Modification of Kraft Lignin to Expose Diazobenzene Groups: Toward

Article pubs.acs.org/Biomac

Modification of Kraft Lignin to Expose Diazobenzene Groups: Toward pH- and Light-Responsive Biobased Polymers Antoine Duval,†,‡ Heiko Lange,‡ Martin Lawoko,*,† and Claudia Crestini*,‡ ‡

Department of Chemical Sciences and Technologies, University of Rome “Tor Vergata”, Via della Ricerca Scientifica 1, 00133 Rome, Italy † Wallenberg Wood Science Center (WWSC), Department of Fiber and Polymer Technology, KTH Royal Institute of Technology, SE-100 44 Stockholm, Sweden S Supporting Information *

ABSTRACT: A pH- and light-responsive polymer has been synthesized from softwood kraft lignin by a two-step strategy that aimed to incorporate diazobenzene groups. Initially, styrene oxide was reacted with the phenolic hydroxyl groups in lignin, to offer the attachment of benzene rings, thus creating unhindered reactive sites for further modifications. The use of advanced spectroscopic techniques (1H and 31P NMR, UV and FTIR) demonstrated that the reaction was quantitative and selective toward the phenolic hydroxyl groups. In a second step, the newly incorporated benzene rings were reacted with a diazonium cation to form the target diazobenzene motif, whose formation was again thoroughly verified. As anticipated, the diazobenzene-containing kraft lignin derivatives showed a pH-dependent color change in solution and light-responsive properties resulting from the cis−trans photoisomerization of the diazobenzene group. lacking.18,19 The commercial utilization of technical lignins is limited due to their heterogeneity, multifunctional character, and the lack of structural understanding. Despite these drawbacks, lignin, and in particular kraft lignin, is a multifunctional polymer, displaying many aliphatic and phenolic OH groups, stilbene and aryl enol ether moieties, which offer plenty of opportunities for functionalization once the selective control of each of the reactive groups is achieved. The specific control of the different lignin moieties constitutes both a challenge and huge opportunity for wood chemists to create innovative smart materials. In polymer science, the development of stimuli-responsive polymers attracts much attention for the development of smart materials able to change some of their properties in response to an external stimulus, such as temperature, light, pH or ionic forces. The development of such kind of materials from widely available and low-cost natural resources constitutes a challenge when it comes to bringing them to high added value applications. Recently, poly(N-isopropylacrylamide) (PNIPAM) has been grafted on lignin to confer thermal- and ionic-responsive properties.20−22 Controlled aggregation of lignin with a triblock copolymer of poly(dimethylamino ethyl methacrylate) was exploited to form pH-responsive gels.23 A lignin-based thermal responsive gel has been generated in the form of a mixture of

1. INTRODUCTION Lignin is a naturally abundant biopolymer and the main renewable source of aromatic structures on Earth. It is formed in vivo in vascular plants by the radical polymerization of phenylpropane units, which in softwood almost entirely consist of the guaiacyl type.1−3 The Kraft pulping process generates some 70 Mt yr−1 of the so-called Kraft lignin (KL), which is mostly burnt on site to generate energy and recover the pulping chemicals.4,5 The amount of commercially available KL is only around 100 000 t yr−1.6 Nevertheless, recent development of new industrial processes for KL isolation7,8 and the emergence of forest-based biorefineries9,10 should contribute to bring to the market larger amounts of lignin. New applications are therefore sought to add value to such “technical” lignins. The most investigated routes deal with the depolymerization of lignin to generate aromatic chemicals,11,12 the production of fuels,13 the production of carbon fibers,14 or its incorporation in polymer materials, with or without chemical modifications.15 Despite intense efforts devoted toward the structural characterization of lignin during the past several decades, a clear idea of its molecular structure emerged only recently with the development of advanced heteronuclear 2D NMR techniques.16,17 Lignin presents a wide diversity and variability due to different botanical origins and isolation processes. The structural characterization of technical lignins and more specifically of kraft lignin is a challenging task, and, to date, a general understanding of its molecular structure is still © 2015 American Chemical Society

Received: July 3, 2015 Revised: August 8, 2015 Published: August 19, 2015 2979

DOI: 10.1021/acs.biomac.5b00882 Biomacromolecules 2015, 16, 2979−2989

Article

Biomacromolecules

Table 1. Amounts of Functional Groups of the Functionalized Lignins As Measured by 31P NMR and UV (in mmol g−1) by 31P NMRa,b −1

by UVc

sample name

StOx eq (mol mol )

