The Recovery of Lignin from Reed Using Solid-DES Extraction (SDESE) ^†

Abstract

The current work presents a sustainable, environmentally friendly method for lignin extraction by applying deep eutectic solvents (DES), which are biodegradable. The Solid-DES extraction (SDESE) method of recovering lignin from reeds is presented. DES based on betaine hydrochloride and lactic acid (BeHCl, LA) was prepared and applied in SDESE at a ratio of 1:20 wt %. Fourier-transform infrared spectroscopy (FTIR) was used to characterize the materials. The DES-extracted lignin (DEL) from the reeds exhibited an interesting structure due to the rearrangement of syringyl, guaiacyl, and parahydroxyphenyl units that occurred as a result of DES breaking the initial raw material structure.

1. Introduction

Reeds are one of the most widespread species of wetland plants on Earth. Initially, reed was widely used in papermaking, but due to its fast-growing properties and high biomass yield, reed is recognized as a promising source of renewable energy [1]. Reed is used as a raw material in the wood industry and is considered to be a biotechnological product with potential for valorization, both from the point of view of organic biomass and as a possible inorganic source, especially due to its high silicon content [2]. In the current work, analyses of reed biomass as a potential source of lignin valorization were carried out.

Lignin has gained interest due to its particular advantage as a source available for biorefineries, but also in other fields of application, such as the pharmaceutical industry [3] and biocomposite production as a char agent [4], and other applications [5] that make lignin an invaluable product for recovery and valorization. Its biopolymer structure allows for numerous structural rearrangements depending on the applied treatments. Lignin is found in plant biomass as having a protective and supportive role for plants. It contains three phenylpropane units of syringyl, guaiacyl, and parahydroxyphenyl. These units can be variously linked, either by C-C bonds or β-O-4’ [6] ether bonds. Due to the high structural heterogeneity induced by its number and types of functional groups, and the way the atoms are arranged, as well as the type of bonds established between the three units, lignin is a material with multiple advantages.

The use of deep eutectic solvents (DESs) in extraction processes has become a well-known approach exploited by many research groups. The formation of eutectic mixture occurs through the association of two components: a hydrogen bond acceptor (HBA) and hydrogen bond donor (HBD) [7]. DESs have many advantages compared to conventional solvents: they are biodegradable, non-flammable, easy to prepare, and have some particular characteristics based on the formation and number of hydrogen bonds between the precursors, which give them special properties depending on the type of each constituent.

2. Materials and Methods

2.1. Materials

The raw material (reed, Phragmites australis) was harvested from the Danube Delta (Mila 23, 45°13′45″ N 29°14′54″ E, 2 m altitude). Sulfuric acid 95–97%, used for the determination of the lignin content, was from Merck (Merck Group, Darmstad, Germany), and barium chloride dihydrate 99% and acetone were from Scharlau (Scharlab, Barcelona, Spain). Commercial lignin and sodium hydroxide 97% were supplied by Sigma-Aldrich (Merck Group, Darmastad, Germany). To prepare the deep eutectic solvents (DES), D,L lactic acid 88–90% from Scharlau (Sclarlab, Barcelona, Spain) and betaine hydrochloride 99% purchased from Merck (Merck Group, Darmstadt, Germany) were used.

2.2. Composition of the Raw Material

2.2.1. Determination of Extractables Content

One gram of dry and well-crushed reed was weighed and placed in a round-bottomed flask. A volume of 60–80 mL of acetone solution [8] was added to the flask and subjected to a thermal process at 90 °C for 2 h at 450 rpm in a reflux installation built in the laboratory. The precipitate obtained was washed with double-distilled water in a Universal 320R Centrifuge (Hettich, Tuttlingen, Germany) at 12 °C, 8078 rcf, 30 min. After being well washed, the precipitate was placed in the freezer at −80 °C for 3 h. Afterwards, the frozen precipitate was freeze-dried in a SCANVAC CoolSafe freeze-dryer (Labogene, Lillerød, Denmark) at −50 °C. The resulting sample, after drying, represented Extractables-Free-Reed (EF-Reed). This sample was used to determine the contents of cellulose, hemicellulose, and lignin from the biomass. The amount of extractables lost was calculated according to the following formula:

$$Reed Extr.(\%)=\frac{m_i-m_{dry}}{m_i}\times 100,$$

(1)

where m_i = the initial mass of reed and m_dry = the dry mass of reed without extractables (EF-Reed).

