Fast, Easy, and Economical Quantification of Lignin Phenolic Hydroxyl

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Biofuels and Biomass

Fast, easy and economical quantification of lignin phenolic hydroxyl groups. Comparison with classical techniques Luis Serrano Cantador, Esakkiammal Sudha Esakkimuthu, Nathalie Marlin, Marie-Christine Brochier-Salon, Gerard Mortha, and Frederique Bertaud Energy Fuels, Just Accepted Manuscript • DOI: 10.1021/acs.energyfuels.8b00383 • Publication Date (Web): 17 Apr 2018 Downloaded from http://pubs.acs.org on April 18, 2018

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Fast, easy and economical quantification of lignin phenolic

hydroxyl

groups.

Comparison

with

classical techniques Luis Serrano1,2,3,4*, Esakkiammal Sudha Esakkimuthu1,2,3, Nathalie Marlin1,2,3, Marie-Christine Brochier-Salon1,3 ,Gerard Mortha1,2,3 ,Frederique Bertaud5 1

Univ. Grenoble Alpes, LGP2, F-38000 Grenoble, France

2

CNRS, LGP2, F-38000 Grenoble, France

3

Agefpi, LGP2, F-38000 Grenoble, France

4

University of Cordoba, Chemical Engineering Department, Faculty of Sciences, Building

Marie-Curie, Campus of Rabanales, 14014 Cordoba-Spain 5

Centre Technique du Papier, CTP, CS90251, 38044 Grenoble cedex 9, France

*Correspondence to: [email protected]

ACS Paragon Plus Environment

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Abstract

The quantification of lignin functional groups constitutes an essential step for effective assessment of lignin transformation. A new, fast, economical and simple method without the use of harmful solvents is presented for lignin phenolic hydroxyl group quantification. The new method is based in simultaneous conductometric and acid-base titration based on pH in aqueous medium and it provides, in a single measurement, phenolic hydroxyl and carboxyl group quantification. The new method provokes an inherent error due to the phenolic structure of lignin. This error should be 10% maximum for the low amount of condensed structures in the analyzed technical lignin samples. Four industrial lignin samples (PB-Protobind lignin, ORGOrganolsolv lignin, KF-Kraft lignin, IND-Indulin lignin) have been analyzed using the new method and the results were compared with six classical methods (UV, 31P-NMR, 1H-NMR, 13CNMR, Folin Ciocalteu and aminolysis). The results showed good statistical correlation of the new method with the UV and

31

P-NMR, the most significant methods, with coefficients of

variation among 6 and 14%. Moreover, the carboxyl content results presented a high accuracy with a traditional method as tetra-n-butylammonium hydroxide (TnBAH) potentiometric titration.

Keywords: lignin; hydroxyl groups; titration; carboxyl groups


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Introduction

Lignin, the most important aromatic polymer in the nature, constitutes between 15-40% of the dry matter content in biomass.1,2 The copolymerisation of three alcohol monomers: p-coumaryl alcohol, sinapyl alcohol and coniferyl alcohol gives rise to the structural units of lignin: guaiacyl, syringyl, and p-hydroxyphenyl.3,4 The complex structure of lignin is built depending of the proportion of each structural unit and the linkages between them (the most common is the aryl ether bond, β-O-4). The final structure is strongly dependent of many factors such as the plant species, cultivation and growth conditions, and isolation methods, among others.5 Lignin is extracted from lignocellulosic biomass during the delignification process in pulp production by the pulp and paper industry. During this process, lignin is partly hydrolyzed and oxidized giving new functionalities, such as phenol and carboxyl groups. Lignin has recently attracted considerable interest turning to a new vision under the biorefinery concept for its use in high added value applications. Potential applications as low cost carbon fibers, plastics and thermoplastics elastomers, foams, membranes, fuels and chemicals1,6 are transforming the lignin in a new challenge for the research community. The lignin functional groups constitute the key for its transformation into high added value products. The main functional groups in the lignin molecule, having a great impact on its reactivity, are the hydroxyl, methoxyl, carbonyl and carboxyl groups.7 One of the most promising applications for lignin is the substitution of petrochemical-based aromatic compounds in the production of polymer materials.8 For this purpose, the determination of lignin molecular weight and the content of free phenolic hydroxyl groups are essential. The presence of phenolic hydroxyl groups has an important influence on the reactivity and solubility of lignin, being also key functions for further derivatization to functionalize the molecule for specific applications. Its


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quantification provides useful information about the lignin structure and its conversion mechanism.7 Up to date, many physical and chemical methods have been developed to estimate the total phenolic hydroxyl groups in lignin and compared between them in several publications.7-11 The UV method, described the first time by Zakis12 and updated by Tamminen and Hortling,13 is based on the absorption difference of some phenolic units at 300 and 350 nm. The absorbance of ionized phenolic groups in alkali solutions compared with the neutral solution at the mentioned wavelengths is proportional to the free phenolic hydroxyl group content. The Folin and Ciocalteu method is also a UV method recently proposed by De Sousa et al.14 derived from a procedure developed by Folin and Ciocalteu15 in the determination of protein content. The phosphotungstic-phosphomolybdenum reagent used in this method provides a blue color proportional to the total amount of phenolic hydroxyl groups and easy to detect by UV spectrophotometry.14 One of the traditional established methods typically used to determine the phenolic hydroxyl group content in lignin samples is the aminolysis method. This multi-step method was originally developed and tested on acetylated lignin model compounds16 and on milled wood lignin samples.9 After acetylation, aromatic acetyl groups can be selectively deacetylated by pyrrolidine reagent. Thus, the resulting 1-acetyl pyrrolidine can be easily quantified by gas chromatography, which is proportional to the number of phenolic hydroxyl groups present in the lignin samples. Another way to determine the phenolic hydroxyl groups of lignin is by FTIR spectroscopy using the aliphatic and aromatic IR ester bands of acetylated lignin samples at 1745 and 1765 cm-1, respectively.17 It is a simple and fast method, but the use of acetylation can cause problems with the incomplete derivatization and results are only qualitative.


