**Observation of Potential Contaminants in Processed Biomass Using Fourier Transform Infrared Spectroscopy**

**Jingshun Zhuang <sup>1</sup> , Mi Li <sup>2</sup> , Yunqiao Pu <sup>3</sup> , Arthur Jonas Ragauskas 2,3,4,\* and Chang Geun Yoo 1,3,\***


Received: 2 June 2020; Accepted: 22 June 2020; Published: 24 June 2020

**Abstract:** With rapidly increased interests in biomass, diverse chemical and biological processes have been applied for biomass utilization. Fourier transform infrared (FTIR) analysis has been used for characterizing different types of biomass and their products, including natural and processed biomass. During biomass treatments, some solvents and/or catalysts can be retained and contaminate biomass. In addition, contaminants can be generated by the decomposition of biomass components. Herein, we report FTIR analyses of a series of contaminants, such as various solvents, chemicals, enzymes, and possibly formed degradation by-products in the biomass conversion process along with poplar biomass. This information helps to prevent misunderstanding the FTIR analysis results of the processed biomass.

**Keywords:** poplar; FTIR; contaminants; by-products

### **1. Introduction**

A proper understanding of biomass characteristics is important for the effective utilization of biomass. It not only provides natural properties of biomass but also tells the influences of the applied processes on the biomass structures. Characteristics of biomass have been investigated in different aspects, including physical, chemical, thermal, mineral, and surface properties. For a better understanding of biomass and its products, several different analytical approaches have been developed using diverse analytical techniques such as high-performance liquid chromatography (HPLC), gas chromatography (GC), gel permeation chromatography (GPC), nuclear magnetic resonance (NMR), time-of-flight secondary ion mass spectrometry (ToF-SIMS), X-ray, transmission electron microscopy (TEM), scanning electron microscopy (SEM), differential scanning calorimetry (DSC), thermogravimetric analysis (TGA), and Fourier transform infrared (FTIR) for measuring carbohydrate contents, identification and quantity of products, molecular weights, structural information such as linkages and composition, spatial distribution of molecules and chemical structures on surface, crystallinity, morphological characteristics in nano- and micro-scales, thermal properties (melting point, glass transition temperature, etc.), functional groups and chemical bonds, and other important information of biomass and its products and by-products [1–9]. Among these methods, FTIR spectroscopy is one of the most widely applied analytical methods to study the functional groups of biomass by measuring the absorption bands of samples [10]. It provides qualitative and semi-quantitative information for functional groups of biomass by determining the presence of fundamental molecular vibrations [11]. Its Fellgett and Jacquinot advantages allow for rapid and ready characterization compared to many other biomass analysis methods [12]. Moreover, it does not need any modification and/or deconstruction of biomass; therefore, original properties can be monitored as the sample is. Despite these advantages, the characterization of biomass using FTIR is still challenged by overlapping the bands from different biomass components and/or unexpected impurities from the applied catalysts and solvents. In particular, fingerprint regions are complicated to identify because of many series of absorptions. For fast and reliable analysis of the substances from different processing, detection, and identification of possible contaminants are very important.

Lignocellulosic biomass is a heterogeneous matrix. Due to the complicated composition and structural properties of biomass, single or multi-stage pretreatment/preprocessing is necessary for its utilization, isolation, and analyses. Various chemicals, such as organic solvents, acids, alkalis, and inorganic salt solutions have been applied for isolation, pretreatment, conversion, and other reactions on biomass [13–19]. Biological catalysts, such as enzymes, have also been used in many biomass conversion processes or characterization methods [20]. Besides, each biomass component could be decomposed and/or modified under severe process conditions [21]. The presence of these chemicals and by-products are considered as impurities and could potentially affect their characterization results; therefore, they should be completely removed after the processes. Unfortunately, these components are possibly retained on the surface of biomass after these preprocessing and cause misinterpretation of the targeted biomass structure by their overlapped FTIR spectra. Besides the misreading of the biomass properties, the detection of contaminants can be used to determine the necessity of biomass washing step. Although the IR assignments of many chemicals and solvents are available individually, their actual contaminations are not easily detected due to the spectra of biomass itself. In this study, poplar biomass was mixed with known chemicals and enzymes, which are potential contaminants, and their overlapped FTIR spectra in each sample were identified and discussed.

#### **2. Materials and Methods**

#### *2.1. Materials*

Poplar was harvested in the Oak Ridge National Laboratory in 2008. Prior to the FTIR analysis, the sample was Wiley-milled and screened to 0.42 mm. Extractives were removed from the original poplar sample (~10 g) by toluene/ethanol Soxhlet extraction (2:1, *v*/*v*, 200 mL) for 8 h followed by 6 h of water extraction. All chemicals (acetone, ethanol, methanol, tetrahydrofuran, dioxane, toluene, glycerol, chloroform, pyridine, sulfuric acid, hydrochloric acid, phosphoric acid, acetic acid, sodium hydroxide, ammonium hydroxide, 1-butyl-3-methylimidazolium chloride, 1-benzyl-3-methylimidazolium chloride, choline chloride, urea, *p*-hydroxybenzoic acid, 4-hydroxybenzaldehyde, *p*-coumaric acid, hydroxymethylfurfural, and furfural) and enzymes (cellulase and β-glucosidase) used in this study were purchased from VWR, Sigma-Aldrich, or Fisher Scientific. Deep eutectic solvents (DESs) formed by combining hydrogen bonding donors (HBDs: urea, *p*-hydroxybenzoic acid, 4-hydroxybenzaldehyde, *p*-coumaric acid) and hydrogen bonding acceptor (HBA: choline chloride) at 80 ◦C prior to the FTIR analysis.

#### *2.2. Isolation of Cellulose, Hemicellulose, and Lignin*

Cellulose, hemicellulose, and lignin were isolated from the extractives-free poplar, as described in the previous studies [2,22]. In brief, the biomass was delignified using peracetic acid at 25 ◦C with 5% (wt/wt) solid loading for 24 h. The remaining solid, holocellulose, was air dried for 24 h. Two-step alkali extraction with 17.5% (wt/wt) and 8.75% (wt/wt) sodium hydroxide was conducted at 25 ◦C for 2 h in each step. The remaining solid fraction was called α-cellulose after being air dried, and the

liquid fraction was neutralized with anhydrous acetic acid and mixed with ethanol three times to precipitate hemicellulose.

Cellulolytic enzyme lignin (CEL) was separated from the poplar samples. The extractives-free poplar was ball-milled using Retsch PM 100 at 600 rpm for 2 h. The ball-milled sample was hydrolyzed at 50 ◦C with the CTec2 enzyme in acetate buffer solution (pH 4.8) for 48 h twice. The residual solid was extracted with 96% dioxane for 48 h. The dioxane-extracted fraction was recovered at 40 ◦C by rotary evaporation and freeze drying and used for further analysis.

#### *2.3. Fourier Transform Infrared (FTIR) Analysis*

To observe the FTIR spectra of contaminants from the spectra of biomass clearly, about 30–50 µL of contaminant was loaded to 0.3 g of extractives-free poplar in 20 mL glass vial and mixed by vortexing prior to the analysis. The prepared DESs were loaded to biomass and physically mixed using a glass rod due to their relatively high viscosity. FTIR analysis was conducted using the Spectrum One FTIR spectrometer (PerkinElmer, Wellesley, MA, USA) equipped with a universal attenuated total reflection (ATR) accessory. ATR-FTIR spectra between 4000 and 600 cm−<sup>1</sup> were measured at a 4 cm−<sup>1</sup> resolution and averaging 16 scans per sample.

#### **3. Results and Discussion**

#### *3.1. Major Components of Biomass*

Cellulose, hemicellulose, and lignin are three main components in lignocellulosic biomass. Table 1 and Figure 1 present the FTIR band assignments and spectra of poplar and its major components isolated from the same biomass. Prior to the analysis, other extractives in the poplar sample were removed by two-step extraction: 8 h toluene/ethanol Soxhlet-extraction followed by 6 h water extraction. Isolated cellulose, hemicellulose, and lignin were analyzed using FTIR and compared with the extractives-free poplar. The assignment of each band was identified according to the previous studies [23–31].



**Figure 1.** FTIR spectra of poplar and its major components: cellulose, hemicellulose, and lignin. (Note: The assignments of the numbered bands in the figure are described in Table 1).

The IR spectra of poplar and its components showed strong O-H stretching and C-H stretching absorptions at 3367 and 2914 cm−<sup>1</sup> , respectively. These two strong absorptions are because all three major components in biomass (cellulose, hemicellulose, and lignin) have hydroxy groups and many C-H bonds in their structures. The absorption at 1745 cm−<sup>1</sup> was due to C=O stretching in hemicellulose and lignin. The absorption at 1618 cm−<sup>1</sup> represented asymmetric stretching band of the carboxyl group of glucuronic acid in hemicellulose and C=O stretching in conjugated carbonyl of lignin. The band at 1650 cm−<sup>1</sup> in the IR spectrum of cellulose was possibly caused by adsorbed H2O. Higher absorption at 3367 cm−<sup>1</sup> was also observed because of the moisture content in the biomass. In addition, the bands due to symmetric CH<sup>2</sup> bending vibration in cellulose, carboxyl vibration in glucuronic acid with xylan, and C-H in plane deformation with aromatic ring stretching in lignin were observed at 1424 cm−<sup>1</sup> . The IR absorption bands at 1582 and 1508 cm−<sup>1</sup> assigned to aromatic ring stretching and vibration (C=C-C) in lignin. The band at 1457 cm−<sup>1</sup> was observed in lignin due to its C-H deformation in methyl and methylene. The C-H bending in cellulose, hemicellulose, and lignin (aliphatic C-H stretching in methyl and phenolic alcohol) was observed at 1370 cm−<sup>1</sup> . The CH<sup>2</sup> wagging in cellulose and hemicellulose and the C-O stretching of C<sup>5</sup> substituted aromatic units, such as syringyl and condensed guaiacyl units, were assigned at 1317 cm−<sup>1</sup> . Similarly, the C-O stretching of guaiacyl unit in lignin was assigned at 1235 cm−<sup>1</sup> . The bands at 1160 and 896 cm−<sup>1</sup> arise from C-O-C stretching at the β-(1→4)-glycosidic linkages in cellulose and hemicellulose. The absorption at 1108 cm−<sup>1</sup> was associated with aromatic C-H in plane deformation for the syringyl unit. The band at 1053 cm−<sup>1</sup> was assigned to the C-OH stretching vibration of cellulose and hemicellulose. Moreover, this band was for C-O deformation in secondary alcohols and aliphatic ethers. The C-O stretching of cellulose and primary alcohols and C-H in plane deformation for guaiacyl unit exhibited at 1032 cm−<sup>1</sup> . Aromatic C-H out of plane bending in lignin was presented at 846 cm−<sup>1</sup> . Although several FTIR bands of different biomass components were overlapped, the IR spectra of samples still provide important clues, including changes of chemical composition, functionalization, and other transformation of biomass for understanding the applied biomass processing.