aliphatic OH

COOH

condensed phenolic OH

uncondensed phenolic OH

total phenolic OH

total phenolic OH

KL KL_StOx0 KL_StOx0.25 KL_StOx0.5 KL_StOx0.75 KL_StOx1 KL_StOx1.3

0 0.25 0.5 0.75 1 1.3

2.00 1.41 2.05 2.66 3.07 3.46 3.76

0.51 0.41 0.42 0.43 0.39 0.36 0.35

2.14 1.52 1.07 0.71 0.48 0.31 0.15

2.47 1.57 1.10 0.54 0.27 0.09 0.04

4.61 3.09 2.17 1.25 0.75 0.40 0.18

3.20 2.86 1.72 1.31 0.48 0.00 0.00

a

Spectra available as Supporting Information (Figure S3). bAnalysis based on literature-known integral ranges.31 cAverages and standard deviations of three measurements. The reaction mixture was subsequently acidified by the addition of 10% (v/v) aqueous HCl and the lignin was isolated as described above under 2.3 until the point at which the sample was freeze-dried. Instead of freeze-drying immediately, the lignin sample was subsequently dispersed in 10 mL of water, and the suspension was divided equally into two parts. One part was subsequently freeze-dried, and the other part was placed in a dialysis tube exhibiting a cutoff between 100 and 500 Da. The filled dialysis tube was subsequently stirred for 12 h in water at pH 7, acidified water at pH 6, and acidic water at pH 2. The lignin-containing emulsion was then isolated using a centrifuge and freeze-dried. 2.4. Lignin Benzylation. Five hundred milligrams of KL were dissolved in water containing 100 mg of NaOH (2.5 mmol, corresponding to 1 equiv to total acidic groups in KL, i.e., phenolic OH and COOH). After 1 h of stirring, styrene oxide was added (from 0.25 to 1.3 equiv to KL phenolic OH), and the reaction mixture was stirred at 50 °C overnight. After cooling to room temperature and acidifying to pH 2 using 10% (v/v) aqueous HCl solution, the suspension was centrifuged to recover the precipitated lignin. The functionalized lignin was then washed three times with 50 mL acidic water (pH 2) and freeze-dried. 2.5. Azolignin Synthesis. Two hundred milligrams of benzylated lignin were dissolved in 5 mL of aqueous NaOH (2 M). N,N-dimethylp-phenylenediamine (1 equiv to benzene groups attached to the lignin) was dissolved in 5 mL aqueous HCl (2 M) in an ice bath. NaNO2 (1 equiv to N,N-dimethyl-p-phenylenediamine) was dissolved in 2.5 mL of water, and the solution was added dropwise to the diamine solution. The solution turned brown, indicating the formation of the diazonium cation. This mixture was then added dropwise to the lignin solution previously placed in an ice/water bath at 0 °C. The reaction was allowed to stir for 2 h. The reaction mixture was subsequently acidified by the addition of 10% (v/v) aqueous HCl and centrifuged to recover the precipitated lignin. Part of the lignin was recovered at pH 6, and the reaction mixture was further acidified to pH 2 to precipitate the remaining lignin. The functionalized lignins were then washed 3 times with 50 mL of acidified water (adjusted at pH 2 or pH 6, respectively) and freeze-dried. 2.6. 31P NMR Analysis. An accurately weighed amount of lignin (about 30 mg) was dissolved in 400 μL of anhydrous CDCl3/pyridine solution (1:1.6 v/v). One hundred microliters of a standard solution of cholesterol (0.1 M in anhydrous CDCl3/pyridine solution) containing Cr(III) acetylacetonate as relaxation agent was then added. Finally, 100 μL of 2-chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane (ClTMDP, 95%, Sigma-Aldrich) was added, and the mixture was stirred at room temperature for 2 h. The mixture was then transferred into 5 mm NMR tubes and the spectra were measured on a Bruker 300 MHz spectrophotometer (256 scans at 20 °C). All chemical shifts reported are relative to the reaction product of water with Cl-TMDP, which gives a sharp signal in pyridine/CDCl3 at 132.2 ppm. Quantitative analysis was performed based on previous literature reports.31 2.7. 1H NMR Analysis. An accurately weighed amount of lignin (about 20 mg) was dissolved in 500 μL of DMSO-d6. One hundred microliters of a standard solution of 2,3,4,5,6-pentafluorobenzaldehyde

furan- and maleimide-functionalized lignins that is capable to undergo reversible Diels−Alder reactions.24 In this study, we aim at linking pH- and light-responsive diazobenzene groups at lignin chain ends. Diazobenzene moieties have been widely studied for their pH-sensitivity25 as well as their light-induced trans−cis isomerization26 thus offering attractive possibilities for the creation of materials for, e.g., photoinduced motion27,28 or photoswitches.29,30 In this work a two-step lignin functionalization strategy was developed to introduce the diazobenzene groups at the lignin terminal units. During the developed synthetic approach reactive benzene groups were first linked to the lignin’s phenolic OH groups, and in a second step these were funtionalized by its reaction with a diazonium cation. After an evaluation of the functionalization efficiency by a set of spectroscopic (1H and 31 P NMR, FTIR, UV) and chromatographic (SEC) techniques, the pH- and light-responsivity of the modified lignins were assessed.