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Finally, NMR spectrometry is amply used by many authors to determine the phenolic hydroxyl groups of lignin. 1H-NMR provides information about the characterization, classification and determination of lignin structure.18,19 The 1H-NMR spectrum of acetylated lignin gives access to hydroxyl group content, but the own lignin structure hinders the accuracy of the measurements due to the existence of a great diversity of protons and linkages between different units. Due to the lack of resolution in the 1H-NMR spectrum using acetylated samples, the use of raw lignin samples, without derivatization, directly dissolved in DMSO or with the presence of D2O facilitates the phenolic hydroxyl groups determination.10 Similarly,

13

C-NMR also provides a

large amount of lignin structural information and it is a very attractive technique for functional group quantification.20 Quantitative analysis can be carried out using acetylated lignin samples to determine the amount of phenolic hydroxyl groups.11 Furthermore, this technique requires large lignin sample quantities, long acquisition times and high purity sample level (free of carbohydrates, ashes or extractives). Finally,

31

P-NMR is a powerful technique to quantify

hydroxyl groups in lignin samples. It was developed by Granata and Argyropoulos21 involving the lignin derivatization with the phosphitylation agent 2–chloro–4,4,5,5–tetramethyl–1,3,2– dioxaphospholane (TMDP). Previously, the agent 2-chloro-1,3,2-dioxaphospholane was tested,21 but the use of TMDP makes possible the discrimination in different spectra areas between phenolic and aliphatic hydroxyl groups, condensed phenol units and carboxyl groups. Other methods such as pyrolytic gas chromatography, periodate oxidation and non-aqueous potentiometric titration with tetra-n-butylammonium hydroxide (TnBAH) or potassium methylate,7,22 are able to quantify the phenolic hydroxyl groups in lignin. All the described methods present important limitations. Some take time and are more or less practical or feasible in simple industrial research and development laboratories (case of non-


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aqueous titrations), or require specific equipment (as spectrometers or chromatographs), or are semi-quantitative (like pyrolytic methods). In this work, a new, easy, fast and economical method to quantify the phenolic hydroxyl groups in lignin samples is presented. It is based on simple simultaneous conductometric and pH-metric titrations in aqueous medium. Four different industrial lignin samples (PB-Protobind lignin, ORG-Organolsolv lignin, KF-Kraft lignin, IND-Indulin lignin) have been used to determine the phenolic hydroxyl group content using six classical methods (UV, NMR,

31

P-NMR, 1H-

13

C-NMR, Folin Ciocalteu and aminolysis) and the comparison with the new method is

presented.

Experimental Section

Raw materials

Four industrial lignin samples were used in this study. (i) Soda lignin from wheat straw (Protobind 1000) (PB) was purchased from Green Value Enterprises LLC. (ii) Kraft lignin from pine (KF) was provided by the Centre Technique du Papier (CTP) (Grenoble, France). (iii) Organosolv lignin (ORG) (namely BioLignin® CIMV process using formic acid/acetic acid/water at 185-210ºC) from wheat straw was purchased from CIMV Company. (iv) Kraft Indulin AT lignin (IND) was purchased from DKSH Switzerland Ltd. and incorporated as reference. All the lignin samples were analysed for chemical composition. Dry matter content was gravimetrically determined by drying samples at 105 °C to constant weight. The ashes content of all the lignin samples was obtained gravimetrically after in-furnace calcinations for 4 h at 525


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°C. Klason lignin was determined by the fraction left insoluble after acid hydrolysis (TAPPI standard T-13) and acid soluble lignin by spectrophotometric method at 205 nm (TAPPI UM250 um-83). Monosaccharides were determined after the injection of 25 µL of acid-soluble lignin filtrate in high performance anion exchange chromatography (HPAEC) with pulsed amperometric detection (PAD).23 Sugars were eluted on a Dionex Carbopac PA1 column (4 mm x 250 mm) using 2 mM of potassium hydroxide at a flow rate of 1 mL/min. Fucose was used as internal standard to calculate the sugars concentration.24 Lignin powder was also studied by ATR-IR to determine the structural composition of samples and the acetylation degree. Samples were analysed by attenuated-total-reflection infrared (ATR-IR) spectroscopy using direct transmittance in a single reflection ATR system (ATR top plate fixed to an optical beam condensing unit with a ZnSe lens) with an MKII Golden Gate SPECAC instrument. Each spectrum was recorded over 16 scans in the range from 4000 to 600 cm-1 with a resolution of 8 cm-1.