#### *3.2. Commonly Used Pretreatment and Preprocessing Solvents*

Table 2 and Figure S1 show the band assignment for common biomass processing solvents. Water is the most common solution in biomass analysis and the conversion processes. It also exists

in the air, and a certain amount can be accumulated in biomass during its storing and processing. The existence of water in biomass remarkably increased the bands at 3354 and 1653 cm−<sup>1</sup> because of its O-H stretching and O-H-O scissors bending, respectively [32]. Acetone, ethanol, and methanol are common organic solvents for the diverse pre- and post-processing of biomass, and they are also produced from biomass [33]. Acetone contamination on poplar was observed at 3005, 2908, 1713, 1431, 1364, and 1222 cm−<sup>1</sup> representing its CH<sup>3</sup> degenerated stretching, CH<sup>3</sup> symmetrical stretching, C=O stretching, CH<sup>3</sup> degenerated deformation, CH<sup>3</sup> symmetrical deforming, and C-C stretching, respectively [34]. A decrease in the bands at 3354 and 1653 cm−<sup>1</sup> is possibly due to the displacement of water in biomass by acetone. The spectra of ethanol impurity were shown at 3350, 2980, and 1056 cm−<sup>1</sup> for O-H stretching, C-H stretching, and C-O stretching, and those of methanol were at 3352, 2952, 2879, 1465, 1450, 1336, 1053 and 1026 cm−<sup>1</sup> for O-H stretching, C-H stretching (asymmetric), C-H stretching (symmetric), C-H bending (asymmetric), C-H bending (symmetric), O-H bending, CH<sup>3</sup> rocking, and C-O stretching, respectively [34–36]. Besides these chemicals, tetrahydrofuran (THF), dioxane, toluene, glycerol, pyridine, and chloroform are well-known solvents for diverse biomass pretreatment, isolation/purification, and analyses [1,3,19,37]. In addition, some chemicals, such as toluene, can be produced from biomass components [38]. The assignments of these impurities were assigned according to the previous studies. Contamination of poplar by THF appeared at 2977 and 2875 cm−<sup>1</sup> for its C-H stretching, 1063 cm−<sup>1</sup> for ring deformation, and 912 cm−<sup>1</sup> for CH<sup>2</sup> twisting [39]. The bands of dioxane were observed at 2960, 2890, 1457, 1322, 1255, 1119, 1057, 889 and 872 cm−<sup>1</sup> to show its equatorial (higher frequency) C-H stretching, axial (lower frequency) C-H stretching, symmetric CH<sup>2</sup> deformation, CH<sup>2</sup> wagging, CH<sup>2</sup> twisting, C-O-C symmetric stretching, ring trigonal deformation, C-C stretching, and C-O-C stretching, respectively [40]. The addition of toluene on poplar caused three bands including 3069 cm−<sup>1</sup> for C-H stretching, 1497 cm−<sup>1</sup> for C-C stretching, and 728 cm−<sup>1</sup> for C-H out of plane bending [41]. Glycerol on poplar had the bands for O-H stretching at 3341 cm−<sup>1</sup> , C-H stretching at 2948 and 2897 cm−<sup>1</sup> , C-H deformation of secondary alcohol at 1333 and 1239 cm−<sup>1</sup> , C-O stretching of primary alcohol at 1034 cm−<sup>1</sup> , and O-H bending at 923 cm−<sup>1</sup> [42,43]. Chloroform contaminants also showed at 1220 and 755 cm−<sup>1</sup> for C-H bending and CCl<sup>3</sup> stretching [34]. Pyridine contamination resulted in additional bands for C-H stretching at 3036 cm−<sup>1</sup> , C-C bonding at 1583 cm−<sup>1</sup> , C-N stretching at1485 cm−<sup>1</sup> , C-H in plane wagging at 1438 cm−<sup>1</sup> , symmetric C-H wagging at 1203 cm−<sup>1</sup> , C-H wagging at 1069 cm−<sup>1</sup> , C-C in plane wagging at 1032 cm−<sup>1</sup> , C-H out of plane bending at 750 and 693 cm−<sup>1</sup> [44,45]. The intensities of the bands at 3353 and 1653 cm−<sup>1</sup> decreased with the contaminants that do not contain OH groups such as acetone, THF, dioxane, toluene, chloroform, and pyridine due to the displacement of moisture in biomass by these solvents. On the other hand, the intensity increased with the contaminants having OH groups such as water, ethanol, methanol, and glycerol.


**Table 2.** FTIR band assignments of common biomass processing solvents on poplar [32–45].


**Table 2.** *Cont*.

#### *3.3. Acids and Alkalis*

Sulfuric acid, hydrochloric acid, acetic acid, phosphoric acid, ammonium hydroxide, and sodium hydroxide on the poplar sample were observed. As Table 3 and Figure S2 present, most acids on poplar, including sulfuric acid, hydrochloric acid, and phosphoric acid, commonly had the bands at 3370 and 1660 cm−<sup>1</sup> to represent O-H bonding and O-H-O scissors bending, respectively, because of water content. The contamination bands from sulfuric acid in the literature at 1362 and 750 cm−<sup>1</sup> for S=O (1362 cm−<sup>1</sup> ) and S-O stretching (750 cm−<sup>1</sup> ) were not clearly appeared in this study [46]. A relatively low concentration of sulfuric acid (4%) could be the reason for weak intensities of the contaminant. Hydrochloric acid showed H-Cl stretching at 2942 cm−<sup>1</sup> , while phosphoric acid had a P-OH bond and P=O stretching at 2904 and 1161 cm−<sup>1</sup> , respectively [34,47,48]. Acetic acid bands appeared at 3351, 2916, 1706, 1427, 1234, and 1031 cm−<sup>1</sup> to indicate its O-H stretching, symmetric CH<sup>3</sup> stretching, C=O stretching, CH<sup>3</sup> deformation, O-H bending and CH<sup>3</sup> rocking, respectively [34]. Sodium hydroxide had the bands caused by water at 3360 and 1660 cm−<sup>1</sup> , but there were no other clear contamination

bands observed. Similarly, ammonium hydroxide had the IR bands at 3350 and 1660 cm−<sup>1</sup> from both water and NH<sup>3</sup> content but N-H stretching of NH<sup>4</sup> <sup>+</sup> also appeared at 2914 cm−<sup>1</sup> . Previous study also said that adsorption of ammonia increased the overall polarity and resulted in the absorbance of several bands (e.g., 1115 and 1036 cm−<sup>1</sup> in this study) not from the N-H vibrations [49].


**Table 3.** FTIR band assignments of acids and alkalis contaminants on poplar [34,46–49].

#### *3.4. Ionic Liquids*

Besides the aforementioned chemicals, FTIR spectra and the band assignments of ionic liquids, enzymes, and biomass-derived chemicals on poplar are presented in Table 4 and Figures S3–S5. The bands from 1-butyl-3-methylimidazolium chloride contaminant were observed at 3341, 1658, and 1604 cm−<sup>1</sup> representing the formation of quaternary amine salt formation with chlorine, C=C stretching, and C=N stretching, respectively. However, the band at 835 cm−<sup>1</sup> representing C-N stretching vibration was not clearly observed [50]. The bands from 1-benzyl-3-methylimidazolium chloride were 2961, 1574, 765, and 633 cm−<sup>1</sup> from C-H stretching, C-C stretching of ring vibration, and C-N/C-Cl in-plane bending, respectively [51]. Moreover, two bands at 1383 and 1176 cm−<sup>1</sup> were observed; however, further study is needed to identify them.

**Table 4.** FTIR band assignments of ionic liquids, enzymes, and biomass-derived chemicals on poplar [49–65].



**Table 4.** *Cont*.

The bands of choline chloride-urea, which is a well-known DES, were at 3435 and 3340 cm−<sup>1</sup> , which ascribed to the stretching of –NH<sup>2</sup> (asymmetric and symmetric), 1669 cm−<sup>1</sup> for the bending vibration of –NH2, 1597 cm−<sup>1</sup> for bending vibration of -OH possibly due to the existence of water, 1474 cm−<sup>1</sup> for CH<sup>3</sup> rocking, 1152 cm−<sup>1</sup> for asymmetric C-N stretching, 1062 cm−<sup>1</sup> for CH<sup>2</sup> rocking, 961 cm−<sup>1</sup> for asymmetric stretching of CCO from choline structure and 790 cm−<sup>1</sup> from C=O bonding [52,53]. Three lignin-based DESs, choline chloride–*p*-hydroxybenzoic acid (PHA), choline chloride–4-hydroxybenzaldehyde (PB), and choline chloride–*p*-coumaric acid (PCA), were mixed with poplar sample to observe the possible contamination bands. The bands of choline chloride–PHA were observed at 3180 cm−<sup>1</sup> for O-H stretching, 1681 cm−<sup>1</sup> for C=O stretching, 1581 cm−<sup>1</sup> for the asymmetric stretch of COO vibrations, 1282 cm−<sup>1</sup> for C-O stretching vibration, 1082 cm−<sup>1</sup> for C-O stretching, 953 cm−<sup>1</sup> for C-N stretching, 861 cm−<sup>1</sup> for CH<sup>2</sup> rocking vibrations, 838 cm−<sup>1</sup> for aromatic C-H out-of-plane bending, 786 cm−<sup>1</sup> for C–C stretching [54,55]. The bands from choline chloride–PB were

observed at 3122 cm−<sup>1</sup> for the stretching vibration of the phenolic O-H group exhibiting intermolecular hydrogen bonding, 1667 cm−<sup>1</sup> for the stretching vibration of carbonyl group, 1272 cm−<sup>1</sup> for the methylene, 1030 cm−<sup>1</sup> for C-H binding vibration [56]. The bands from choline chloride—PCA DES were observed at 3126 cm−<sup>1</sup> , 1675 cm−<sup>1</sup> , 1606 cm−<sup>1</sup> , 1160 cm−<sup>1</sup> , 848 cm−<sup>1</sup> from bending vibration of –NH2, C=O stretch of carboxylic acid, C=C stretching, C-OH stretching, C-H stretching and 771cm−<sup>1</sup> from stretching of the -OH group on the second carbon of the choline chloride [57–59].

#### *3.5. Enzymes*

Enzymes such as cellulase and β-glucosidase break polysaccharides in biomass to fermentable sugars. The bands at 3353, 2942, 2900, 1642, 1334, and 1036 cm−<sup>1</sup> were observed from cellulase (Table 4 and Figure S4). The bands at 3353, 2942, and 2900 cm−<sup>1</sup> were from N-H/O-H stretching and the C-H stretching (asymmetric and symmetric) of cellulase. The bands at 1642, 1432, 1334, and 1036 cm−<sup>1</sup> were possibly from NH<sup>2</sup> scissoring, C-C stretching, C-N stretching, and C-O stretching, respectively [60–62]. β-glucosidase also showed similar bands at 3351, 1646, 1432, and 620 cm−<sup>1</sup> , which represented N-H stretching, N-H bonding and C=O stretching, N-H bending, and N-H out of plane bending, respectively [63].

