °C), NMMO (130 °C for 30 min), and CPA (85% at room temperature) was reported to be 80%, 82%, and 92%, respectively.⁶³⁵ The results also showed that the hydrolysis rate for the IL pretreated sample was higher than that of the CPA pretreated sample. These results were possibly due to the transformation of cellulose I to amorphous cellulose and cellulose II in IL and CPA pretreatments, respectively. CPA was reported as a better cellulose solvent than NMMO and [BMIM][Cl] in improving the saccharification rate and yield of cotton cellulose due to the high specific surface area and low DP for CPA pretreated cellulose.⁶³⁶

On the other hand, IL pretreatment ([EMIM][Br] and [EMIM][P]) was reported to be more effective than CPA pretreatment in converting cellulosic substrates to levulinic acid.⁶³⁷ Decrease in cellulose crystallinity, solubilization of cellulose, and IL interaction with cellulose were the determinant factors for higher yields.

In summary, the effectiveness of different cellulose solvents in the pretreatment of lignocelluloses is strongly dependent on biomass type and final chemicals produced. In contrast to phosphoric acid, residual ILs and NMMO cause inhibitory effects on biotechnological downstream processes.

10 Concluding remarks

Regarding the high worldwide fuel demand and the significant potential for biomass conversion to offset fossil fuel usage, a high number of studies and efforts have been made in the past several decades in the cellulosic fuel area. The cellulosic fuel production process involves four major steps of biomass preparation, pretreatment, hydrolysis, and fermentation, with the pretreatment step being one of the most cost contributing and the rate and yield limiting step. Given their significant ability to fractionate the lignocellulosic structure, cellulose solvents are excellent starting points for industrial biorefinery applications. Although this category of pretreatment has several advantages over other conventional pretreatments, which was the focus of this review, several issues should be addressed to make the process economically and environmentally viable. One of the promising features of cellulose solvents for the pretreatment of lignocelluloses is associated with their properties as “green” solvents. They should have negligible vapor pressure to help their recyclability, and should be easily biodegradable. Using cellulose solvents for the pretreatment of lignocelluloses is advantageous due to their application at high solid loadings and relatively low pressure/temperatures with no/less chemical modification and inhibitory byproduct formation. The solvents can be oriented towards different purposes for the pretreatment of lignocelluloses. They can be selected for (1) the separation of mainly cellulose, (2) the separation of mainly hemicellulose, (3) the separation of mainly lignin, (4) opening the compact structure, (5) regeneration and structural modification, and (6) cellulose crystallinity reduction, although more than one of these actions typically take place. Organic cellulose solvents, e.g., concentrated phosphoric acid and concentrated NaOH, are capable of removing and/or reorganizing the hydrogen bond network structure of cellulose, decreasing cellulose crystallinity, and enhancing cellulose accessibility to cellulases, and consequently enhancing glucan digestibility even at low cellulase loadings that is vital to reduce the overall process cost. There are some cellulose solvents, e.g., NMMO, that modify the structure of lignocelluloses by dissolution and regeneration of the whole biomass, without significant lignin/hemicellulose removal. Meanwhile, ILs are target-oriented solvents that can be designed to separate specific parts of lignocelluloses. The high revenues from co-products (acetic acid, lignin, and hemicelluloses) of the pretreatment can drastically improve the economy of cellulosic fuels. However, the cellulose solvent-based pretreatment of lignocelluloses is still in its early stage of development, mainly in the laboratory scale, and facing several technological and economic challenges. The efficient recovery of the solvents, which is usually a high energy intensive process, is necessary because of not only the high price of the solvents, but also because of the inhibitory effects of the solvents on subsequent processes. Furthermore, substantial reduction in the use of chemicals (both the cellulose solvents and organic solvents) is required in order to have an economically competitive process.

Conflicts of interest

There are no conflicts to declare.

Abbreviation

AD	Anaerobic digestion
[BMIM][Cl]	1-Butyl-3-methylimidazolium chloride
CBM	Cellulose binding module
CPA	Concentrated phosphoric acid
CBP	Consolidated bioprocessing
CAC	Cellulose accessibility to cellulase
COSLIF	Cellulose solvent- and organic solvent-based lignocellulosic fractionation
CrI	Crystallinity index
[EMIM][OAc	1-Ethyl-3-methylimidazolium acetate
GHG	Greenhouse gas
IL	Ionic liquid
LOI	Lateral order index
NREL	National renewable energy laboratory
NMMO or NMO	N-Methyl-morpholine-N-oxide
RTIL	Room-temperature ionic liquid
SCO	Single cell oil
SAA	Soaking in aqueous ammonia
SS	Simons' staining
SSA	Specific surface area
TCI	Total crystallinity index
TRS	Total reducing sugar
VFA	Volatile fatty acid
WRV	Water retention value

Acknowledgements

The corresponding author would like to acknowledge support from the Office of Biological and Environmental Research in the US Department of Energy (DOE) Office of Science through the BioEnergy Science Center (BESC) and the Center for Bioenergy Innovation (CBI), both at Oak Ridge National Laboratory.

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