Classical methods for phenolic hydroxyl group determination

UV method

A solution dissolving 20 mg of lignin in a mixture of 10 mL dioxane and 10 mL 0.2M aqueous NaOH was prepared. The solution was filtered by nylon 0.45 µm to remove possible undissolved particles. 4 mL of the initial solution was diluted to 50 mL with three different aqueous solutions (0.2M NaOH, buffer pH 6 and buffer pH 12) to reach a final concentration of 0.08 g/L. The UV measurements were carried out with a Shimadzu UV-1800 in the range 200-600 nm using the lignin solution with buffer pH 6 as reference. The absorbance of the maxima observed


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at 300 nm and 350 nm were used for the calculations. The total phenolic hydroxyl group content was calculated according to the Equation (1).13 Total OH phenolic content mmol⁄g = 0.425 x A NaOH + 0.182 x A% NaOH& x a 1 where, A = absorbance, a (L·g-1·cm-1) = correction term = 1/(c·l)·10/17, c (g/L) = lignin solution concentration and l (cm) = path length.

Folin and Ciocalteu method

A calibration curve was obtained using vanillin solution in DMSO. Stock solution of 2 mmol/L in DMSO was prepared and suitable dilutions were made until 0.012 mmol/L, 0.02 mmol/L, 0.04 mmol/L and 0.06 mmol/L. A solution of 50 mg of lignin with 50 mL DMSO was prepared. The dissolution of 1 ml of each sample/reference/standard solution, 3 mL Folin reagent and 30 mL DMSO was kept for 5-8 minutes in a 50 mL gauged flask followed by the addition of 10 mL of aqueous Na2CO3 (200 g/L) and the flask was filled with DMSO until 50 mL. The mixture was let to react for 2 hours with every 20 minutes shaking. Finally, the absorbance at 760 nm was measured using a Shimadzu UV-1800. The use of DMSO facilitates the lignin dissolution. Some authors proposed the addition of small quantities of NaOH during the sample preparation but it has been demonstrated that the Folin reagent is not stable in alkali solutions. The total phenolic hydroxyl group content was calculated according to the Equation (2).25 Total OH phenolic content mmol⁄g = A'()* ⁄k x 50 x 50⁄m

(2)

where, A = absorbance, k (mmol/L) = slope derived from vanillin calibration curve and m (mg) = mass of lignin.


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Acetylation

Acetylated lignin samples were used, first to develop the aminolysis reaction, and second, for quantitative

13

C-NMR analysis.26 More details have been provided in the Supplementary

information.

Aminolysis

The acetylated lignin samples (100 mg) were dissolved in 5 mL dioxane containing 25 mg of internal standard (1-methyl naphtalene). To initiate the aminolysis reaction, 5mL of dioxanepyrrolidine (1/1: v/v) reactant mixture were added. The rates of 1-acetyl pyrrolidine formation were calculated by injecting 0.5 µL reaction mixture into the gas chromatograph with a periodic time intervals (for instances: 5, 10, 20, up to 60 minutes). A GC-Trace (ThermoQuest) gas chromatograph was used with a flame ionization detector, Helium as carrier gas, and a DB-5 capillary column (30 m length, 320 µm internals diameter, 1 µm film thickness). The 1-acetyl pyrrolidine content was calculated according to the Equation (3). Total OH phenolic content mmol⁄g =

, - ./ - 0 ,/ - .1 - 2 - 00

(3)

where, A = chromatographic area corresponding grams to the 1-acetyl pyrrolidine signal, As = chromatographic area corresponding grams to the internal standard signal, WL = weight of lignin (dry matter), WS = weight of internal standard (1-methyl naphthalene) and k = calibration constant. The content of phenolic hydroxyl groups was calculated by extrapolation of the curve at zero time, as described by Dence and Lin, 1992.7


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NMR analysis

NMR spectroscopic measurements were conducted on a Bruker AVANCE400 spectrometer equipped with a 5 mm BB/19F-1H/d Z-GRD probe operating at 100.612 MHz for 13C, 161.982 MHz for

31

P and 400.130 MHz for 1H. Acquisition and data treatment were done using the

LINUX TopSpin 3.2 software. Detailed information about

31

P-NMR,

13

C-NMR and 1H-NMR

analysis has been provided in the Supplementary information.10,11,21

Fast method for phenolic hydroxyl group determination

A rapid method in aqueous medium based on simultaneous conductometric and acidic-base titration by pH was evaluated to determine the amount of phenolic hydroxyl groups and compared with the traditional methods previously presented. At the same time, this method is able to provide the carboxyl content amount.27 A solution containing 1 g of lignin sample, 500 mL of fresh deionized water, free of carbonates, and 8 mL of sodium hydroxide 1M (large excess) was prepared. After stirring until total dissolution (around 15 min for most of the studied lignin samples), the solution was titrated back with 1M hydrochloric acid using a Schott automatic burette. Simultaneous measurements of conductivity and pH were recorded with a Crison pH GLP 21 equipment. During titration, the titration vase was protected from air by a plastic film, to prevent sodium hydroxide neutralization by atmospheric CO2.


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Carboxyl group determination

The carboxyl group content, quantified by the fast method, was compared with results originating from

31

P-NMR and the classic non-aqueous potentiometric titration method using

Tetra-n-butylammonium hydroxide (TnBAH).7 In the potentiometric method (TnBAH method), 175 mg of lignin and 25 mg of hydroxybenzoic acid were dissolved in 30 mL of DMF with constant agitation during 5 minutes. After lignin dissolution, the solution was titrated using TnBAH 0.05M in isopropanol under N2 flow. Two inflexion points appear during the titration, the first point corresponded to the carboxyl content and the second to the total weak acids (COOH + ArOH). The use of the fast method would help to avoid the use of DMF and the necessity of a special electrode for the measurements.