#### *3.6. Biomass-Derived Chemicals*

Biomass can be contaminated by its decomposed fractions. For instance, furan-based chemicals such as furfural and hydroxymethylfurfural can be produced through the dehydration of hexoses and pentoses in biomass. As Figure S5 presents, HMF contamination showed at 3364, 1661, and 1561 cm−<sup>1</sup> from O-H stretching, C=O stretching (carbonyl), and C=C stretching of furan ring, respectively [64]. Furfural also showed bands at 3134 cm−<sup>1</sup> from C-H stretching of furan ring, at 2859 cm−<sup>1</sup> from the C-H vibration of aldehyde group, 1671 cm−<sup>1</sup> from C=O in the conjugated carbonyl group, 1465 cm−<sup>1</sup> from C=C stretching of furan ring, 1276 and 1021 cm−<sup>1</sup> from C-O stretching vibration, 928, 884, and 755 cm−<sup>1</sup> from C-H bending out of plane peaks [65,66].

#### **4. Conclusions**

The identification of contaminants on the biomass surface after preprocessing is important to avoid the unwanted misleading of analysis data. This study investigated and discussed diagnostic FTIR bands from 26 potential chemicals, including organic solvents, acids and alkalis, ionic liquids, enzymes, and biomass-derived components through diverse biomass preprocessing. The observation of these contaminants will improve the FTIR analysis with diverse biomass and bioproducts in the biorefinery.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2076-3417/10/12/4345/s1, Figure S1: FTIR spectra of preprocessing solvent contaminants on poplar, Figure S2: FTIR spectra of preprocessing acid and alkaline contaminants on poplar, Figure S3: FTIR spectra of ionic liquid contaminants on poplar, Figure S4: FTIR spectra of enzyme contaminants on poplar, Figure S5: FTIR spectra of biomass-derived chemical contaminants on poplar.

**Author Contributions:** C.G.Y. and A.J.R. conceived and designed the research. J.Z., C.G.Y., M.L., and Y.P. carried out the experiment. J.Z., M.L., and C.G.Y. wrote the manuscript. All the authors discussed data and revised the paper. All authors have given approval to the final version of the manuscript.

**Funding:** This research received no external funding.

**Acknowledgments:** This manuscript has been authored by UT-Battelle, LLC under Contract No. DE-AC05- 00OR22725 with the U.S. Department of Energy. This study was supported and performed as part of the BioEnergy Science Center (BESC) and Center for Bioenergy Innovation (CBI). The BESC and CBI are U.S. Department of Energy Bioenergy Research Centers supported by the Office of Biological and Environmental Research in the DOE Office of Science. The United States Government retains and the publisher, by accepting the article for publication, acknowledges that the United States Government retains a non-exclusive, paid-up, irrevocable, world-wide license to publish or reproduce the published form of this manuscript, or allow others to do so, for the United States Government purposes. The Department of Energy will provide public access to these results of federally sponsored research in accordance with the DOE Public Access Plan (http://energy.gov/downloads/doe-public-access-plan). The views and opinions of the authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof. Neither the United States Government nor any agency thereof, nor any of

their employees, makes any warranty, expressed or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights. The views and opinions of the authors expressed herein do not necessarily state or reflect those of the United States Government or any agency thereof. Neither the United States Government nor any agency thereof, nor any of their employees, makes any warranty, expressed or implied, or assumes any legal liability or responsibility for the accuracy, completeness, or usefulness of any information, apparatus, product, or process disclosed, or represents that its use would not infringe privately owned rights.

**Conflicts of Interest:** There are no conflicts to declare.

### **References**


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### *Article* **Drying E**ff**ect on Enzymatic Hydrolysis of Cellulose Associated with Porosity and Crystallinity**

**Bonwook Koo 1,\*, Jaemin Jo <sup>1</sup> and Seong-Min Cho <sup>2</sup>**


Received: 27 July 2020; Accepted: 7 August 2020; Published: 11 August 2020

**Featured Application: Drying causes irreversible structural changes in cellulose and the changes are intimately associated with porosity, including pore volume and surface area. These changes must be considered for the application of cellulose in high value industry such as sustainable polymers that use cellulose nanofiber and sustainable sugar production.**

**Abstract:** The effect of drying on the enzymatic hydrolysis of cellulose was determined by analysis of porosity and crystallinity. Fiber hornification induced by drying produced an irreversible reduction in pore volume due to shrinkage and pore collapse, and the decrease in porosity inhibited enzymatic hydrolysis. The drying effect index (DEI) was defined as the difference in enzymatic digestibility between oven- and never-dried pulp, and it was determined that more enzymes caused a higher DEI at the initial stage of enzymatic hydrolysis and the highest DEI was also observed at the earlier stages with higher enzyme dosage. However, there was no significant difference in the DEI with less enzymes because cellulose conversion to sugars during hydrolysis did not enhance enzymatic hydrolysis due to the decrease in enzyme activity. The water retention value (WRV) and Simons' staining were used to measure pore volume and to investigate the cause of the decrease in enzymatic hydrolysis. A decrease in enzyme accessibility induced by the collapse of enzymes' accessible larger pores was determined and this decreased the enzymatic hydrolysis. However, drying once did not cause any irreversible change in the crystalline structure, thus it seems there is no correlation between enzymatic digestibility and crystalline structure.

**Keywords:** drying effect; cellulose; enzymatic hydrolysis; hornification; porosity

#### **1. Introduction**

Biorefinery platforms using biomass have been studied widely in recent years because of their low carbon footprint [1]. The U.S. Department of Energy (DOE) announced the top 12 platform chemicals that can be produced from biomass and that it would consider the market and potential application of these in industry [2,3]. Most of the platform chemicals can be produced from sugars, which indicates that low cost sugar production from biomass is important to achieve a low carbon footprint [4]. Lignocellulosic biomass has been suggested as the feedstock for sustainable sugar production and various conversion processes including pretreatment and saccharification have been studied to facilitate effective sugar production from lignocellulosic biomass [5–7].

Sustainable sugar is mainly produced by enzymatic saccharification and pretreatment must be performed to improve enzyme accessibility to cellulose [8,9]. The pretreatment exposes cellulose by partial removal of hemicellulose and lignin, and it facilitates enzyme access to cellulose. After pretreatment, the exposed cellulose is likely to be partially dried and the cellulose drying affects its enzymatic digestibility [10]. Once cellulose has been dried, the dimensions of the cellulose are changed due to the collapse of pores and shrinkage of the internal volume [11,12]. The structural change stiffens the cellulose via a process known as "hornification" [13].

The process of hornification is frequently explained as irreversible, or partially irreversible hydrogen bonding upon drying or water removal [14] due to the aggregation of the cellulose chains [15]. The cellulose chains are aggregated due to the removal of water by heating and then they are not able to fully open up to the next exposure to water [16]. The aggregation may collapse the pore structure, and thus a decrease in pore volume and surface area can be expected [12,17]. Crystallinity in cellulose fiber has been considered as one of the major properties altered by the aggregation [18]. It was reported that several cycles of drying and rewetting caused irreversible change in crystalline structure, which could not be recovered due to the growth of crystalline domains through co-crystallization [19]. However, no change in crystalline index by the recycling of paper has been reported [20].

It has been reported that pore volume is significantly decreased by the process of repetitive drying and rewetting. Repetitive recycling of delignified and alkali-extracted pulp was carried out by rewetting and drying, and it decreased the Brunauer–Emmett–Teller (BET) surface area and pore volume of cellulose, as measured by the water retention value (WRV). In a five-times recycling procedure, the BET surface area and the WRV decreased by 22.9% and 35.9%, respectively, compared with non-dried and rewet pulp [17].

In addition, in previous studies on the effect of drying, it was reported that hornification induced by drying affects enzymatic hydrolysis [21,22]. The enzymatic digestibility of dried substrate was significantly lower than that of never-dried substrate, and the reduction in enzymatic conversion was caused by a decrease in pore volume, which can be evaluated by WRV [22]. Hornification induced by drying caused the collapse of larger pores while increasing the number of smaller pores, which are not accessible to enzymes [21]. This suggests that the collapse or closure of larger pores could be a primary reason for a reduction in enzyme accessibility to cellulose, given the size of the cellulase core.

Although numerous studies have been performed to investigate the drying effect on enzymatic hydrolysis [23], little research on the effect with regard to enzyme dosage and the kinetics of hydrolysis has been performed. This study aims to investigate the drying effect on enzymatic hydrolysis at different enzyme dosages to determine the hydrolysis kinetics and the limitation of enzymatic digestibility. In addition, several properties of the cellulose structure including porosity, enzyme accessible surface area, and crystallinity were determined to investigate their effect on enzymatic hydrolysis.

#### **2. Materials and Methods**

#### *2.1. Sample Preparation*

A fully bleached and never-dried hardwood pulp was obtained from a mill in the southeast of the United States and used in this study. The pulp was washed thoroughly with plenty of tap water. Handsheets were made using the washed pulp to prevent the formation of fiber flocks, which might affect the enzymatic digestibility of pulp due to their effect on the enzyme-accessible surface area. It has been reported that fiber bundles in a wet state can easily form very strong fiber flocks of varying size upon drying due to inter- and intra-fiber hydrogen bonding [22]. The washed hardwood pulp, which weighed about 2.0 g was disintegrated in a standard disintegrator (Lorentzen & Wettre, Kista, Sweden) for 5 min with 5% solid consistency at room temperature and then diluted to 1% solid consistency with deionized water. The diluted pulp suspension was used to make handsheets according to the TAPPI Standard method, T205 sp-95 [24] using a standard laboratory handsheet mold. The weight was 2.0 g per sheet (based on the oven-dried weight) and the handsheets were stored in a cold room at 4 ◦C using a plastic bag to prevent biological decomposition and drying.

#### *2.2. Compositional Analysis*

The compositional analysis of the pulp was performed according to NREL (National Renewable Energy Laboratory) Standard Procedures [25]. Sulfuric acid hydrolysis w performed in two stages with 72% and 4% of sulfuric acid. First, the pulp was hydrolyzed with 72% of sulfuric acid at 30 ◦C for 1 hr and then the acid was diluted to 4% for the second hydrolysis at 121 ◦C for 1 hr. After two stages of sulfuric acid hydrolysis, the hydrolysate from the pulp was filtrated by a 0.2 µm Milipore filter and analyzed by high-performance liquid chromatography (Agilent 1200; Agilent Technologies, Palo Alto, CA, USA) to measure the amount of structural sugars. The sugars were separated with a Shodex SP0810 column at a temperature of 80 ◦C with a flow rate of 0.5 mL/min using deionized water as an eluent. A refractive index detector was used to quantify the arabinose, galactose, glucose, xylose, and mannose in the hydrolysates. The chemical composition of the pulp is shown in Table 1.