2. EXPERIMENTAL SECTION 2.1. Materials. Kraft lignin (KL) from softwood was obtained from the Lignoboost process.7 Prior to use, it was washed with acidified water (pH 2) to remove the remaining salts and dried in a vacuum oven (24h at 40 °C). Its content in functional groups as measured by 31 P NMR is given in Table 1. Styrene oxide, sodium nitrite (NaNO2) and N,N-dimethyl-p-phenylenediamine were purchased from SigmaAldrich and used as received. 2.2. Model Compounds Functionalization. Two reactions were carried out on lignin dimeric models, one blocked phenolic model(1(4-ethoxy-3-methoxyphenyl)-2-(2-methoxyphenoxy)propane-1,3-diol) (1) and one phenolic model (4-(1-hydroxy-2-(2-methoxyphenoxy)ethyl)-2-methoxyphenol) (2). Model compound (0.5 mmol) was dissolved in 3 mL of aqueous NaOH (2 M). N,N-dimethyl-pphenylenediamine (1 equiv) was dissolved in 3 mL aqueous HCl (2 M) in an ice bath. NaNO2 (1 equiv to N,N-dimethyl-p-phenylenediamine) was dissolved in 2.5 mL of water, and the solution was added dropwise to the diamine solution. After 2 h at 0 °C, the reaction mixture was acidified, stirred for 20 min, and then neutralized to pH 6−7. The aqueous phase was extracted using ethyl acetate (3 × 10 mL). The combined organic phases were washed with brine, dried over magnesium sulfate, filtered, and concentrated under reduced pressure. 2.3. Lignin Reaction with Diazonium Ion. Three hundred eighty-three milligrams of KL (corresponding to 1 mmol G-type aromatics) was dissolved in 6 mL of aqueous NaOH (2M). N,Ndimethyl-p-phenylenediamine (1 equiv to G-type aromatics present in the lignin sample) was dissolved in 6 mL aqueous HCl (2M) in an ice bath. NaNO2 (1 equiv to N,N-dimethyl-p-phenylenediamine) was dissolved in 2.5 mL of water, and the solution was added dropwise to the diamine solution. This mixture was then added dropwise to the lignin solution previously placed in an ice/water bath at 0 °C. The reaction was allowed to stir for 2 h. 2980


Article

Biomacromolecules

Figure 1. Structural motifs found in kraft lignin. Nonphenolic β-O-4′ motifs are highlighted in red, phenolic β-O-4′-motifs are highlighted in green. 2.10. Size-Exclusion Chromatography (SEC). SEC was performed on a SECcurity 1260 system (Polymer Standard Services, Mainz, Germany) equipped with a GRAM PreColumn and two GRAM 1000 analytical columns in series (Polymer Standards Services, Mainz, Germany), using dimethyl sulfoxide (DMSO, HPLC grade, Scharlab, Sweden) with 0.5% w/w LiBr (ReagentPlus) as eluent, with a flow rate of 0.5 mL min−1 at 60 °C. The detection was performed by a refractive index detector and a UV detector set at 280 nm. The lignin fractions were dissolved without derivatization in the SEC eluent at 5 mg mL−1, and filtered through a 0.45 μm PTFE syringe filter prior to injection. Standard calibration was performed using a set of narrow polydispersity pullulan standards of known molecular weights (MW range 342−3.44 × 105 g mol−1). 2.11. GC−MS Analysis. Approximately 1 mg of analyte was dissolved in 1 mL of ethyl acetate. In case the sample was not volatile enough, in situ silylation of the sample was performed by means of addition of 100 μL of pyridine, followed by addition of 50 μL of N,Obis(trimethylsilyl)trifluoroacetamide 60 min prior to analysis. Analysis was done on 5 μL aliquots using a Shimadzu GCMS QP2010 Ultra equipped with an AOi20 autosampler unit. A SLB-5 ms Capillary GC Column (L × I.D. 30 m × 0.32 mm, df 0.50 μm) was used as stationary phase, He as the mobile phase, 100 kPa pressure, 240 °C injection temperature, 200 °C interface temperature; program: 50 °C start temperature for 1 min, 10 °C min−1 heating rate, 240 °C final temperature for 15 min); mass analysis after electron ionization using