Statistical analysis

Multivariable analysis was carried out to compare the methods for phenolic hydroxyl group determination in lignin samples by applying paired two-sided t-test at 95% confidence level. The analysis was developed using STATGRAPHICS Centurion XV 15.2.06 version.

Results and discussion

Lignin samples structural characterisation

The lignin samples were analysed by ATR-IR to observe its structural composition and the effect of the extraction process used and the provenance of the raw material. Figure S1 shows the


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spectra of the four technical lignin samples used in this study. All the spectra had a similar profile, with slight bands intensity variation. The similarity makes difficult the comparison of the different lignin structures and their possible relation with the raw material and extraction method carried out for lignin recovery. A detailed analysis of the spectra is presented in the Supplementary information.

Chemical composition

Table 1, presenting the chemical composition of lignin samples, gives structural information and provides the level of purity for each sample. All the investigated samples presented total lignin content around 90%, but it is important to highlight the percentage of ashes and sugars indicative of the impurities level. High percentages of sugars and ashes could disturb the phenolic hydroxyl group determination depending of the applied method. On this sense, Organosolv lignin presented the highest sugar contamination due to the lighter process used for its extraction. Indulin lignin from industrial Kraft process presented the highest ashes content, possibly due to the use of minerals for its recovery, whereas, Kraft lignin extracted at CTP (Centre Technique du Papier, Grenoble, France) from pine Kraft industrial black liquor showed the lowest ashes value due to successive washes for impurities removal. All these impurities/interferences can be eluded using the fast method proposed in this study because it is a non-sensitive method to lignin impurities.

Table 1 Chemical composition of lignin samples. Components (%w/w)

PB-Protobind lignin

ORGOrganosolv lignin

KF-Kraft lignin

IND-Indulin lignin


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Total lignin

92.4

89.2

93.7

95.7

Acid insoluble lignin

89.5 ± 1.2

87.8 ± 1.2

92.8 ± 0.3

94.7 ± 0.4

Acid soluble lignin

2.9 ± 0.2

1.4 ± 0.1

0.9 ± 0.2

1.0 ± 0.1

3.1 ± 0.2

7.2 ± 0.3

2.5 ± 0.1

2.6 ± 0.2

Glucose

1.0

4.3

0.2

0.3

Galactose

0.2

0.8

1.6

1.3

Xylose

1.5

1.3

0.2

0.5

Mannose

-

0.2

-

-

Arabinose

0.4

0.6

0.5

0.5

1.6 ± 0.1

1.2 ± 0.1

0.3 ± 0.1

2.9 ± 0.3

Total sugars

Ash

Fast method for phenolic hydroxyl and carboxyl group determination

A fast and easy method for lignin phenolic hydroxyl and carboxyl group determination has been developed. It is based on the combination of aqueous conductometric titration and acid-base titration based on pH determination. A back titration with HCl after neutralisation of phenolic and carboxyl acid groups is carried out by an excess of sodium hydroxide. Figure 1 shows the three different equivalence points appearing along the conductometric and pH-metric titrations in the analysed lignin samples. V1 is detected by change in the conductometric curve slope from negative to slightly positive and V3 is detected by a sharper increase. They correspond to the end-point titration volume of the hydroxyl ions added in excess (V1) and the start of the hydrochloric acid excess introduced in the medium (V3). Between V1 and V3, phenolic hydroxyl groups first, then carboxyl groups are protonated. To distinguish


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these two kinds of functional groups, a pH-metric titration is done in parallel. The titration of free phenols with pKa’s in a range of 8-11.5 can be obtained by the difference volume between V2, corresponding to pH 7, and V1. The remaining acidic groups, i.e. carboxyl groups, correspond thus to the volume difference between V3 and V2. It should be noticed that the conductivity curve presents no rupture between V3 and V1. Equal conductivities of lignin moieties bearing either carboxyl or phenolic hydroxyl functions make it necessary to use pH detection to discriminate them, because of their different pKas.

Figure 1 Conductometric and acid-base titration curves for lignin samples. All the lignin samples presented a similar profile during the titrations independently of the starting raw material and the process applied to recover the lignin samples.


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However, while the V3 equivalence volume was always rather precisely determined (because most carboxylic acids in lignin have a pKa above 4.5, as reported by Zakis),12 a significant imprecision may come from the V1 equivalence volume determination due to the presence of phenolic components with high pKa values (up to 11.5 in condensed phenolic structures bearing an α-carbon adjacent to the phenolic carbon). Therefore, to estimate the extent of error due to the presence of high pKa phenols, Excel software associated to Visual Basic, based on a knowledge mathematical model, was used to simulate the conditions of lignin titration. The titration model consisted in calculating each species concentration by solving the electroneutrality equation (Equation 4) at each titration step, then calculating the theoretical conductivity of the solution by the Kohlrausch equation (Equation 5) [assuming limiting ion conductivities (S·m2·eq-1) at 25°C: 5.01×10-3 for Na+, 7.63×10-3 for Cl-, 1.98×10-2 for OH-, 3.50×10-2 for H3O+, and 2.00×10-3 for a carboxylic or phenolic ionized lignin moiety]. A detailed description of the equations and model used for numerical calculation is presented in the Supplementary information. ∑5 45 . 65 = ∑5 47 . 67