**Table 1.** Chemical composition of the bleached hardwood pulp.

All values were calculated based on the oven-dried weight of pulp; Glu: glucan; Xyl: xylan; Man: mannan; Gal: galactan; Ara: arabinan; KL: Klason lignin; ASL: acid soluble lignin; ND: not detected.

#### *2.3. Drying and Rewetting of the Pulp*

The handsheets were subject to drying and rewet to induce fiber hornification. Two different drying methods, oven and freeze drying, were applied to determine the drying effect on enzymatic hydrolysis. Oven drying of the handsheets was conducted in a drying oven at 105 ◦C for 24 h and freeze drying was conducted at −40 ◦C in a freeze dryer for 48 h. After each drying, the dried handsheets were stored in plastic bags for enzymatic hydrolysis and analysis of those properties. Rewetting the dried pulp was performed by disintegration for 5 min at 5% of consistency and then the water was drained out by centrifuge. Rewetted pulp fibers were also stored in a plastic bag and used for enzymatic hydrolysis and analysis of those properties.

#### *2.4. Enzymatic Hydrolysis*

Commercial cellulase including β-glucosidase (C-tec2) and a xylanase (H-tec2) provided by Novozymes (Franklinton, NC, USA), were used for enzymatic hydrolysis. Cellulase dosages of 4, 10 and 20 FPU per g of pulp (5.2, 13.3 and 26.7 enzyme protein mg per g of pulp) and a ninth of the xylanase were used to determine the drying effect depending on enzyme dosage. Enzymatic hydrolysis was performed in 20 mL of 100 mM acetate buffer (pH 5.0) at a 5% (*w*/*v*) solids loading. The substrates and enzymes were incubated in a shaker at 50 ◦C and 180 rpm for 120 h. The enzymatic digestibility was determined in duplicate by the amount of sugars released during enzymatic hydrolysis based on the amount of structural sugars in the original pulp. The amount of released sugars in the enzymatic hydrolysate was quantified by the HPLC (high pressure liquid chromatography) [26].

#### *2.5. Analysis of Pulp Properties*

#### 2.5.1. Porosity by Water Retention Value and Simons' Staining Method

When pulp fibers are dried, internal fiber volume shrinks [11] and the shrinkage prevents the accurate measurement of porosity due to the pore collapse caused by drying. Thus, the measurement needs to be done in a wet state [27] and two independent methods, water retention value (WRV) and Simon's staining method were selected to evaluate the porosity of pulp in wet state.

The WRV has been used to estimate the fiber saturation point, which correlates to the amount of water in the cell wall pores or the volume of pores, and it has been found that the drying effect induced by fiber hornification is reflected in a reduction in the WRV [22]. The WRV was determined using the TAPPI Useful method with a centrifugal force (Eppendorf North America, Hauppauge, NY, USA) of 900 rcf (2400 rpm) for 30 min [28]. The centrifuged wet pulp was first weighed and then oven dried at 105 ◦C overnight. The WRV was calculated by the percentage of water retained in the dried pulp, i.e.,

$$\text{WRV (\%)} = [\text{(W}\_{\text{wet}}\text{-W}\_{\text{drried}}) / \text{W}\_{\text{dried}}] \, ^\ast \, 100 \tag{1}$$

in which:


In addition to the WRV, the Simon's staining method was performed to analyze the porosity, including the enzyme accessible surface area of the pulp, depending on the drying method. For the Simons' staining method, the 1:1 staining method were used in this study [29]. Blue dye (DB) and orange dye (DO) were dissolved in nanopure water for the preparation of each dye solution and the final concentrations of DB and DO were 1% (*w*/*v*), respectively. For the orange dye, only higher molecular fractions were used after the fractionation. The dye solution was then prepared with 1 mL of the DB solution, 1 mL of the DO solution, 10 mL of 10% (*w*/*v*) NaCl and 70 mL of nanopure water. For staining, a 25 mg of sample (dry weight) in the dye solution was incubated in a 75 ◦C shaking incubator at 200 rpm for 48 h. The stained samples were then filtered, rinsed with a minimum amount of distilled water and stripped with 25% (*w*/*v*) aqueous pyridine at 45 ◦C for 18 h. The dye stripping solution was then analyzed using a UV-Vis spectrophotometer to determine the concentration of DO and DB (the maximum absorbance of DB and DO was at 624 and 455 nm, respectively). The concentration of DO and DB dyes in the dye stripping solution (C<sup>O</sup> and CB) was determined using the following two equations, Equations (2) and (3) (using the Lambert–Beer law for a binary mixture), which were solved simultaneously [30].

$$A\_{455nm} = \varepsilon\_{\text{O}\text{455}} \, L\text{C}\_{\text{O}} + \varepsilon\_{\text{B}\text{455}} \, L\text{C}\_{\text{B}} \tag{2}$$

$$A\_{624nm} = \varepsilon\_{\text{O}\nmid624} \, \text{LC}\_{\text{O}} + \varepsilon\_{\text{B}\nmid624} \, \text{LC}\_{\text{B}} \tag{3}$$

in which:


• ε*O*/*<sup>455</sup>* = 35.62, ε*B*/*<sup>455</sup>* = 2.59, ε*O*/*<sup>624</sup>* = 0.19, ε*B*/*<sup>624</sup>* = 15.62 L·g −1 ·cm−<sup>1</sup> and L = 1 cm

#### 2.5.2. Crystallinity by <sup>13</sup>C CPMAS Solid-State NMR Analysis

High-resolution <sup>13</sup>C CPMAS NMR spectra were collected by a Bruker Avance 200 MHz spectrometer (Bruker BioSpin Corporation, Billerica, MA, USA) with a 7 mm probe, operating at 50.13 MHz for <sup>13</sup>C, at room temperature. The spinning speed was 7000 Hz, contact pulse 2 ms, acquisition time 51.3 ms and delay between pulses was 4 s, with 20,000 scans. The adamantine peak was used as an external reference (δC 38.3 ppm). The samples were hydrated with deionized water (ca. 43%) before recording the spectra. The <sup>13</sup>C chemical shifts were given in δ values (ppm) and each peak was assigned according to the values in the literature [19]: C6-amorphous (61.3 to 62.1 ppm), C6-crystalline (65.2 to 65.8 ppm), C2, C3 and C5 (71.2 to 75.6 ppm), C4-amorphous (83.5 to 84.4 ppm), C4-crystalline (88.2 to 89.6 ppm), and C1 (104.2 to 107.8 ppm).

#### **3. Results and Discussion**

#### *3.1. Drygin E*ff*ect on Enzymatic Hydrolysis*

The effect of drying on enzymatic hydrolysis was evaluated by enzymatic digestibility, which was determined by the amount of sugars released during enzymatic hydrolysis with 10 FPU/g-pulp of cellulase. It was calculated based on the amount of structural sugars in the original pulp.

Compared with oven-dried (OD) pulp, the enzymatic digestibility of never-dried (ND) pulp increased rapidly in the initial stage of the hydrolysis, but the hydrolysis rate of the ND pulp reduced gradually as the hydrolysis proceeded (Figure 1b). A decrease in hydrolysis rate has been ascribed to several factors such as the transformation of cellulose into a structurally resistant form, inhibition of enzyme action by accumulated products, and the completeness of hydrolysis [31]. Here, it seems that the decrease in hydrolysis rate of the ND pulp can be attributed to the completeness of the hydrolysis because the digestibility leveled off at 90%. Though the enzymatic digestibility of freeze-dried (FD) pulp was slightly lower than that of the ND pulp, a similar trend was observed in the hydrolysis of the FD pulp. This suggested that freeze drying caused mild hornification, unlike oven drying [21].

**Figure 1.** Enzymatic digestibility (sugar recovery) depending on the enzyme dosage of (**a**) 4 FPU/g biomass, (**b**) 10 FPU/g biomass, and (**c**) 20 FPU/g biomass.

Enzymatic hydrolysis of the ND pulp showed the highest digestibility (96.6%) when the hydrolysis was carried out with 10 FPU of cellulase for 120 h. However, the OD pulp showed significantly low digestibility though that of the FD pulp was almost similar to that of the ND pulp (96.1%) (Figure 1b). Pore volume has been considered as one of the most important properties affecting enzymatic hydrolysis [32,33] and fiber hornification induced by drying produces irreversible reduction in pore volume due to shrinkage and collapse of pores [22,32]. The reduction in pore volume decreases enzyme accessibility [22] and it reduces enzymatic digestibility due to less enzyme accessible surface area [32]. Therefore, the difference in enzymatic digestibility depending on drying, as shown in Figure 1b, should be caused by the reduction in pore volume induced by drying. However, there was a significant difference in the enzymatic digestibility of the FD and OD pulp. Again, it was considered that the better hydrolyzability of the FD pulp compared to the OD pulp, was due to the milder hornification induced by freeze drying [21]. The mild hornification can be explained by the rigidity of fibers at initial freezing [34]. The freezing gave a sort of rigidity to the fibers and this maintained the pore structure, thus minimizing pore collapse during drying. Therefore, enzymatic hydrolysis of the FD pulp showed better hydrolyzability compared to the OD pulp.

The difference in digestibility between the ND and OD pulp decreased gradually as the hydrolysis progressed because the digestibility of the OD pulp kept increasing while that of the ND pulp had already leveled off. The OD pulp had less pore volume due to fiber hornification, and thus there was a big difference in enzymatic digestibility compared to the ND pulp in the initial stages [17]. However new pores were formed by cellulose conversion to sugars during hydrolysis and this can give enzyme the space to access remaining cellulose [35]. Thus, the difference in digestibility between the ND and OD pulps was reduced gradually as the hydrolysis progressed. It is considered that the lower enzymatic digestibility of the OD pulp at the initial stage was induced by pore collapse; however, it can be recovered at a later stage because the pore volume increased due to cellulose conversion to sugars during enzymatic hydrolysis, even though hornification leads to irreversible or partial irreversible collapse of pore and it causes the difference in digestibility at the final stage of hydrolysis [32]. In order to determine the kinetics of enzymatic hydrolysis, the Langmuir equation style fitting was used, which

enabled the prediction of the final digestibility after 120 h of enzymatic hydrolysis. The Langmuir equation style for the determination is as follows:

$$S\_t = (S\_{max} \cdot t) / (k + t) \tag{4}$$

where *S<sup>t</sup>* is enzymatic digestibility at a time, *Smax* is the maximum digestibility predicted (%), *k* is the Langmuir style equilibrium constant and *t* is time (h). This equation can be re-organized by the Lineweaver–Burk method to linearize the rate of expression as follows:

$$\mathbf{1}/\mathbf{S}\_{t} = k \cdot (\mathbf{1}/\mathbf{S}\_{\max}) \cdot (\mathbf{1}/t) + \mathbf{1}/\mathbf{S}\_{\max} \tag{5}$$

The kinetic parameters were inferred with an intercept at 1/*Smax* and a slope of *k*·(1/*Smax*) and are summarized in Table 2. The predicted final digestibility of the OD pulp with 10 FPU was 82.6%, which was less than that of the ND pulp (102.0%) (Table 2). It was proved that there was a big difference between the ND and OD pulps with regard to enzyme accessibility. The difference was caused by the irreversible transformation of cellulose into a structurally resistant form by hornification and the inhibition of enzyme action by accumulated products [31]. All of the predicted digestibilities depending on enzyme dosage and drying, were similar to the real values.