in DMSO-d6 was then added, and the mixture was transferred into 5 mm NMR tubes. The spectra were acquired on a Bruker 300 MHz spectrometer (64 scans at 20 °C). 2.8. FTIR Analysis. FTIR spectra were measured on a PerkinElmer 100 FTIR spectrometer. The spectra were acquired on KBr pellets as the average of 32 scans between 450 and 4000 cm−1 with a resolution of 4 cm−1. Spectra were baseline corrected using Spectrum software (PerkinElmer). Prior to quantification, the spectra were normalized with respect to the carbonyl band at 1710 cm−1, which remained unaffected by the chemical modifications carried out in this work, since none of them introduced additional CO groups. 2.9.1. UV−Vis Analysis. Twenty-five microliters of stock solutions of the lignin in DMSO (4 g L−1) were diluted to 2 mL with DMSO to give 0.05 g L−1 solutions. UV−vis spectra were acquired on a Shimadzu dual beam UV-2550 UV−vis spectrophotometer between 250 and 800 nm. To study the pH-dependence of the lignin solutions, 25 μL of stock solutions of the lignin in DMSO (4 g L−1) were diluted to 2 mL with deionized water previously adjusted to pH1 to pH5, and the spectra were recorded in the same conditions. 2.9.2. Phenolic Content by UV Spectroscopy. The phenolic content of the lignins was measured according to the previously reported UV method, which is based on the difference of absorption of the phenolic moieties in neutral solution and the corresponding phenolate anions in alkaline environments.32,33 Spectra were recorded between 200 and 400 nm on a Shimadzu UV-1800 spectrophotometer. 2981


Article

Biomacromolecules Scheme 1. Three-Step Functionalization of Lignin to Expose the pH-Reponsive Diazobenzene Groupsa

a

(a) Benzylation of lignin phenolic OH groups with styrene oxide (highlighted in blue). The structural option (1) is practically not found in mixture of kinetically and thermodynamically opened epoxides; only the structural option (2) is actually detected. (b) Diazotization of N,N-dimethyl-pphenylenediamine. (c) Reaction of diazonium cation with lignin benzene rings yielding the diazobenzene group (highlighted in red).

Figure 2. Reaction of lignin phenolic and nonphenolic model compounds with in situ-generated diazonium ions: (a) blocked phenolic β-O-4′ model (1) and (b) free phenolic β-O-4′ model (2).

2982


Article

Biomacromolecules ca. 70 eV. Analysis was done using Shimadzu GCMS Solution software.

ring as chain end, as depicted in Scheme 1a. The reaction was conducted in NaOH solution, using a stoichiometric amount of NaOH with respect to lignin acidic groups (i.e., phenolic OH and COOH). The applied conditions ensured that all phenolic OH groups would be deprotonated, whereas aliphatic OH’s would remain unaffected because of their higher pKa. The reaction was performed with increasing amounts of StOx, ranging from 0 to 1.3 equiv with respect to lignin phenolic OH. The functional group content of the resulting lignins was monitored by 31P NMR and UV spectroscopy (Table 1). All the 31P NMR spectra are given as Supporting Information (Figure S3). The decrease in total phenolic OH is represented in Figure 3a. The values obtained by both methods

3. RESULTS AND DISCUSSION 3.1. Model Compounds Study. In order to synthesize diazo lignins it is necessary to efficiently functionalize its aromatic rings. In softwood lignins, the majority of aromatic units present are already substituted in at least positions 1, 3 and 4, as shown in Figure 1. This substitution pattern might play a crucial role in the reactivity of lignin toward diazonium ions. In fact, the position para to the free phenolic groups, which is the most activated toward electrophilic substitution, is always functionalized. The only position available for further direct functionalization on the ring itself is thus position 5, which is both ortho to the phenolic or aryl ether moiety and meta to both the methoxy and alkyl substituent (Figure 1). In order to assess the lignin subunit reactivity with diazonium ions, we selected two lignin model compounds carrying the βO-4′ aryl glycerol lignin subunit (Scheme 1 β-O-4′), which is the most abundant interunit bonding motif in lignin. Since the phenolic OH group appeared to be a significant discriminant for reactivity, we selected a nonphenolic model 1 and a phenolic model compound 2, respectively, as shown in Figure 2. Under the alkaline conditions typically used for electrophilic substitution with diazonium ions the nonphenolic model 1 did not react. The phenolic model 2 showed some reactivity yielding traces of the expected product as reported in Figure 2. The reactivity of 2 constitutes a proof of concept that the reaction can indeed occur on the lignin phenolic end groups. The yields obtained in the reaction, however, were not significant given that the major species detected by GC-MS analyses were substrates and deaminated para-dimethylaminoaniline. While β-O-4′ subunits are the most abundant interunit pattern in native lignin, this does not apply to kraft lignin. The alkaline pulping process in fact largely depletes such bonding and leaves a heavily modified polymer; to date the structure of kraft lignin is still a matter of debate. The structure reported in Figure 1 is indicative of the main bondings generated during kraft pulping. In any case, the model served its purpose as model for free and blocked phenols present in all types of lignins. In order to assess the reactivity toward electrophilic aromatic substitution of this material, we proceeded to its treatment with the diazonium ion. When kraft lignin was treated under the same diazotization conditions, 31P NMR and SEC results suggested that no structural changes took place (Figure S1). The 1H NMR spectrum revealed only a very weak signal in the characteristic region for the H atoms ortho to the −NN− bond (7.66 ppm, Figure S2). After dialysis using a dialysis tube with low molecular weight cutoff between 100 and 500 Da, the signal could practically not be detected any more, suggesting that it arose from molecular species smaller than 500 Da. Based on the combined results of the model compound and kraft lignin reactions, it is unlikely that diazo groups can be directly linked to lignin in a significant yield. In order to introduce an activated functional group for reaction with diazonium ions, we thus adopted the strategy to link unfunctionalized and sterically more accessible benzene groups to the lignin backbone. 3.2. Benzylation of Lignin. KL was reacted with styrene oxide (StOx), to expose a nonsubstituted and reactive benzene