(4)

σ = ∑: z: . C: . λ: (5) The presented simulations reproduced the titration of 1 g of lignin sample, containing about 4 meq of protonated phenolic hydroxyl groups, dissolved by adding 8 meq of NaOH 1M in a total volume of 500 mL of fresh deionized water. Then titration is carried out by adding 16 mL of HCl 1M as titrant. Medium ionic conductivity and pH are calculated after each titrant addition (0.05 mL), and results are plotted against titrant volume (Figure 2). Four titration cases were studied, in which high, mid and low pKa phenolic components varied in abundance. The extent of the range of phenolic hydroxyl group pKas is due to the linkage


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environment of the aromatic nucleus in the lignin molecule. Some structural examples are briefly presented in Supplementary information. With an electron-attracting group such as a carbonyl in a position on side chain, the pKa is low, such as in vanillin or syringaldehyde, with pKas of the order of 8 or even below. Conversely, a carbon-condensation at the C5 position of the aromatic ring results in an increased pKa, above 11 (electron-donating effect). Indeed, normal pKa values in normal, non-condensed lignin moieties are of the order of 9 to 10, similarly as in vanillic alcohol or guaiacol. Data for titration simulations are summarized in Table 2, where the total equivalent amount of components was kept constant (3.6 meq) but the amount of phenolic hydroxyl groups with different pKas varied in proportion. Table 2 Abundance of components (meq) of low, medium and high pKa values in the titration model. Simulation cases

low-pKa components (pKa of 8.0, 8.5 and 9.0)

mid-pKa components (pKa of 9.5, 10.0 and 10.5)

high-pKa components (pKa of 11.0 and 11.5)

Case 1

mid-pKa components dominant, low-pKa present, high-pKa absent

1.2

2.4

0

Case 2

low-pKa components dominant, mid-pKa present, high-pKa absent

2.4

1.2

0

Case 3

equal number of high, mid and low pKa components

1.2

1.2

1.2

Case 4

mid-pKa components dominant, high and lowpKa minor

0.6

2.4

0.6


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Figure 2 illustrates the different titration cases obtained. The theoretical V1 equivalence volume is 4.40 mL, since 4.40 mmol of OH- remains after the consumption of 3.6 mmol of OHby the initially-protonated phenolic units. It is found at the intercept of best extrapolated straight lines plotted in the different portions of the titration curve, around the V1 volume. In cases 1 and 2, a V1 volume of 4.40 ± 0.05 mL of HCl can be read rather accurately at the straight line intercept. Therefore, the encountered quantification error will be quite small at the V1 volume, of the order of ± 1-2% only. This is indeed caused by the absence of high pKa phenols. In case 3, V1 is found at 4.95 ± 0.05 mL, which leads to 12.5 % underestimation of total phenolic hydroxyl group content. In Case 4, possibly with the highest probability since mid-pKa phenols largely dominate, the error keeps rather low, of about 8%, with a volume of 4.75 ± 0.05 mL of HCl.


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Figure 2 Model titration curves simulating lignin titration in the conditions applied for the real lignin samples. Cases corresponding to Table 2. The curves analysis reflected that the fast method has some inherent weakness for phenolic hydroxyl group determination but not for carboxyl groups. However, the magnitude of error for phenolic hydroxyl groups may not exceed 10-15%, since most extracted lignin samples for commercial applications contain rather low amount of condensed structures. When these structures are present, a rather smooth curvature is observed around the V1 point, as demonstrated on the model graphs. Our analyzed samples (Figure 1) showed a sharp curvature


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characteristic of a low presence of high pKa structures. Therefore, the maximum error could be around 10% in the lignin samples investigated. The main advantage of the fast titration method is that it provides a simple, rapid, cheap and non-sensitive to lignin contamination way for accurate testing of industrial lignin samples, often available in larger amounts than 1 g and presenting contamination by sugar and ashes. Meanwhile, the traditional methods present some limitations which will be exposed in the next section. Conversely, the method may not be as well suited to the generally smaller amounts of lignin samples from extracted pulps at lab scale (like HCl-dioxane extraction), it is not able to quantify the structural lignin units (S, G, and H) and total dissolution of lignin samples is necessary for correct quantification. However, the limitation about the lignin amount is expected to be solved in future works diminishing the samples dilution degree.

Phenolic hydroxyl group determination. Comparison with classical methods

The new method presents a useful way for routine determination of phenolic hydroxyl groups substituting tedious and expensive techniques with important drawbacks previously described in the introduction section. Table 3 shows the obtained results for the analyzed lignin samples using the fast method and the comparison with six classical methods. All the measurements were triplicated and average values and standard deviations are presented.


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Table 3 Method comparison for lignin phenolic hydroxyl group determination (mmol/g).