**Table 2.** The prediction of final enzymatic digestibility depending on enzyme dosage and drying.


#### *3.2. Drying E*ff*ect Depending on Enzyme Dosage*

The drying effect was determined depending on enzyme dosage. Enzymatic hydrolysis was performed with 4 FPU and 20 FPU of cellulase and the enzymatic digestibility after 120 h was predicted using the same method as above with 10 FPU (Table 2).

The predicted enzymatic digestibility after 120 h increased as the enzyme dosage increased, regardless of drying. The increase in the rate of digestibility was more remarkable in the hydrolysis of the OD pulp, and the rate increased by 20.0 (ND), 24.1 (FD) and 32.3% (OD) when the enzyme dosage increased from 4 to 20 FPU. It is believed that a higher enzyme dosage can convert more cellulose to sugars in a short period of enzymatic hydrolysis, and thus the digestibility of the OD pulp increases significantly as the enzyme dosage is increased. There was a 26.0% increase in the digestibility of the OD pulp when the enzyme dosage was increased from 4 to 10 FPU, which was much higher than the 5.1% increase in digestibility when the enzymes were increased from 10 to 20 FPU. This suggested that an enzyme dosage 10 FPU was enough to improve enzymatic digestibility by pore formation during hydrolysis.

The kinetics of enzymatic hydrolysis with 4 FPU and 20 FPU are shown in Figure 1a,c. There was no leveling off on the enzymatic hydrolysis with 4 FPU. Enzymatic hydrolysis with lower enzyme dosages requires longer hydrolysis time to obtain maximum enzymatic digestibility [36], and 120 h of hydrolysis time might not be enough to achieve maximum digestibility with 4 FPU of enzyme dosage. However, the predicted digestibility with 4 FPU was 62.9%, which was slightly lower than the actual digestibility of 70.4% after 120 h (Tables 2 and 3). It was considered that hydrolysis with 4 FPU progressed steadily over 120 h and it had already reached the maximum digestibility even though it did not appear to level off. There was no further increase in digestibility with 4 FPU as a result of enzyme inhibition caused by accumulated products such as released sugars [37]. When 20 FPU was used, the enzymatic digestibility of the ND and FD pulps leveled off in 24 h and both final digestibilities were almost 100% after 120 h. Although the digestibility of the OD pulp also increased

when the enzyme dosage increased to 20 FPU, it was still lower than that of the ND and FD pulp due to the drying effect induced by fiber hornification. The enzymatic digestibility after 120 h and the predicted final digestibility of the OD pulp with 20 FPU was 93.2% and 90.1%, respectively. It indicated that the drying effect on enzymatic hydrolysis can vary depending on enzyme dosage, but there is likely to be a difference in the final enzymatic digestibility of the ND and OD pulps due to irreversible structural change in the cellulose [38]. The drying effect described by the differences in the enzymatic digestibility of the ND and OD pulp with 20 FPU was clearer at the initial stage and it was different compared to the drying effect with 4 FPU, which did not show any big differences as the enzymatic hydrolysis progressed. Thus, the drying effect can vary depending on enzyme dosage and the term "Drying effect index" (DEI) was suggested to define the drying effect. The equation for calculating the DEI is as follows (Equation (6)), and the DEI was plotted, as shown in Figure 2.

$$\text{DEI} \left( \% \right) = 100 - \text{ (Digestibility of OD pulp/Digestibility of ND pulp)} \* 100 \tag{6}$$

The highest DEI, which represents the biggest drying effect, increased as enzyme dosage increased and were revealed at the earlier stages of hydrolysis (Figure 2). The highest DEIs were 22.3 (4 FPU), 24.1 (10 FPU) and 29.3 (20 FPU) and these were observed at 96, 24 and 6 h, respectively. Enzymatic hydrolysis of the OD pulp began in the non-collapsed part first, followed by hydrolysis of the collapsed part because enzyme access to the non-collapsed part was easier than access to the collapsed part [39]. Higher enzyme dosage resulted in the biggest difference in digestibility (DEI) at the earlier stage (29.3 at 6 h) because there was a considerable difference in the amount of collapsed pores in the ND and OD pulp. The ND pulp was hydrolyzed quickly due to the small amount of collapsed pores and the fast hydrolysis of the non-collapsed pores in the ND pulp by more enzymes resulted in a big difference in the DEI in 6 h. As the hydrolysis progressed, the reversible collapsed pores were converted to partial collapsed pores by rewetting in buffer solution and the cellulose conversion to sugars induced by progress in the hydrolysis provided more pore volume in the cellulose [40]. Thus, the DEI with 20 FPU decreased as hydrolysis progressed, which meant that there was less difference in enzymatic digestibility between the ND and OD pulp. Therefore, it was concluded that more enzymes caused a higher drying effect at the initial stage of enzymatic hydrolysis and the time for the highest drying effect as revealed earlier. Once the drying effect reached maximum, the drying effect was reduced as hydrolysis progressed when 10 and 20 FPU of enzymes were used. When 4 FPU of enzymes was used, however, there was little difference in the DEI as hydrolysis progressed, unlike 10 and 20 FPU of enzymes. It was considered that the slower rate of hydrolysis with less enzymes caused more inhibition due to a decrease in enzyme activity, and thus, new pores formed during hydrolysis could not dramatically enhance enzyme accessibility in 120 h. In summary, the drying effect on enzymatic hydrolysis increased as the enzyme dosage increased, and the highest DEI occurred in the earlier stages with higher enzyme dosage. As enzymatic hydrolysis with 10 and 20 FPU of enzymes progressed, the drying effect decreased gradually due to the formation of pores in the OD pulp during hydrolysis. When 4 FPU was used, however, a decrease in the DEI was not observed in 120 h because there was not enough pore formation from enzyme inhibition. − ∗

**Figure 2.** Drying effect index depending on enzyme dosage and hydrolysis time.

#### *3.3. Evaluation of Drying E*ff*ect Using Water Retention Value and Simons' Staining Method*

The water retention value (WRV) has been used to characterize the drying effect on enzymatic hydrolysis induced by fiber hornification because it can easily measure the pore volume of pulp in a wet state by measuring the amount of water in cell wall [22].

It was previously found that the WRV of the OD pulp decreased significantly due to collapse and shrinkage of pores and the decrease in the WRV correlated well with enzymatic digestibility [22,32]. In this study, the OD pulp with lower enzymatic digestibility, as shown above, revealed a significantly lower WRV, which decreased by 36.4% and 33.3% based on the ND and FD pulps, respectively (Table 3). The decrease in WRV related to the lower enzymatic digestibility was also caused by less enzyme accessibility to cellulose, induced by the decrease in pore volume [22]. However, as enzymatic hydrolysis progresses, new pores are formed through cellulose conversion to sugars and the pore formation increases the pore volume in cellulose. Therefore, the WRV of the OD pulp hydrolyzed for 24 h increased drastically from 105.7 to 203.0%. As described, the kinetics of enzymatic hydrolysis showed that the bigger drying effect at initial stages, which is due to less pore volume caused by the collapse of pores and pores formed by cellulose conversion during hydrolysis, can supply enzymes with space to access cellulose. Therefore, the difference in enzymatic digestibility between the ND and OD pulps will decrease.



The Simons' staining method can be used to evaluate the enzyme accessible surface area on cellulose using adsorption and desorption of dyes [26,30]. The pores are segregated depending on the diameter of two different dyes, which have a different affinity for hydroxyl groups in cellulose, and thereby, enzyme accessible surface area of cellulose can be measured in a wet state. The large orange-colored dye molecules can only penetrate larger pores and adsorb in preference to the small blue-colored dye molecules as a result of their greater affinity for hydroxyl groups in cellulose [29]. The large orange-colored dye molecule is mainly composed of two fractions that have hydrodynamic diameters of 5–7 and 12–36 nm [29]. The 5–7 nm diameter is very similar to that of the catalytic core of *Trichoderma reesei* endoglucanase [21]. Therefore, enzyme accessible surface area can be evaluated by the amount of adsorbed orange dye [26], even though the adsorption behavior onto the cellulose surface may be different to enzymes, and also the drying effect on enzymatic hydrolysis has been assessed effectively by the Simons' staining [21].

As result of the Simons' staining, we found that the amount of orange dye adsorbed in the ND and OD pulps were 154.4 and 75.9 mg per g of cellulose, respectively, and the drying process decreased more than half of the accessible surface area in larger pores by pore collapse (Table 4). However, the amount of blue dye, which adsorbed on the surface area of smaller pores, increased from 36.7 to 62.1 mg per g of cellulose. The increase in the adsorption of blue dye was likely caused by the change of larger pores to smaller pores due to the collapse of larger pores, and thus, the surface area of smaller pore increased [21]. The increase in enzyme accessible surface area enhanced enzymatic digestibility through better enzyme adsorption, and it was more remarkable at the initial stage. The total amount of adsorbed dye decreased from 191.0 to 138.0 by drying, which suggested that the formation of the smaller pores and disappearance of larger pores by pore collapse in cellulose decreased the total surface area. It is well known that most pretreatment processes open up the cell wall structure by mechanical shearing force and/or chemical removal of components and this forms new pores in cellulose [41]. Thus, enzymatic hydrolysis could be enhanced by increasing the accessibility through

pore formation. The drying process decreased enzyme accessibility and reduced enzymatic hydrolysis. This can be explained by using the opposing mechanism of pretreatment on enzyme accessibility and enzymatic hydrolysis. The drying effect was more remarkable at the initial stage and then it decreased as hydrolysis progressed. However, the irreversible change in structure made a difference to the maximum enzymatic digestibility.


**Table 4.** Amount of adsorbed dye depending on drying methods.

#### *3.4. Evaluation of Drying E*ff*ect on Crystalline Structure Using <sup>13</sup>C CPMAS Solid-State NMR Analysis* ± ± 2 ± 4 ± 3 ± 5 ± 8

The ultrastructure of cellulose has mainly been determined by <sup>13</sup>C CPMAS solid-state NMR spectroscopy [42,43], and the solid-state NMR characterized changes associated with drying pulp [44] and bleached pulp by hornification has also been reported [19].