Figure 3. Evolution of the phenolic content with the amount of styrene oxide used for functionalization: (a) total phenolic content as measured by 31P NMR and UV; (b) condensed and uncondensed phenolic OH groups as measured by 31P NMR.

correlate well, but the UV method yields slightly lower values than 31P NMR. This is most likely due to the fact that some phenolic structures cannot be determined by the UV method.34,35 In any case, the reaction was found to be quantitative within the accuracy of the determination methods when a slight excess of StOx was used (1.3 equiv), since no more phenolic OH groups could be detected. The reaction is thus more selective than reported for oxypropylation, for which a large excess of propylene oxide (2.5 equiv) is required under similar reaction conditions to achieve a full conversion of phenolic OH groups.36 The reactivity of C5-condensed and noncondensed phenolic units is relatively similar (Figure 3b), even though the full 2983


Article

Biomacromolecules

intermediate stabilized at the benzylic position by the adjacent ring, is thus fully followed, indicating no limitations due to steric hindrance. Figure 4b presents the correlation between the increase in aliphatic OH, resulting from the opening of the epoxide, and the decrease in phenolic OH, caused by the ether formation in course of the linkage of StOx onto lignin. Though the correlation is linear, it does not follow a 1 to 1 ratio, because of the increase in molecular weight associated with the StOx attachment. Since data are expressed in mmol g−1, the apparent decrease in phenolic OH groups is more important than the concomitant increase in aliphatic OH. A loss of some soluble low molecular weight fragments during the workup could also contribute to this difference, since they are known to have the highest phenolic content,38−40 and would also explain the decrease in phenolic OH groups measured between the initial lignin and the blank reaction (KL_StOx0, see Table 1). All the functionalized lignins present similar molecular weight distributions, independent of the total amount of StOx linked (Figure 5). It seems that the loading factor does

conversion of condensed units seems slightly more difficult to achieve, probably due to steric hindrance. As anticipated, carboxyl groups are mainly unaffected by the reaction (Table 1 and Figure 4a). Only a slight decrease is measured, whose significance lies at the limit of the accuracy of the measurement technique.

Figure 5. Molecular weight distributions of the benzylated lignins as a function of the amount of styrene oxide used for functionalization.

Figure 4. (a) Evolution of aliphatic OH and COOH groups with the amount of styrene oxide used for functionalization, (b) correlation between the increase in aliphatic OH (ΔAl−OH) and the decrease in phenolic OH (Δϕ−OH) as a result of the functionalization.

not influence the hydrodynamic volume in a detectable way, probably because the chemical structure of the linked molecule is very close to the original lignin (Scheme 1a). It is noteworthy that no additional peak can be seen in the low molecular weight region, confirming that the styrene oxide (M = 120.15 g mol−1) is successfully attached to the lignin and that the potential excess of reagent has been effectively removed during the washing sequence. The functionalized lignins were analyzed by 1H NMR, and all relevant spectra are supplied as Supporting Information (Figure S5). A strong new signal appeared in the region of the aromatic protons, centered at 7.28 ppm, as a result of the 5 new protons belonging to the benzene ring. Nevertheless, since it overlaps with the aromatic protons initially present in the KL, this creates difficulties and limitations for quantification. Another new signal appearing at 5.20 ppm is assigned to the newly formed benzylic proton (Scheme 1a) and is thus an additional evidence indicative for the covalent bonding of the benzyl group. FTIR spectra of the functionalized lignins, presented in Figure 6a, also confirm the successful chemical transformation. The shape of the OH band between 3100 and 3600 cm−1 was

As depicted in Scheme 1a, the opening of the epoxy ring of StOx during the reaction produces new aliphatic OH groups: this can clearly be evidenced in the 31P NMR spectra, in which the aliphatic OH peak centered around 147.2 ppm increases markedly (Figure S3). Figure 4a shows the increase in aliphatic OH groups with respect to the amount of StOx used for functionalization. It was, however, not possible to determine by this method at which position the epoxide preferentially opens, i.e., which one of the forms (1 or 2) presented in Scheme 1a predominates. To assess the regioselectivity of the reaction, selected samples were functionalized with 2-chloro-1,3,2dioxaphospholane and submitted to 31P NMR, as this reagent allows discriminating primary from secondary aliphatic OH groups.37 The spectra are given as Supporting Information (Figure S4). The content in secondary aliphatic OH groups is constant, whereas an increase in primary aliphatic OH groups is measured, with the growing of a new signal at 133.7 ppm, indicating that only form 2 is produced under these reaction conditions. The electronically favored pathway, with the 2984