Method

PBProtobind lignin

ORGKF-Kraft Organosolv lignin lignin

INDIndulin lignin

Fast method

2.4 ± 0.03

1.5 ± 0.03

2.4 ± 0.03

2.7 ± 0.05

UV-method

2.6 ± 0.02

1.7 ± 0.01

2.8 ± 0.01

3.4 ± 0.01

Aminolysis

3.4 ± 0.18

2.4 ± 0.12

4.0 ± 0.07

3.6 ± 0.10

Folin-Ciocalteu

3.6 ± 0.03

1.9 ± 0.04

3.0 ± 0.04

3.4 ± 0.02

31

P-NMR

2.7 ± 0.1

1.3 ± 0.06

3.2 ± 0.16

3.2 ± 0.10

1

H-NMR

1.8 ± 0.13

0.9 ± 0.07

2.7 ± 0.04

3.1 ± 0.23

2.4 ± 0.04

2.0 ± 0.03

4.2 ± 0.06

3.6 ± 0.05

13

C-NMR

Analysing the results, Indulin lignin presented more homogeneous results between all the classical methods, but the fast method provided a lower value. The highest results found using the conventional determinations could be due to the interferences induced by the large sugar (2.6%) and ashes (2.9%) contamination leading to an over-estimation. This interference is not reflected in the fast method values. As far as other lignin samples are concerned, values obtained using the fast method are in the range of those given by the classical methods. Statistical analysis (p-test) was carried out to compare results obtained with the fast and conventional methods (Table 4). The results revealed that the fast method presented significant statistical correlation with

31

P-NMR and UV-method, medium correlation with Folin-Ciocalteu

and 1H-NMR and poor correlation with aminolysis and 13C-NMR methods.


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Table 4 Paired t-test (two-sided p-values) for methods comparison. 31

1

Method

Fast method

UVmethod

Aminolysis

FolinCiocalteu

PNMR

HNMR

Fast method

-

0.0298

0.1235

0.0743

0.0358

0.0878

0.3051

UV-method

-

-

0.1759

0.1812

0.0671

0.0362

0.2539

Aminolysis

-

-

-

0.2417

0.0295

0.1190

0.1159

FolinCiocalteu

-

-

-

-

0.1539

0.3094

0.5777

31

P-NMR

-

-

-

-

-

0.0580

0.1597

1

H-NMR

-

-

-

-

-

-

0.1064

-

-

-

-

-

-

-

13

C-NMR

13

C-NMR

The analysed lignin samples do not contain great amounts of condensed, high pKa structures. Then, the correlation between the fast method and

31

P-NMR and UV-method is even better

because an underestimation not higher than 10% in the results should be considered. The good correlation between the fast method and

31

P-NMR and UV-method is an excellent

result because these methods represent the most recognized in scientific publications for the phenolic hydroxyl group determination of lignin samples. Nowadays,

31

P-NMR could be

considered the most referenced method. The use of phosphorylating agent and internal standard (cholesterol) leads the clear differentiation of hydroxyl (aliphatic and aromatic) and carboxyl groups. At the same time,

31

P-NMR is a non-sensitive method to impurities which are not

reactive after phosphorylation28 however, comparing with the fast method, it is a more tedious and complex determination since it requires a derivatization step consuming time and expensive chemicals, a rapid measurement after phosphorylation and an access to costly NMR equipment. The main benefit of the

31

P-NMR method is that it allows a detailed quantification of syringyl,


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guaiacyl and p-hydroxyphenyl units.24 Figure 3 shows a representative 31P-NMR lignin spectrum and Table 5 the quantification of different types of hydroxyl groups.

Figure 3 PB-Protobind lignin sample 31P-NMR spectrum. Table 5 Detailed hydroxyl group quantification (mmol/g) by 31P-NMR.

Lignin samples

Aliphatic OH

Syringyl OH

Condensed OH

Guaiacyl OH

p-Hydroxy OH

PB-Protobind lignin

1.50 ± 0.07

0.67

0.86

0.82

0.35

ORG-Organosolv lignin

1.23 ± 0.06

0.25

0.39

0.45

0.21

KF-Kraft lignin

1.87 ± 0.09

0.24

1.30

1.50

0.16

IND-Indulin lignin

2.06 ± 0.10

0.20

1.25

1.54

0.21

The results reveal the origin of each lignin sample. Softwood lignin samples (Kraft lignin and Indulin lignin) presented a combination between high amount of guaiacyl units and small amounts of syringyl and p-hydroxyphenyl units. On the other hand, non-wood lignin samples


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from agricultural residues (Organosolv lignin and Protobind lignin) presented similar structures proportion.24 UV-method is a rapid and easy way for phenolic hydroxyl group measurements as the fast determination presented in this work. Also, this method only provides the total amount of phenolic hydroxyl groups without any structural specification. However, the method presents some limitations; especially for the quantification of some phenolic structures bearing more than one phenolic hydroxyl group.10 Another important limitation is obtaining full lignin solubility in the water-dioxane mixture used as solvent at buffered pH 6, which is not straightforward for some samples. For the studied lignin samples, the UV-method led to higher values than the fast method. As it was mentioned, the results are equated if an underestimation of 10% is considered in the fast method measurements. Anyway, the presence of contaminants in the samples could be the reason for the slightly increase in the UV-method values. The medium correlation between the fast method with Folin-Ciocalteu and 1H-NMR analysis is significant with the peculiarities of these methods. The Folin-Ciocalteu method was initially proposed for protein content determination. On this way, the application of this method to lignin samples has not been amply extended and seems to present some deficiencies since the assay also quantifies oxidized structures.29 This probably leads to an overestimation of the phenolic hydroxyl groups in comparison with the fast method (Table 3). Definitely, the Folin-Ciocalteu method should be more deeply studied to verify its adequacy for lignin samples analyses. The very low values obtained with the 1H-NMR in Protobind lignin and Organosolv lignin samples left in evidence the main drawback of this technique. To avoid the sample acetylation, the use of deuterated solvents (D2O) requires extreme care in the sample preparation and total