The <sup>13</sup>C solid-state NMR analysis was performed in wet conditions for the ND, OD and FD pulps and the spectra with signals assigned are shown in Figure 3 [19]. The <sup>13</sup>C NMR spectrum of the pulp was mainly dominated by signals assigned to cellulose. Signals assigned to C-4 and C-6 were partially separated into two clusters, labeled i and s, which were assigned to the interior and surface of the crystalline domains, respectively. Broadening of the NMR signal and relative sharpening of C-4 and C-6 signals on the surfaces of crystalline domains by partial drying has been reported, but the signal broadening and sharpening of C-4 and C-6 signals can be reversed by rewetting the pulp [19]. The <sup>13</sup>C NMR spectra of all three pulps were quite similar because both the FD and OD pulps were analyzed in a wet state. Although it has been reported that several repetitions of drying and rewetting cause the difference to peak at 89.4 ppm due to the irreversible drying effect [19], it was not observed in this study because drying was performed only once and rewetting for NMR analysis might reverse the drying effect. Therefore, it can be concluded that drying once did not cause an irreversible change in crystalline structure, and therefore, the decrease in enzymatic digestibility does not seem to be related to the structure of cellulose. However, it might be related to cellulose structure if the pulp was dried more than once.

**Figure 3.** 13C solid-state NMR spectra of never-dried, freeze-dried and oven-dried pulps.

### **4. Conclusions**

The drying effect was evaluated by the difference in enzymatic digestibility in the ND and OD pulp, which was found to vary depending on enzyme dosage. The drying effect index (DEI) was defined as the difference in enzymatic digestibility between the OD and ND pulps and it was concluded that more enzymes cause a higher DEI at the initial stage of enzymatic hydrolysis, and the highest DEI was also observed in the earlier stages with the higher enzyme dosage. Once the DEI reached maximum, it began to reduce as enzymatic hydrolysis progressed due to pore formation by cellulose conversion to sugars. However, there was no big difference in the DEI with lower enzyme dosage because slower hydrolysis with less enzymes caused more enzyme inhibition. Thus, the pores formed by cellulose conversion during hydrolysis with less enzymes could not improve enzyme accessibility after 120 h.

It was also found that the drying effect was well correlated with porosity in cellulose, as measured by the water retention value (WRV) and Simons' staining. The porosity in cellulose was reduced during drying by pore collapse and thus, enzyme accessibility decreased accordingly. The pore collapse was partially reversed by rewetting the pulp, however, there was an irreversible structural change that decreased the final enzymatic digestibility of the OD pulp. However, irreversible change in crystalline structure by drying once was not observed through the 13C solid-state NMR, and thus, decreases in enzymatic digestibility did not seem to be related to the crystalline structure of cellulose.

**Author Contributions:** Conceptualization, B.K.; methodology, B.K.; formal analysis, B.K. and J.J.; investigation, B.K. and S.-M.C.; resources, B.K.; data curation, S.-M.C.; writing—original draft preparation, B.K.; writing—review and editing, B.K.; visualization, J.J.; supervision, B.K.; project administration, B.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was carried out with the support of the R & D Program for Forest Science Technology (Project No. 2020226A00-2022-AC01) provided by the Korea Forest Service (Korea Forestry Promotion Institute).

**Acknowledgments:** The NMR analysis in this work was performed by Rui Katahira, a research chemist at the National Renewable Energy Laboratory.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


© 2020 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Lignin to Materials: A Focused Review on Recent Novel Lignin Applications**

**Osbert Yu <sup>1</sup> and Kwang Ho Kim 2,3,\***


Received: 4 June 2020; Accepted: 2 July 2020; Published: 3 July 2020

**Abstract:** In recent decades, advancements in lignin application include the synthesis of polymers, dyes, adhesives and fertilizers. There has recently been a shift from perceiving lignin as a waste product to viewing lignin as a potential raw material for valuable products. More recently, considerable attention has been placed in sectors, like the medical, electrochemical, and polymer sectors, where lignin can be significantly valorized. Despite some technical challenges in lignin recovery and depolymerization, lignin is viewed as a promising material due to it being biocompatible, cheap, and abundant in nature. In the medical sector, lignins can be used as wound dressings, pharmaceuticals, and drug delivery materials. They can also be used for electrochemical energy materials and 3D printing lignin–plastic composite materials. This review covers the recent research progress in lignin valorization, specifically focusing on medical, electrochemical, and 3D printing applications. The technoeconomic assessment of lignin application is also discussed.

**Keywords:** lignin valorization; lignin applications; 3D printing; electrochemical material; medical application

#### **1. Introduction**

The overdependence on fossil fuels has raised increasing concerns about climate change and an energy crisis, which has warranted research to search for renewable and clean energy alternatives. Lignocellulosic biomass is the most abundant and renewable source of organic carbon on Earth, presenting the best option to achieve a sustainable biorefinery in the future [1]. In recent decades, there has been significant research conducted to convert biomass components into biofuel and value-added products. Although some sectors in biorefineries, including lignocellulosic bioethanol, have been widely researched, the economic feasibility is often discussed when it comes to commercialization. From the technoeconomic perspective, the success of future biorefineries is highly dependent on lignin valorization.

Lignin can be obtained from a variety of natural sources, including woody biomass, agricultural residues, and energy crops. Regardless of the type of lignin, there are typically two pathways for lignin valorization. One pathway uses the lignin as a macro-polymer to produce valuable materials; the other pathway involves the depolymerization of lignin into low-molecular weight monomers [2]. Lignin monomers can undergo derivatization through various chemical processes to be converted into desired products [3].

The world annually produces around 100 million metric tons of lignin, worth approximately 732.7 million USD [3]. Since lignin is a cheap and abundant natural polymer, much research has been spent on valorizing lignin. In recent decades, perspectives on lignin have changed from a waste

product used as a low-grade fuel and animal feed to valuable products such as polymers, adhesives, and others [3–6]. More recently, lignin valorization has received interest in other sectors, like the medical and electrochemical energy materials sectors. The scope of this review will focus on the valorization of technical lignin in advanced materials.

#### **2. Technical Lignins**

In nature, lignin is an amorphous phenolic polymer that is randomly branched and crosslinked with cellulose and hemicellulose [7]. To utilize lignin, it needs to be isolated from biomass through various processes, and the extracted lignin is called technical lignin. Since most technical lignins are available in the pulp and paper industries as byproducts, the production of lignin will continue to increase as lignocellulosic ethanol emerges. Considering that the lignin extraction method modifies the native structure of lignin, it is important to understand how the process affects lignin structures to develop effective lignin valorization technologies. There have been many review articles that thoroughly discuss lignin separation methods and their influence on the structures and properties of the extracted lignin [8–11]. The following overview briefly introduces common industrial processes in which lignin is extracted from biomass.

Figure 1 illustrates several processes to extract technical lignins from lignocellulosic biomass. Among the technical processes, Kraft pulping is the major chemical pulping process, accounting for 85% of the total lignin production in the world [12]. The process is performed at a high pH, and about 90–95% of the lignin is dissolved into the black liquor. Kraft lignin is typically precipitated and recovered from black liquor by the addition of acidifying agents. Predominantly, the acidification is carried out by adding either mineral acid (e.g., sulfuric acid) or carbon dioxide, followed by filtering, washing, and drying for the recovery of Kraft lignin. Around 630,000 tons of Kraft lignin are annually produced, and most Kraft lignin is combusted for heat generation, resulting in low-value utilization. The sulfite pulping process is conducted between a pH of 2–12, depending on the cationic composition of the pulping liquor [13]. Lignosulfonates, isolated lignins from the sulfite process, contain significant amounts of sulfur in the form of sulfonate groups. Since lignosulfonates are widely available, lignosulfonates were used in a wide range of applications, such as dispersants, flocculants, concrete additives, and composites [14].

**Figure 1.** Processes for the extraction of technical lignins.

The organosolv process constitutes the fractionation of biomass components through treatment using an organic solvent, such as ethanol, ethylene glycol, acetone, tetrahydrofuran, and γ-valerolactone [15,16]. Since the organosolv process is conducted in the absence of sulfur, it has recently been utilized more so than Kraft and sulfite pulping. Furthermore, the large-scale production of organosolv lignin is expected from the emerging cellulosic ethanol sectors, which offers significant opportunities for lignin valorization.

In addition to the conventional pulping and biomass pretreatment processes such as steam explosion, dilute acid, and ammonic fiber explosion, there have been significant advances in biomass fractionation to extract high-quality lignin using novel solvents. For example, ionic liquids (ILs) and deep eutectic solvents (DESs) have attracted considerable attention as promising agents for biomass fractionation due to their high solvation capacity in the dissolution of biomass components [17,18]. The technical lignins obtained from ILs and DESs have been found to retain their original structures; the eco-friendly properties of such solvents offer new options for lignin extraction.

#### **3. Medical Applications**

#### *3.1. Wound Dressings*

Hydrogels are three-dimensional (3D) networks of polymers; their hydrophilic structures make them capable of absorbing and holding large amounts of water in their 3D networks [19,20]. Hydrogels have gained significant attention in biomedical sectors because they can be used for drug delivery, tissue engineering, and antimicrobial materials [21]. Recently, lignin has been viewed as a promising material for the production of hydrogels, because lignin possesses great antioxidant and antibacterial properties [22]. Furthermore, hydrogels have a high absorption capacity, allowing them to effectively remove undesirable metabolites from the wound [23]. Due to lignin's high mechanical strength, the use of lignin in hydrogels helps protect the wound from further injury or contamination [23]. However, due to the relatively low amount of hydroxyl groups and its rigid structure, lignin requires pretreatment for it to be effectively used as a medical material. In this respect, many studies have focused on developing chemical modification strategies of lignin. Such modifications include the introduction of new active sites on lignin by combining other materials and the functionalization of the hydroxyl group to enhance its reactivity.

There are several ways that lignin can be combined with other materials to construct composite wound dressings that are antimicrobial and biocompatible. A study that used a lignin model polymeric compound, dehydrogenate polymer (DHP), in alginate (Alg) hydrogel, determined that the lignin model DHP in hydrogel produced antimicrobial effects against several clinical bacterial strains and did not have toxic effects on human epithelial cells [24]. A stock suspension of 10 mg/mL DHP and 20 mg/mL Alg was used to test for antibacterial properties. The DHP–Alg demonstrated minimum inhibitory concentration values of 0.002–0.90 mg/mL and a minimum bactericidal concentration of 0.004–1.25 mg/mL. Furthermore, DHP–Alg showed higher antibacterial activity against *L. monocytogenes*, *P. aeruginosa,* and *S. Typhimurium* than streptomycin and ampicillin [24]. The mechanism of antibacterial action was speculated to be that DHP in hydrogel interacts with bacterial cell wall synthesis and/or cell wall structure, which could lead to the disorganization of the cell wall. Although more studies need to be conducted on elucidating the antibacterial mechanism of DHP, the results show the potential application of lignin as a wound-healing agent.