2.2.2. Determination of Hemicellulose Content

One gram of EF-Reed was mixed with a volume of 100 mL of 0.5 M NaOH solution, and the mixture was heated at 80 °C for 3 h 30 min [9]. After this stage, the precipitate was cooled, centrifuged (8078 rcf, 30 min, 12 °C), and washed with ddH₂O for 5–6 cycles, until a neutral pH was reached. The precipitate was freeze-dried. This precipitate was considered to be Hemicellulose-Free-Reed (HF-Reed). The amount of hemicellulose solubilized in the alkaline solution was determined using the following formula:

$$Hemicellulose (\%)=\frac{m_i-m_f}{mi}100,$$

(2)

where m_i = the initial mass of BSG represented by EF-Reed and m_f = the dry mass of the precipitate without hemicellulose (HF-Reed).

2.2.3. Determination of Lignin Content

The acid hydrolysis method was applied to determine the lignin content. A volume of 80 mL of H₂SO₄ 98% [10] was added over 1 g of EF-Reed. The mixture was kept at room temperature for 24 h under magnetic stirring at 200 rpm. After this step, it was heated at 30 °C for one hour. After cooling, the residue (containing mostly cellulose) left after the hydrolysis was separated and the lignin was precipitated with double-distilled water and centrifuged at 8078 rcf and 4 °C for 30 min. After this step, a 10% barium chloride solution was added to neutralize the sulfonate groups attached to the lignin. Afterwards, the lignin precipitate was introduced to the centrifuge (the same parameters were maintained) and washed with double-distilled water 10 times.

The precipitate was freeze-dried. The amount of lignin was determined using the formula:

$$Lignin (\%)=\frac{m_{dry}}{m_i}\times 100,$$

(3)

where m_dry = the dry mass of the precipitate from the supernatant after acidic hydrolysis and m_i = the dry mass of initial EF-Reed.

2.2.4. Determination of Cellulose Content

Since the main components of reed biomass are cellulose, hemicellulose, extractables, and lignin, the cellulose content of reed [11] can be determined as follows:

$$m_{cellulose}=100-(m_{hemicellulose}+m_{lignin}+m_{extractables}),$$

(4)

2.3. Lignin Extraction, Characterization of the Raw Material and of Extracted Lignin

2.3.1. DES Preparation

The DES was prepared based on betaine hydrochloride (BeHCl) as a hydrogen bond acceptor and lactic acid (LA) as a hydrogen bond donor at a molar ratio of 1:5. For the solubilization of BeHCl, it was necessary to add water (W) in molar ratio of 5, with a final composition of BeHCl:LA:W in a 1:5:5 molar ratio. The first thermal process was carried out at a temperature of 60 °C in the ultrasonic bath Elmasonic P30H, Singen, Germany, at 37 pulses/minute with power of 100 W for 2 h. After the BeHCl solubilization, the mixture was introduced to the reflux installation at 60 °C for 2 h at 250 rpm.

2.3.2. Solid-DES Extraction (SDESE)

The SDESE method [12] was applied for the lignin extraction. A mixture composed of 1 g of Reed and 20 g of DES was introduced and processed in the reflux installation at 80 °C and 250 rpm for 24 h. After this step, the mixture was cooled to room temperature and treated with a volume of 20 mL of acetone: water 1:1 v/v. This treatment was repeated 5 times. The supernatant was recovered and collected in a container. To remove the acetone, the supernatant was introduced into a Buchi Rotavapor at 60 °C and 40 rpm for 30 min. The pH of the solution was controlled and adjusted to 2.00 to precipitate the lignin [13], then the solution was placed in the freezer at −80 °C for at least 3 h, freeze-dried, and finally weighed. The extraction yield was calculated considering the amount of lignin determined in one gram of reed using acid hydrolysis with sulfuric acid.

2.3.3. Physicochemical Analysis

The FTIR analysis was performed with an IRTracer—100 spectrophotometer (Shimadzu, Tokyo, Japan). The samples were analyzed in transmittance mode over a wavenumber range of 4000–400 cm⁻¹ with a resolution of 4 cm⁻¹.

The density measurement was performed using an EasyD40 densimeter (Mettler Toledo, Greinfensee, Switzerland). The pH measurement was performed with a Seven Compact pH meter (Mettler Toledo, Greinfensee, Switzerland) equipped with an InLab^® Viscous Pro—ISM electrode.

3. Results

3.1. Composition of the Raw Material

Figure 1 describes the main stages carried out for the biomass characterization. Stage 1 describes the harvested reed sample (raw material). In step 2, the reed sample was ground, followed by obtaining the EF-Reed in step 3. From the EF-Reed sample, the hemicellulose content was determined in stage 4 with the formation of the HF-Reed sample, the cellulose mass in stage 5, and the lignin content in stage 6.