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Page 24 of 36

dryness in the sample and solvents. These requirements make the method very tedious, time consuming and sometimes lead to underestimated results. Moreover, the use of narrow spectra hinders the spectra integration due to overlapping and lack of resolution.28 Finally, the poorest correlation was found with aminolysis and

13

C-NMR methods caused by

the samples acetylation. Aminolysis presented higher values than fast method for all the analysed lignin samples. In theory, the steric effect due to the presence of methoxyl groups in lignin samples22 should provoke incomplete acetylation reactions, giving rise to underestimated results. In our case, more phenolic hydroxyl groups have been detected. This could be due to the lignin sugar contamination, increasing the phenolic hydroxyl group value or disturbing the aminolysis reaction during the acetylation/deacetylation process. Despite the good resolution of

13

C-NMR

spectra, in case of incomplete acetylation reaction this method is limited. Figure 4 shows a representative

13

C-NMR lignin spectrum where numerous peaks of pyridine residues derived

from the acetylation process can be observed even after several evaporation and lyophilisation steps carried out to remove the acetylation reagent excess. The region between 165-175 ppm revealed the aliphatic and aromatic hydroxyl groups and after peak integration the total hydroxyl groups could be quantified. The poor correlation found between the

13

C-NMR and the fast/reference methods (UV and

31

P-NMR) could be due to the

signal overlapping which makes difficult the exact quantification in the carbonyl region of 13CNMR spectrum.


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Figure 4 KF-Kraft lignin sample 13C-NMR spectrum and detail of the carbonyl zone.

Carboxyl group determination

The fast method is able to provide the phenolic hydroxyl and carboxyl content of lignin samples in a single measurement. Table 6 shows the value of carboxyl groups found in the analysed lignin samples using the fast method and the comparison with

31

P-NMR and non-

aqueous potentiometric titration by TnBAH. The TnBAH method has been introduced as the only method in this work using titration methodology and

31

P-NMR constitutes nowadays a

powerful technique to quantify carboxyl groups in lignin samples.


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Table 6 Method comparison for carboxyl group determination (mmol/g).

Method

PBProtobind lignin

ORGKF-Kraft Organosolv lignin lignin

INDIndulin lignin

Fast method

1.6 ± 0.02

2.3 ± 0.05

1.2 ± 0.04

1.4 ± 0.02

TnBAH method

1.6 ± 0.04

1.3 ± 0.01

1.2 ± 0.03

1.1 ± 0.01

31

1.4 ± 0.07

1.1 ± 0.05

0.9 ± 0.04

0.9 ± 0.04

P-NMR

The results were in agreement with some bibliographic values found in the literature using traditional methods for carboxyl group determination.30 Curiously, the analysis revealed lower values using

31

P-NMR and great similarity between the two titration methods except for the

Organosolv lignin sample. Moreover, the carboxyl content of all the analysed lignin samples was lower than the phenolic hydroxyl group amount, except again for the Organosolv lignin sample. A higher carboxyl content compared to phenolic hydroxyls is possible since Organosolv lignin sample extraction process uses oxidative conditions.

Conclusions

A new method for phenolic hydroxyl and carboxyl group quantification on lignin samples was developed. The method, based on a combination of simple aqueous conductometric titration and acid-base titration based on pH determination, provides a fast, economical and simple way to quantify the phenolic hydroxyl groups without using harmful chemicals or costly equipment. Large amount of lignin sample (more than 1 g) and its total dissolution are required for suitable quantification. Moreover, an error around 10-15% may be found for phenolic hydroxyl group determination if the lignin samples present phenolic structures with high pKa values. In the


26

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analysed technical lignin samples low amount of condensed structures were found leading to a maximum error around 10%. Four technical lignin samples were analysed and results obtained with the fast method were compared with those issued from traditional methods. Statistical analysis demonstrated that the fast method results were comparable with the most significant traditional methods as UV and 31

P-NMR analyses. Nowadays, the

31

P-NMR method is considered the reference method for

phenolic hydroxyl groups in lignin samples. However, the complexity of the NMR method and the high cost of the chemicals and equipment could contribute to the use of the fast method providing similar results in a more simple way. On the contrary, 31P-NMR provides information about the structural lignin units (S, G and H) and the fast method is limited in this aspect. On the other hand, the new method presented higher differences with methods those have important limitations as incomplete acetylation (13C-NMR, aminolysis), overestimation probably due to samples contamination (aminolysis), signal overlapping or resolution lack (1H-NMR) or quantification of oxidized structures (Folin-Ciocalteu). Finally, the new method is able to provide the carboxyl content with high accuracy in the same sample measurement. The comparison with a traditional method as TnBAH potentiometric titration showed great results similarity with the exception of the Organosolv lignin sample which is probably partly oxidized.

Acknowledgements

This work has been partially supported by the PolyNat Carnot Institute (Investissements d’Avenir - grant agreement n°ANR-11-CARN-007-01). Luis Serrano gratefully acknowledges


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Page 28 of 36

support from Spanish MINECO via the concession of a Ramon y Cajal contract (ref. RYC-201517109).

Supporting Information

Supporting information includes additional details for: ATR-IR analysis of lignin samples; methodology about acetylation of lignin samples,

31

P-NMR, 1H-NMR and

13

C-NMR analysis;

equations for the titration model calculations; and the influence of lignin phenolic structures in the pKa range.