Lignin amine and sulfite-pulped lignin can also be used to construct two different wound dressings. Lignin amine can be synthesized from sodium lignosulfonate by a Mannich reaction; the lignin amine can be crosslinked with poly(vinyl alcohol) (PVA) to form a hydrogel. When a solution of silver nitrate was added to the lignin-based hydrogel, the biocompatible hydrogel not only demonstrated enhanced antimicrobial properties against *E. coli* and *S. aureus*, but the hydrogel also exhibited good elasticity [25]. Additionally, the lignin obtained from sulfite pulping can be combined with chitosan dissolved in an acetic acid solution and PVA dissolved in water to form a lignin–chitosan–PVA composite hydrogel. The sulfonate groups in the lignin formed ionic bonds with amino groups in the chitosan; this is what gives the hydrogel a high mechanical strength and high antioxidant activity [23]. The lignin–chitosan–PVA composite hydrogel demonstrated inhibitory effects on *S. aureus* by penetrating its cell membrane. Results from the study suggested that a hydrogel with a greater lignin concentration resulted in an increase in the swelling ratio, hydrophilicity, protein adsorption capacity, tensile strength, and elongation.

Three-dimensional-printed wound dressings can also be constructed from lignin. Poly(lactic acid) (PLA) pellets coated with castor oil can also be combined with a small amount of Kraft lignin (3 wt % or less) and an antibiotic, tetracycline, to create a composite used for 3D printing wound dressings. The printed filament material was treated with the 2,2-diphenyl-1-picrylhydrazyl (DPPH), a free radical standard, to assess its radical scavenging activity, which can potentially be applied in wound care applications [26]. Figure 2 shows photographs of meshes that were 3D-printed from PLA and 2 wt % Kraft lignin. While lignin in the 3D-printing composite provided antioxidant properties to the composite, small amounts of lignin in the composite did not demonstrate antimicrobial properties with *S. aureus*. However, the tetracycline in the composite was what contributed to the composite's antimicrobial properties. More interestingly, the mesh of the PLA–tetracycline–lignin wound dressing can be 3D-printed based on the patients' needs, leading to the possible customization of wound dressings in the future.

**Figure 2.** Meshes 3D-printed from poly(lactic acid) (PLA) and 2 wt % lignin with grid sizes of 1.5 mm and 1 mm. The middle column shows the side view of each mesh when slightly bent, while the right column shows the close-ups of each mesh. Reprinted from reference [26], with permission from the MDPI.

#### *3.2. Pharmaceuticals*

There are specific compounds derived from lignin that can not only be used to construct pharmaceuticals to alleviative the symptoms of diseases, but can also be used to construct drug delivery materials. While sulfite and Kraft lignins are more commonly used lignins to synthesize pharmaceutical products, recently lignins obtained from steam explosion and the organosolv process have been investigated for pharmaceutical use. The mechanism commonly employed to utilize lignin is to depolymerize lignin and obtain derivatives to construct biologically active compounds. These biologically active compounds can be used to treat the Herpes simplex virus, influenza virus, and other viruses [27]. Since the toxicity of drugs is an important factor in the drug discovery phase, the cytotoxicity of lignin was studied; findings from a study show that lignins are generally safe to consume and no not disrupt cell viability [27]. When synthesizing pharmaceutical drugs, the antioxidant property of lignins is highly valued because its hydroxyl functional groups in their phenolic rings neutralize free radicals and protect molecules from oxidation [27]. For instance, lignins isolated from Bagasse, steam explosion lignins, and lignosulfonates have half maximal inhibitory concentrations of 44.9 µg/mL, 74.6 µg/mL, and 133.6 µg/mL, respectively. Compared to a common antioxidant present in tea, epicatechin, which has a half maximal inhibitory concentration of 42.3 µg/mL, these lignins have enhanced antioxidant properties [28].

Lignin-derived components and products, including polyphenols, may help in controlling diseases, like diabetes, coronary heart disease, and Alzheimer's disease. Results of a study suggest that lignosulfonic acid can be taken with glucose to cause a delay in the uptake of glucose to potentially treat diabetes [29]. The study administered a combination of lignosulfonic acid and glucose as well as glucose alone to determine their effects on 2-deoxyglucose uptake in human colorectal adenocarcinoma cells. Since lignosulfonic acid is a non-competitive inhibitor of α-glucosidase, lignosulfonic acid can inhibit α-glucosidase activity and delay the absorption of glucose in the intestines. Lignosulfonic acid was also demonstrated to inhibit human immunodeficiency virus (HIV) and herpes simplex virus (HSV) transmission when tested with human T-cell leukemia cells, human embryonic kidney cells, and peripheral blood mononuclear cells [30]. Furthermore, lignosulfonates have the potential to be developed into drugs that could decrease oxidative activity, thereby boosting the immune system [3]. Lignin may also helpful in controlling obesity. Out of the dietary fiber components, lignin is reported to be the strongest bile acid adsorbent due to the presence of methoxyl and β-carbonyl groups in lignin. When the lignin binds to the bile, micelles are unable to be produced, thereby resulting in decreased lipid absorption [27].

Table 1 summarizes several studies describing the value products that can be synthesized from technical lignins. In a study, lignophenols derived from the native lignin of Japanese cedar were shown to decrease oleate-induced apolipoprotein-B secretion and reduce cholesterol in HepG2 cells (human liver cells), thereby potentially preventing coronary heart disease [31]. These lignophenols were derived using a phase separation process that included cresol and sulfuric acid; lignophenols were then dissolved in dimethyl sulfoxide before being cultured with HepG2 cells. Another study presents three steps in which lignocellulose can be depolymerized to form platform chemicals, which can then be used to synthesize biologically active compounds with the use of a DES [32]. The DES used was a mixture of choline chloride and oxalic acid. Biologically active compounds that can be synthesized include tetrahydro-2-benzazepines. Tetrahydro-2-benzazepines are present in alkaloids, such as galantamine, that could treat Alzheimer's disease. In this three-step process, the only by-product was water. Furthermore, biologically active compounds derived in this study were also shown to be effective against *S. aureus*. Figure 3 summarizes the reaction pathway to synthesize biologically active compounds that likely exhibit antibacterial or anticancer activities.

**Figure 3.** Summary of the reaction pathway to synthesize the biologically active compounds. Reprinted with permission from [32]. Copyright (2019) American Chemical Society.


**Table 1.**Summary of the studies on the pharmaceutical applications of lignin.

Materials containing lignin and lignin derivatives can also be effectively used to deliver drugs inside the human body. Perhaps the largest advantages of lignin-based nanoparticles for delivering drugs are their inexpensive and non-toxic properties. For instance, lignins derived from sugarcane can be used as a drug carrier to deliver methotrexate and treat rheumatoid arthritis [33]. Lignin nanoparticles encapsulating iron oxide could effectively deliver Sorafenib and Benzazulene in alkali media [34]. Hydrogels produced from varying concentrations of starch, Kraft lignin, hemicellulose, and citric acid were tested for their ability to deliver drugs effectively [35]. To characterize the performance of these hydrogels, the swelling capability was analyzed because it indicates the crosslinking density and ultimately the amount of water absorbed in the hydrogel. Based on the experiments testing the diffusion of water into the gel, the hydrogel has the potential to swell to 1380% at a pH of 9 and the potential to swell to 345% at a pH of 4. The pH-dependent swelling behavior is desirable for hydrogels, because the diffusion rates of molecules in and out of the gel can be controlled.

Since pharmaceutical products are a common source of water pollution, lignin can adsorb these pharmaceuticals to alleviate water pollution. Kraft lignin and α-chitin powder from crab shells were mixed with hydrogen peroxide to create a sorbent that adsorbed ibuprofen and acetaminophen, which are commonly used drugs that cause water pollution [36]. Figure 4 summarizes the process to construct the biosorbent with a high removal efficiency of ibuprofen and acetaminophen. Several mechanisms for the adsorption process were proposed. One such mechanism was ion−dipole interactions; there could be electrostatic interaction between pharmaceutical ions and the negatively or positively charged surface of the sorbent surface. Another mechanism could be hydrogen bonds forming between the pharmaceutical drug and the sorbent. Such sorbents can potentially be reused. For instance, ethanol was an effective eluent for ibuprofen with a yield of 82.2%, while methanol was an effective eluent for acetaminophen with a yield of 80.8%.

**Figure 4.** Biosorbent consisting of chitin and lignin removing ibuprofen and acetaminophen with high efficiency. Reprinted from reference [36], with permission from Elsevier.

In another application, lignin was used as a carbonaceous precursor with TiO<sup>2</sup> to form a photocatalyst that degraded acetaminophen, or Tylenol, in one hour of solar radiation [37]. Photocatalysts that experienced thermal treatment in nitrogen rather than in air maintained more carbon from the lignin. This caused a higher light absorption and decreased the photocatalytic efficiency. In another study investigating the use of lignin to adsorb fluoxetine, alkali lignin with low sulfur content and PVA were dissolved to form a solution used for electrospinning [38]. Nanofibers recovered from the electrospinning process then went through a thermostabilization processes and an acidic bath to increase the strength of the fibers. A lignin:PVA ratio of 1:1 resulted in the optimal adsorption capacity; this lignin−PVA nanofibrous membrane adsorbed around 38% of the fluoxetine. The authors

of the study hypothesized the sorption mechanism to be the phenol groups in lignin-forming hydrogen bonds with the amino or Fluor groups of the fluoxetine.

#### **4. Electrochemical Energy Materials**

In the past decade, lignin has been increasingly investigated for its potential incorporation in the production of battery materials and supercapacitors for the main advantage of being environmentally friendly. With the increasing manufacturing of lignin in recent years, lignin has been available at a low cost. The relatively low cost of lignin makes it an attractive ingredient for the production of anodes for lithium batteries, gel electrolytes, binders, and sodium batteries [39]. Lignin is also valued for the production of energy materials due to its high carbon content that is greater than 60 wt %. Specific functional groups of lignin, such as benzyl and phenolic groups, act as active reaction sites for ions to be stored in applications to supercapacitors. Additionally, the abundant oxygen atoms in lignin are integral for facilitating electrolyte ion adsorption and redox reactions for supercapacitors. Lignin can be used for porous carbon structures in supercapacitors due to its favorable crosslinked structure [2]. While lignin types like lignosulfonate can act as a sulfur-doped agent in batteries or supercapacitors, alkali lignins have shown to be suitable for electrospinning processes to construct nanomaterials [39]. There are generally two processes commonly employed to derive carbon from lignin for energy material applications. One process utilizes precursor carbonization and carbon activation, while the other process uses chemical activators before carbonization and activation that happen at the same time [2]. Graphene, a promising carbon material for electrochemical energy materials, can be synthesized from lignin by catalytic graphitization, carbonization, or oxidative cleavage along with aromatic refusion [40–42].