Article

Biomacromolecules

diazonium cation onto the benzene ring as depicted on Scheme 1. N,N-Dimethyl-p-phenylenediamine was selected to create, at the end of the lignin chain, a similar motif as in the commonly used pH indicator methyl yellow (4-dimethylaminoazobenzene). Several reactions were performed, starting from either a partially (KL_StOx0.5) or a fully benzylated lignin (KL_StOx1.3). Each time, 1 equiv of N,N-dimethyl-p-phenylenediamine was used with respect to the benzene ring loading factor. A reaction was also performed in a one-pot fashion, without isolating the lignin after the benzylation step. At the end, the reaction mixture was acidified to precipitate the lignin derivatives. Some material was recovered at pH 6, and the mixture was further acidified to pH 2, before the rest of the material was recovered. In some cases, the amount of precipitated material at pH 6 was negligible, and its characteristics are thus not provided (Table 2). Table 2. Characteristics of the Diazobenzene Lignins sample name

pH of precipitation

xdiazoa

ndiazob (mmol g−1)

KL1_StOx1.3_Diazo KL1_Diazo_1-pot KL1_StOx0.5_Diazo_pH2 KL1_StOx0.5_Diazo_pH6

2 2 2 6

0.24 0.29 0.38 0.76

0.57−0.70 0.68−0.84 0.48−0.70 0.95−1.40

a Conversion of benzene groups into diazobenzene, calculated from eq 1. bAmount of diazobenzene motifs in mmol g−1, calculated from eq 2 considering a content in initial benzene groups calculated from either Δ(aliphatic OH) or Δ(total phenolic OH) (Table 1).

Figure 7 presents the 1H NMR spectra of the lignins after the formation of the diazobenzene group. The strong new signal at 3.0 ppm is related to the 6 protons from the two methyl groups carried by the terminal amine group (Figure 7c). Two distinct doublets are discernible in the aromatic protons regions, at respectively 7.66 and 6.77 ppm. They are assigned to the four protons of the newly attached aromatic ring (Figure 7c) and thus demonstrate the formation of the −NN− bridge. Nevertheless, since they strongly overlap with the other aromatic protons coming from the lignin and the linked benzyl group, the quantification of the formation of the diazo linkage by 1H NMR is impossible. The samples were further examined by FTIR spectroscopy. The spectra are presented in Figure 8. The peak at 1600 cm−1, commonly assigned to the aromatic skeleton of lignin,41 increases strongly after the formation of the diazo linkage, possibly because of a contribution of the −NN− vibration at 1575 cm−1.42 Important information can be obtained once more by the study of the fingerprint region. As a result of the formation of the diazo linkage, the initially monosubstituted benzene rings become para-disubstituted (see Scheme 1c). The two typical peaks of monosubstituted benzene rings at 700 and 760 cm−1, respectively, are thus supposed to be replaced by a unique peak around 825 cm−1.42,40 The remaining peak at 700 cm−1, visible in Figure 8a and to a lesser extent oin Figure 8b, is evidence of the incomplete conversion of benzenes into diazobenzenes. The new peak that appears at 822 cm−1 is, on the contrary, an unequivocal indicator for the formation of the diazo linkage. Based on these observations, it is possible to deduce from the FTIR spectra the amount of benzene rings converted into diazobenzene, by the formula

Figure 6. (a) FTIR spectra of the benzylated lignins as a function of the amount of styrene oxide used for functionalization. (b) Evolution of the height of the peaks at 700 and 760 cm−1 with the amount of styrene oxide used.

apparently modified because of the conversion of phenolic OH’s into aliphatic OH’s. The C−H peak at 2940 cm−1 was seen to markedly increase because of the new methanediyl (CH2) and methanetriyl (CH) groups formed by the incorporation of StOx via the opening of the epoxy ring (Scheme 1a). The most obvious changes took place in the fingerprint region, where two sharp signals, typical for monosubstituted benzene rings, appeared at 700 and 760 cm−1.40 Their intensity is well correlated to the amount of StOx used for the functionalization (Figure 6b). The linkage of free and reactive benzene rings onto lignin phenolic OH has thus been successfully realized under mild conditions, with a good selectivity toward phenolic OH, as shown using complementary spectroscopic and chromatographic techniques. The possibility to modulate the benzyl loading on lignin allowed a subsequent functionalization aimed at creating a stimuli-responsive lignin derivative. 3.3. Formation of the Diazobenzene Groups. The incorporation of a diazobenzene group in softwood kraft lignin was then carried out by the electrophilic substitution of a 2985