References

(1) Ragauskas, A.J.; Beckham, G.T.; Biddy, M.J.; Chandra, R.; Cheng, F.; Davis, M.F.; Davison, B.H.; Dixon, R.A.; Gilna, P.; Keller, M.; Lagan, P.; Naskar, A.K.; Saddler, J.N.; Tschaplinski, T.J.; Tuskan, G.A.; Wyman, C.E. Lignin valorization: improving lignin processing in the biorefinery. Science 2014, 344, 1-10. (2) Tolbert, A.; Akinosho, H.; Khunsupat, R.; Naskar, A.; Ragauskas, A.J. Characterization and analysis of the molecular weight of lignin for biorefining studies. Biofuels, Bioprod. Bioref. 2014, 8, 836–856. (3) Buranov, A.U.; Mazza, G. Lignin in straw of herbaceous crops. Ind. Crops Prod. 2008, 28, 237‒259. (4) Hatakeyama, H.; Hatakeyama, T. Lignin structure, properties and applications. Adv. Polym. Sci. 2010, 232, 1−63.


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(5) Capanema, E.A.; Balakshin, M.Y.; Kadla, J.F. A comprehensive approach for quantitative lignin characterization by NMR spectroscopy. J. Agr. Food Chem. 2004, 52, 1850−1860. (6) Naseem, A.; Shaiza, T.; Zia, K.M.; Zuber, M.; Ali, M.; Noreen, A. Lignin-derivatives based polymers, blends and composites: A review. Int. J. Biol. Macromol. 2016, 93, 296–313. (7) Dence C.W.; Lin, S.Y. Methods in lignin chemistry; Springer, Berlin, Heidelberg, 1992. (8) El Mansouri, N.; Salvado, J. Structural characterization of technical lignins for the production of adhesives: Application to lignosulfonate, kraft, soda-anthraquinone, organosolv and ethanol process lignins. Ind. Crops Prod. 2006, 24, 8-16. (9) Månsson, P. Quantitative determination of phenolic and total hydroxyl groups in lignins. Holzforschung 1983, 37, 143-146. (10) Tianinen, E.; Drakenberg, T.; Tamminen, T.; Kataja, K.; Hase, A. Determination of phenolic hydroxyl groups in lignin by combined use of 1H NMR and UV Spectroscopy. Holzforschung 1999, 53, 529–583. (11) Faix, O.; Argyropoulos, D.S.; Robert, D.; Neirinck, V. Determination of hydroxyl groups in lignins evaluation of 1H-,

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1994, 48, 387–394. (12) Zakis, G.L. Functional analysis of lignins and their derivatives; Tappi Press, Atlanta, 1994. (13) Tamminen, T.; Hortling, B. Isolation and characterization of residual lignin. In Progress in lignocellulosics characterization; Ed. Argyropoulos, D., TAPPI Press, 1999.


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(14) De Sousa, F.; Reimann, A.; Björklund, M.; Nivelbrant, J.; Nivelbrant, N.O. Estimating the amount of phenolic hydroxyl groups in lignin. 11th ISWPC Nice, Conference Proceedings Vol. III, 2001, pp 649-653. (15) Folin, O.; Ciocalteu, V. On tyrosine and tryptophane determinations in proteins. J. Biol. Chem. 1927, 73, 627–650. (16) Månsson, P., Selective deacetylation of aromatic acetates by aminolysis. Tetrahedron Lett. 1982, 23, 1845−1846. (17) Wegener, G.; Strobel, C. Bestimmung der phenolischen Hydroxylgruppen in Ligninen und Ligninfraktionen durch Aminolyse und FTIR-Spektroskopie. Holz als Roh- und Werkstoff. 1992, 50, 417–420. (18) Lenz, B. 1968. Application of nuclear magnetic resonance spectroscopy to characterisation of lignin. Tappi 51, 511–519. (19) Lundquist, K. 1979. NMR-studies of lignins 2. Interpretation of the 1HNMR spectra of acetylated birch lignin. Acta Chem.Scand. B33, 27–30. (20) Robert, D. R. and G. Brunow. 1984. Quantitative estimation of hydroxyl groups in milled wood lignin from spruce and in dehydrogenation polymer from coniferyl alcohol using 13C-NMR spectroscopy. Holzforschung 38, 85 –90. (21) Granata, A.; Argyropoulos, D.S. 2-Chloro- 4,4,5,5-tetramethyl-1,3,2-dioxaphospholane, a reagent for the accurate determination of the uncondensed and condensed phenolic moieties in lignins. J. Agr. Food. Chem. 1995, 43, 1538-1544.


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(30) El Mansouri, N.E. Analytical methods for determining functional groups in various technical lignins. Ind. Crops Prod. 2007, 26, 116-124.


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Figure 1. Conductometric and acid-base titration curves for lignin samples. 144x109mm (150 x 150 DPI)


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Figure 2. Model titration curves simulating lignin titration in the conditions applied for the real lignin samples. Cases corresponding to Table 2. 254x190mm (96 x 96 DPI)


Page 34 of 36

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Figure 3. PB-Protobind lignin sample 31P-NMR spectrum. 81x50mm (300 x 300 DPI)


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Figure 4. KF-Kraft lignin sample 13C-NMR spectrum and detail of the carbonyl zone. 84x50mm (300 x 300 DPI)


Page 36 of 36

Fast, Easy, and Economical Quantification of Lignin Phenolic Hydroxyl

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