The incorporation of lignin results in a high gravimetric capacitance and great cycling durability for energy materials. Table 2 shows the summary of recent studies on the electrochemical applications of technical lignins. In an experiment with electrospun carbon nanofibers produced from alkali lignin−PVA solutions, the capacitance of the supercapacitor reduced by 10% after 6000 cycles of discharging and charging [43]. Furthermore, a higher energy value of 42 W h kg−<sup>1</sup> and a power density of 91 kW kg−<sup>1</sup> were reported. The experiment suggested that an increasing amount of lignin in precursor nanofibers resulted in a decreased average pore size, increased pore volume, and increased specific surface area. In another study, electrodes constructed from 75 wt % alkali lignin and 25 wt % 0.5 M sodium sulfate electrolyte resulted in one of the highest specific capacitances for electrodes produced from biopolymers: 205 F g−<sup>1</sup> [44]. The mesopore range of carbon fibers had a wide pore distribution, which contributed to excellent electrochemical performance. Additionally, hierarchical porous carbons can be derived from steam explosion lignin through the carbonization−activation method. Lignin-based hierarchical porous carbons showed a high capacitance of 286.7 F g−<sup>1</sup> at 0.2 A g−<sup>1</sup> [45]. The structure of lignin provided advantages to the electrochemical performance, including accessible ion transportation pathways and a high surface area.

Lignin-derived materials used to construct batteries have performed similarly with commercial graphite and other materials commonly used in the industry. Ball-milled hydrolysis lignin can be used for low-rate power sources. A study investigated the performance of a battery that used hydrolysis lignin as the lithium battery cathode material [46]. The cathode material had 76 wt % hydrolysis lignin, 13 wt % carbon black, and 11 wt % polytetrafluoroethylene (PTFE)-based binder. Lithium batteries using this hydrolysis lignin as a cathode material achieved a high discharge capacity of 450 m A h g−<sup>1</sup> .

Additionally, acetone was used to extract lignin from a corn stalk lignin precursor; the extracted lignin was used to construct hard carbon materials through stabilization in nitrogen, carbonization in nitrogen, and hydrogen reduction [47]. Figure 5 shows the pyrolysis reaction mechanism that occurred after the lignin was extracted from a precursor with acetone. The hard carbon had an initial discharge capacity of 882.2 m A h g−<sup>1</sup> at 0.1 ◦C and retained a charge capacity of 228.8 m A h g−<sup>1</sup> at 2 ◦C for 200 cycles. In another study, a blend of lignin, PLA, and elastomeric polyurethane was used to create carbon nanofibers through electrospinning. These nanofibers led to increased porosity levels and additional lithium storage sites [48].


**Table 2.** Summary of the studies on the electrochemical applications of lignin.

**Figure 5.** Reaction mechanism using the pyrolysis technology. Reprinted from reference [47], with permission from Elsevier.

#### **5. 3D Printing Lignin**−**Plastic Composites**

Blending lignin with various plastic materials to form 3D printing composites has recently received attention. The ideal 3D printing material has excellent extrudability to ease the process of 3D printing while also being strong so that the final printed material can retain its structure. Lignin has several structures that are advantageous for 3D printing: aliphatic ether groups, β-O-4′ linkages, and oxygenated aromatic bonds [49,50]. Incorporating lignin into plastic materials traditionally used for 3D printing results in composites that can be used for a more environmentally friendly and cost-effective 3D printing process.

There has also been recent discussion with energy sacrificial bonds as a mechanism for strong biomaterials, including lignin. Such sacrificial bonds dissipate energy through rupturing and reform by stretching. In light of this, a study investigated Zn-based coordination bonds between lignin nanoparticles and an elastomer matrix [51]. Such bonds were found to facilitate the dispersion of lignin in the matrix, thereby enhancing the strength of the composite. This study reported that a lignin loading as high as 30 wt % can increase the strength, ductility, and toughness of thermoplastic elastomers, which can be used for 3D printing.

Several studies show that lignin can enhance structural properties of 3D printing materials. Table 3 summarizes recent studies on the application of technical lignins to 3D printing materials. In a study, when Kraft lignin was blended with acrylonitrile butadiene styrene (ABS), the result was a more brittle structure; this composite demonstrated low tensile energy and strength [49]. However, when 10 wt % acrylonitrile butadiene rubber (NBR41) was added to the composite, the chemical and physical crosslinks between the NBR41 and lignin resulted in the improved mechanical properties of the composite relative to common petroleum-based thermoplastics. This composite containing 40 wt % lignin, 10 wt % NBR41, and 50 wt % ABS showed great 3D-printability.


**Table 3.** Summary of the studies on the 3D printing applications of lignin.

In an experiment where organosolv hardwood lignin was blended with nylon, the lignin was found to reinforce the thermoplastic matrix by increasing the stiffness of the structure, leading to increased 3D printability [50]. The melt viscosity was also reduced as a result of blending the lignin, which further increases 3D printability. The proposed mechanism for the bonding in the composite was lignin domains forming hydrogen bonds with the thermoplastic matrix. In addition, a study shows that 20–40% lignin can be used with PLA to form a matrix material for 3D printing. In the study, characterization techniques including thermogravimetric analysis, X-ray diffraction, and scanning electron microscope indicated that lignin is a nucleating agent that increases the crystallization of PLA [52]. This composite material resulted in great extrudability and flowability; it was also observed that lignin did not agglomerate. In another study, alkali lignin and organosolv lignin were acetylated to improve the compatibility with PLA to form a composite [53]. Even though the lignin decreased the crystallization behavior of PLA, it drastically improved the thermal stability of PLA and increased the elongation at break. The acetylated lignin was observed to prevent PLA from undergoing hydrolytic degradation.

Besides combining lignin with PLA, other studies have shown the diverse types of plastics that lignin can be blended with to construct effective 3D printing materials. For instance, authors of a study constructed photoactive acrylate resins used for 3D printing from up to 15 wt % acylated organosolv lignin, resin bases, a reactive diluent, and other compounds [54]. Figure 6 shows the reaction used for lignin acylation to construct resins. Even though the resulting lignin resin had decreased thermal stability compared to the commonly available resin, the resin exhibited characteristics that were favorable for 3D printing. The resin had increased ductility and resulted in 3D prints that were uniformly fused, high-resolution, and tough.

Additionally, a study reports that 20 wt % biorefinery lignin from *P. radiata* combined with polyhydroxybutyrate (PHB) improved the surface quality and reduced the shrinkage of the 3D printing material compared to the PHB alone [55]. Figure 7 below is a flow chart depicting how lignin can be extracted from softwood chips and combined with PHB to construct an improved 3D printing material. The presence of lignin was reported to reduce wrapping by values between 38% and 78% when compared to PHB, used as the sole printing material. This reduction of shrinkage was possibly due to the complex bulk lignin structure in addition to the lack of interfacial tension between the lignin and the polymer matrix.

**Figure 6.** Reaction to synthesize acylated lignin for the construction of resin. Reprinted with permission from [54]. Copyright (2018) American Chemical Society.

**Figure 7.** Flow chart showing 3D printed product constructed from high-temperature mechanical pretreatment (HTMP)-derived lignin and polyhydroxybutyrate (PHB). Reprinted from reference [55], with permission from Elsevier.

#### **6. Perspectives**

In the past decade, there have been more investigations conducted on using lignin to construct medical materials, electrochemical energy materials, and 3D printing composites. From the limited technoeconomic assessments conducted, it is difficult to predict the economic trajectory of lignin valorization. Since lignin valorization is an emerging concept, production costs may be higher than conventional processes [3]. According to a technoeconomic assessment analyzing catechol production from lignin, the lignin depolymerization and product separation stages accounted for a total of 55% [3]. For the production of lignin micro- and nanoparticles, their investment cost was 160 million USD, with an atomizer and separation system costing roughly 65% of the capital investment [56]. Assuming Kraft lignin and lignosulfonates are used, manufacturing costs can vary between 870 and USD 1170 per ton. The production capacity of around 70% of North American Kraft pulp mills is restricted by the operation of recovery boilers. Another challenge is finding the optimal membrane used for the Kraft process that will result in the highest lignin yield and purity [57]. Factors to consider for future scale-up efforts would be the size of the market, price volatility of the desired product [3], and the cost of feedstock in addition to solvents [56]. Lignin commercialization is likely with increased international collaboration, as this will mobilize resources and facilitate the development of innovative pathways [57].

Materials constituting lignin and its derivatives have potential in the medical sector due to their versatility as a wound dressing, pharmaceutical drug, drug delivery system, and drug removal material. However, it is difficult for lignin to be absorbed by the human's small intestine or stomach because of its high molecular weight and polydispersity [27]. There have also been studies suggesting that lignin can adsorb toxins, cholesterol, and surfactants, which can negatively impact human health [27]. Much of the research conducted is still at the proof of concept stage and pharmaceutical drugs will need to go through clinical trials before determining their efficacy. Regulatory bodies, like the US Food and Drug Administration, will need to approve the medical products even before they can be commercialized [58]. Clinical trials are time-consuming and costly; thus, there needs to be sufficient confidence in the research done and the viability of the production process before pursuing clinical trials.

While lignin is cheap and eco-friendly as a feedstock material for battery and 3D printing materials, a significant challenge is that it is hard to control or isolate lignin's complex recalcitrant structures. Even when fractionation is employed to isolate a lignin structure, fractionation often results in a mixture of products [59,60]. The electrochemical performance of bio-based battery materials depends on pore features, surface chemistry, and the structure of the lignin raw material [2]. Even though lignin is a relatively cheap starting material, it is still challenging to produce high performing carbon materials for supercapacitors and batteries. Since some lignins cannot stabilize easily, they convert to carbon nanofibers and consequently their morphologies are difficult to control; this makes high-performing lignin-based electrodes difficult to achieve [61]. Furthermore, lignin reaction mechanisms are not well understood [2].

Incorporating lignin in 3D printing materials decreases the reliance on typical petroleum-based plastic 3D printing materials. Doing so not only makes the 3D printing materials more environmentally friendly, but it also reduces costs as lignin is much cheaper than typical 3D printing materials [62]. Additionally, a myriad of materials can not only be created by 3D printing, but they can also be customized. This is of interest in the medical sector, where wound dressings can be 3D printed and customized to best suit the patients' needs. While lignin and other materials are being developed as 3D printing materials, they will most likely not replace traditional 3D printing materials including plastics [62]. Studies conducted so far in this current review suggest that lignin can be used to enhance the properties of the lignin–plastic composite 3D printing material but will not be the main feedstock material for 3D printing.

Even though the technoeconomic assessments of lignin valorization present several challenges, lignin's versatility has attracted significant efforts to valorize lignin. To illustrate lignin's versatility as a renewable material, Figure 8 shows the number of publications and citations searched in the

Web of Science with the keywords: "lignin application." As presented, the number of publications and citations have substantially increased over the past decade, indicating that significant efforts have been made in lignin valorization. Considering that there has been increased market demand for renewable materials in a wide range of industries, innovative lignin applications will continue to be developed indefinitely.

**Figure 8.** Number of publications (**left**) and citations (**right**) searched in the Web of Science with the keywords "lignin application" (as of May 30th 2020).

**Author Contributions:** Conceptualization, O.Y. and K.H.K.; Methodology, O.Y. and K.H.K.; Writing—Original draft preparation, O.Y.; Writing—Review and editing, O.Y. and K.H.K.; Supervision, K.H.K. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the Korea Institute of Science and Technology–The University of British Columbia Biorefinery onsite laboratory project, grant number 2E30740.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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