The gravimetric calculations resulted in contents of 27.5% of hemicellulose, 48.3% of cellulose, 12% of extractables, and 10.2% of lignin.

3.2. Characterization of Reed

The FTIR spectra of the HF-Reed, EF-Reed, and reed (raw material) are described in Figure 2.

Biomass presents a complex vibration spectrum provided by the existence of three main components: cellulose, hemicellulose, and lignin. In the 3500–2500 cm⁻¹ region, bands corresponding to the OH groups (from the lignin, cellulose, and hemicellulose) at 3449 cm⁻¹ and vibrations of alkyl C–H bonds (predominant in lignin) can be observed. In the region of 2000–1000 cm⁻¹, bands from the structures of the three components were described: the stretching vibrations of the carbonyl linkage at 1734 cm⁻¹ (hemicellulose and lignin) [14], usually attributed to esterified or protonated carboxyl groups, the peak at 1641 cm⁻¹, which could have a contribution from deprotonated, ionic forms of carboxyl (carbonyl vibration), protein amide I, and/or adsorbed water, the peaks at 1450–1319 cm⁻¹, which revealed the deformation vibrations of the CH groups (lignin, cellulose, and hemicellulose), and the peak at 1236 cm⁻¹, which attested to the stretching vibrations of the C–O bond (lignin). At 1024 cm⁻¹, etheric C–O–C bonds were observed, with a shoulder at 1145 and one at 904 cm⁻¹ being given by the presence of C–O groups corresponding to the carbohydrates’ vibrations (cellulose and hemicellulose). In addition, in the “fingerprint” area, there were vibrations of the C–C, C–H linkages.

Compared to the reed spectrum, in the EF-Reed spectrum, the OH band shifted to 3423 cm⁻¹, the band at 1734 cm⁻¹ was shifted to 1726 cm⁻¹ (C=O bond), the band at 1450 cm⁻¹ shifted to 1454 cm⁻¹, the shoulder at 1145 cm⁻¹ was shifted as a distinct peak to 1161 cm⁻¹, the peak at 1024 cm⁻¹ was shifted to 993 cm⁻¹, and the bands with lower frequencies showed some changes given by the specific vibrations of the C–H, C–C linkages. By removing the hemicellulose from the reed (HF-Reed), several transformations of the vibrational bands occurred. The band corresponding to the OH group shifted from 3423 in EF-Reed and showed a reduced intensity at 3333 cm⁻¹. The band corresponding to the carbonyl group at 1726 cm⁻¹ was significantly reduced and the band at 1641 cm⁻¹ was slightly more pronounced, which could be explained mainly by the formation of sodium carboxylate, reducing its protonated form and increasing the content of its ionic form. The deformation bands of the CH bonds underwent changes, with the band at 1454 cm⁻¹ being reduced and becoming less prominent than the bands of the CH deformation from the cellulose (1368 cm⁻¹) and syringyl units (1315 cm⁻¹). The bands from 1161–904 cm⁻¹ were preserved, as these came from both hemicellulose and cellulose, as well as lignin, but they were shifted to 1159–897 cm⁻¹. The bands from lower frequencies were preserved as well, being assigned to the C–H bonds from various structures.

3.3. Characterization of DES

The density of the DES (1.4330 g cm⁻³) was higher than that of water, similar to previous reports [15]. DESs are generally much denser due to the hydrogen bonds formed between HBA and HBD. By adding carboxylic acid (lactic acid) to the system, the pH dropped significantly to 1.20.

The formation of DES was analyzed using FTIR spectroscopy. Figure 3 shows the characteristic spectra of the precursors and DES. The spectra recorded in IR show some shifted bands of DES compared to the spectra of the initial components. There are some groups involved in making new hydrogen bonds [16,17]. The broad band with a minimum at 3337 cm⁻¹ in DES, which reflects vibrations of -OH involved in H-bonds, was much more pronounced than for lactic acid (LA), and shifted from approx. 3441 cm⁻¹ in LA. This red shift was similar to the shift reported previously for a DES formed of BeHCl:LA in the same ratio of 1:5 [18]. The flattened character of the corresponding band in BeHCl, which was almost absent due to less H-bonds, was not reflected in the DES spectrum, confirming that new and more abundant H-bonds were formed in the DES compared with both LA and BeHCl. The involvement of the COOH groups in the H-bonds was indicated by the shift of the C=O vibration band from 1715 (LA)/1720 (BeHCl) cm⁻¹ to 1719 cm⁻¹ in DES.