Article

Biomacromolecules

Figure 7. 1H NMR of the diazobenzene lignins, with an inset of the aromatic protons region (6−8 ppm): (a) fully benzylated lignin (KL_StOx1.3) and subsequent diazobenzene lignins obtained in a stepwise and in a one-pot procedure and (b) partially benzylated lignin (KL_StOx0.5) and subsequent diazobenzene lignins. The signal assignment for the new aromatic protons is given in panel c.

xdiazo =

I822 I822 + I700

(1)

where I700 and I822 represent the relative intensities of the peaks located at 700 and 822 cm−1. The amount of diazobenzene moieties can then be obtained from the initial content in benzene groups nbenzene: ndiazo(mmol ·g −1) = xdiazo × nbenzene(mmol · g −1)

(2)

The results are tabulated in Table 2. The lowest conversion yield is obtained for the sample prepared from the fully benzylated lignin. This low reactivity is most likely related to a limited solubility of the fully benzylated lignin in the NaOH solution used for the reaction. Several attempts using different reaction pH or incorporating DMF as a cosolvent43 failed to give higher conversion yields. The enumerated solubility issue can be partially solved by performing the reaction in one-pot: the lignin is not isolated after the benzylation, but is left in the alkaline solution to which the diazonium cation is added later. In addition to a simplified workup, this one-pot procedure allows one to increase the conversion yield by about 5%. The highest conversion yields have been achieved when a partially benzylated lignin has been used as substrate for the formation of the diazobenzene. Indeed, the partially benzylated lignin still possesses free phenolic OH groups, which, under alkaline conditions, will be deprotonated and thus favor the solubility in form of phenolates. Under these conditions, more than 75% of the benzene rings can be converted into diazobenzene. The total amount of diazobenzene groups thus incorporated within the lignin can finally be higher than when

Figure 8. Detailed FTIR spectra of the diazobenzene lignins in the 450−2000 cm−1 range: (a) fully benzylated lignin (KL_StOx1.3) and subsequent diazobenzene lignins and (b) partially benzylated lignin (KL_StOx0.5) and subsequent diazobenzene lignins. The full spectra (450−4000 cm−1) are available as Supporting Information (Figure S6).

2986


Article

Biomacromolecules

Figure 9. UV−vis spectra measured in DMSO (c = 0.02 g L−1): (a) fully benzylated lignin (KL_StOx1.3) and subsequent diazobenzene lignins, (b) partially benzylated lignin (KL_StOx0.5) and subsequent diazobenzene lignins, and (c) differential spectra of the diazobenzene lignins.

Figure 10. (a) Aqueous solutions (c = 0.2 g L−1) of KL_StOx0.5_Diazo_pH6 at different pH, (b) UV−vis spectra of the same (c = 0.05 g L−1), and (c) the starting KL under identical conditions.

working on a fully benzylated lignin, with the added option to introduce further functional groups on the remaining phenolics as far as needed for envisaged applications. UV−vis absorbance spectra of the samples have been measured in DMSO. DMSO has been chosen for solubility reasons, despite a relatively high wavelength cutoff, which, however, does not affect the measurement on the most interesting range (350−600 nm). The spectra are presented in Figure 9a,b. A strong increase in the absorbance between 350

and 550 nm is obvious for all samples. In order to isolate the contribution of the diazobenzene groups on the absorbance, differential absorbance spectra are presented oin Figure 9c. They were obtained by subtracting the absorbance of the benzylated lignin to the corresponding diazobenzene lignin. All samples exhibit similar spectra, with a maximum at 456 nm and a shoulder at 425 nm, corresponding to the π−π* and the n−π* electronic transitions, respectively. The overlap of the 2987