Most of the bands from BeHCl disappeared or were significantly reduced and shifted, with the spectrum of DES being very similar to that of LA, with small differences. The same was previously reported for BeHCl:LA without water [18]. The bands corresponding to the C–H bonds of alkyl groups were shifted in DES compared to LA: 2893/2897 cm⁻¹ for the symmetric vibration of CH₂ and CH₃ and 2992/2989 cm⁻¹ for the asymmetric vibration of CH₃ in DES/LA). BeHCl had three bands in this region: 2897 cm⁻¹ probably from the symmetric vibration of the CH₃ groups, 2967 cm⁻¹ for the asymmetric vibrations of the CH₃ and/or CH₂, and 3024 cm⁻¹ for the asymmetric stretching vibration of (CH₃)₃N⁺ [19]. The bands 2967 cm⁻¹ and 3024 cm⁻¹ from BeHCl were significantly reduced and/or shifted in DES due to interactions between the methyl groups of BeHCl and the oxygen of LA [17]. There was an additional small peak in LA at 2947 cm⁻¹ which would be characteristic of asymmetrical CH₂ vibrations, but as lactic acid does not have a CH₂ group, this band could come from some impurities. The sharp band at 673 cm⁻¹ in BeHCl is not visible in DES, indicating the involvement of C–OH in H-bonding. This band was not present in LA, which had a small band at 635 cm⁻¹ shifted to 631 cm⁻¹ in DES. Another band from BeHCl still visible in DES was the band at 988 cm⁻¹, which was significantly reduced and shifted to 974 cm⁻¹ in DES. This band could reflect the stretching vibration of C-O and/or C-C as previously assigned in the case of serine [20,21]. Other shifted bands identified were: 1648 (LA) to 1645 (DES) cm⁻¹; 1458 (LA)/1479 (BeHCl) to 1466 (DES) cm⁻¹; 1377 (LA)/1410 (BeHCl) to 1391 (DES) cm⁻¹; 1205 (LA)/1196 (BeHCl) to 1211 (DES) cm⁻¹; 1120 (LA)/1130 (BeHCl) to 1117 (DES) cm⁻¹; 1040 (LA) to 1042 (DES) cm⁻¹; 820 (LA) broadened and shifted to 818 (DES) cm⁻¹; 746 (LA)/770 (BeHCl) to 750 (DES) cm⁻¹. The following additional bands were not visible or were significantly reduced in DES: 1246, 1086, 939, 887 cm⁻¹ (BeHCl) and 922, 868 cm⁻¹ (LA).

3.4. Lignin Extraction and Characterization

The extraction yield was calculated considering the amount of 10.2% existing in one gram of reed. Based on these data, SDESE presented an extraction yield of 52%. This value was in agreement with the extraction yield used by Kumar and team [22], when they used DES based on betaine/lactic acid (1:5 molar ratio) to extract the lignin from rice straw.

The extracted lignin was characterized using ATR-FTIR, comparing it to a commercial lignin sample—Figure 4.

Figure 4 shows the superimposed spectra of the DES-extracted lignin (DEL) and commercial lignin (CL). The numbers marked in yellow denote the similar bands of the two lignins (DEL and CL). The band at 3526 cm⁻¹ in DEL belonged to the stretching vibrations of the O–H bonds attached to the aromatic ring. The bands at 2972 and 2897 cm⁻¹ corresponded to the vibrations of the CH groups (sp³ hybridized) existing in the propenyl radicals attached to the phenolic units. As can be seen from Figure 4, these bands were slightly shifted compared to CL. The bands at 1595 and 1508 cm⁻¹, corresponding to the aromatic ring vibrations, were at the same wavenumbers in the two lignins. The characteristic vibrations of the syringyl units were at 1331 cm⁻¹ in DEL, while in CL, they were slightly shifted to higher frequencies (1365 cm⁻¹). The band characteristic to the deformation vibrations of the guiacyl units was found at the same frequency of 1219 cm⁻¹ in both lignins. In the spectrum of DEL, the peak at 1113 cm⁻¹ denoted the presence of the CH vibration in the syringyl, with a shoulder at 1034 cm⁻¹ reflecting the C–O bonds of DEL and CL. The band at 826 cm⁻¹ represented the C=C bending.

The IR spectra of the two lignins showed similarities given by the vibrations of the specific aromatic units, but also some differences due to the different biomasses from which they were extracted and the applied extraction method that influenced the rearrangement of the structural units in the lignin.

4. Conclusions

Solid-DES extraction (SDESE) with BeHCl:LA:W was applied to extract lignin from reeds, representing a promising method with multiple advantages (biodegradable, relatively cost effective, removable, and recyclable). The extracted lignin showed a similar FTIR spectrum to that of commercial lignin, with some small differences probably determined by the lignin source and method of extraction.