Article

Biomacromolecules two absorption bands has already been reported for diazobenzene compounds with similar substitutions.44 3.4. Stimuli-Responsive Properties of the Functionalized Lignins. The synthetic efforts outlined so far were motivated by a couple of facts: functionalized biopolymers are of increasing interest for use in smart materials; often natural polymers, most often proteins, are thereby functionalized by attachment of functionalized synthetic polymers;45 natural polyphenolic antioxidants are used in the construction of smart materials;46 and diazo motifs are often used exploiting both their pH-responsive as well as their UV-responsive characteristics.47,48 In order to solely demonstrate that the diazobenzenefunctionalized lignin can potentially play the role of diazo motif-containing polymers while maintaining its characteristics as a natural polyphenol, i.e., so that the diazo-motif also shows effects when present in lower loadings that maintain phenolic OH groups untouched and thus reactive for additional functionalizations, we performed experiments targeting pH and UV-sensitivity of the newly generated functionalized lignin derivatives. pH-Responses. The diazobenzene motif incorporated on the lignin is known to be pH-sensitive. It presents the exact same structure with the pH indicator methyl yellow (4-dimethylaminoazobenzene), a compound exhibiting a color change in solution from red to yellow when pH rises above 3. Selected samples were thus examined for their pH-response. The results presented here relate to the sample with the highest content in diazobenzene structure (KL_StOx0.5_Diazo_pH6). Similar data of the samples having the lowest content in diazobenzene (KL_StOx1.3_Diazo) are provided as Supporting Information (Figure S7). Stock solutions in DMSO were diluted with water previously adjusted to a pH within the range of 1 to 5. The modified lignin was soluble over the whole pH range, down to pH 1, whereas the unmodified kraft lignin control always precipitated at the bottom of the vials (Figure 10a,c). A shift from pink to yellow color occurs between pH 2 and 3 (Figure 10a). This is confirmed by the UV−vis spectra, which reveal an important decrease of the peak located at 500 nm and an increase of the absorption around 430 nm. The same trend is also observed for the lignin samples containing less diazobenzene motifs, though the transition is less clearly marked (Figure S8). This reveals that even for a relatively low loading, the diazbenzene motifs confer to the lignin pH-responsive ability. Photoisomerization. After irradiation with UV light (365 nm), some changes can be detected in the UV−vis absorbance spectra, as shown in Figure 11. They originate from the trans− cis photosiomerization of the diazobenzene group,26 which normally causes a strong reduction in the π−π* absorption band and a slight increase in the n−π* absorption band. Interpretation in the current case is, however, difficult due to the overlap of the two bands as discussed above. Nevertheless, the observed change in absorption constitutes an additional proof of the formation of the diazobenzene group; an in-depth investigation of the photoisomerization behavior of the functionalized lignins will be published in due course.

Figure 11. UV−vis spectra of KL_StOx0.5_Diazo_pH6 before and after irradiation of the solution at 365 nm. Similar data for KL_StOx1.3_Diazo are given as Supporting Information (Figure S9).

styrene oxide under mild conditions, to allow the selective incorporation of unhindered benzene rings at the phenolic ends of the lignin chains, via aryl benzyl ether linkages. The reaction was shown to be quantitative when a slight excess of reagent was used, with excellent selectivity toward the phenolic OH. The incorporated benzene groups were then reacted with a diazonium cation to produce a diazobenzene motif exhibiting a similar structure to that found in the pH indicator methyl yellow. Under the selected conditions, up to 75% of the benzene groups were converted into diazobenzene. This conferred to the lignin a pH-sensitivity, visible as a color change in solution when the pH was reduced below 2−3, and confirmed by UV−vis spectroscopy. Upon irradiation at 365 nm, characteristic changes in the UV absorbance were seen to occur as a result of the cis−trans photoisomerization of the diazobenzene group. It is to be noted, however, that the induced photoresponse may be somewhat obscured by the complex and heterogeneous nature of softwood kraft lignin. It is likely that smaller, less heterogeneous and more monodisperse fractions of softwood kraft lignin would respond to the devised scheme more intensely. As such, current efforts in our group are focused at demonstrating the cis−trans isomerism on lignin fragments obtained via novel solvent fractionation techniques.39,49,50 Overall, this work has provided for a first time the framework and a feasibility study that entails to ligninderived stimuli-responsive polymers. More detailed studies of the light-responsive properties, including the cis−trans reverse photoisomerization is currently being conducted in our laboratory in order to expose and technologically exploit the full potential of the described functionalized lignins.

■

ASSOCIATED CONTENT

S Supporting Information *

Supporting Information (Figures S1−S9) include all 31P NMR, 1 H NMR, FTIR and UV−vis spectra, as well as size exclusion chromatograms. The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/ acs.biomac.5b00882.

4. CONCLUSIONS Kraft lignin was chemically modified to incorporate chain ends with the pH-responsive diazobenzene motif. The installation of these groups within the lignin was carried out in two steps. Initially, the lignin’s phenolic OH groups were reacted with

(PDF) 2988


Article

Biomacromolecules

■

(28) Yamada, M.; Kondo, M.; Mamiya, J.; Yu, Y.; Kinoshita, M.; Barrett, C. J.; Ikeda, T. Angew. Chem., Int. Ed. 2008, 47, 4986−4988. (29) Russew, M.-M.; Hecht, S. Adv. Mater. 2010, 22, 3348−3360. (30) Bléger, D.; Schwarz, J.; Brouwer, A. M.; Hecht, S. J. Am. Chem. Soc. 2012, 134, 20597−20600. (31) Granata, A.; Argyropoulos, D. S. J. Agric. Food Chem. 1995, 43, 1538−1544. (32) Zakis, G. F., Ed. Functional Analysis of Lignins and Their Derivatives; Tappi Press: Atlanta, GA, 1994. (33) Gärtner, A.; Gellerstedt, G.; Tamminen, T. Nord. Pulp Pap. Res. J. 1999, 14, 163−170. (34) Tiainen, E.; Drakenberg, T.; Tamminen, T.; Kataja, K.; Hase, A. Holzforschung 1999, 53, 529−533. (35) El Mansouri, N.-E.; Salvadó, J. Ind. Crops Prod. 2007, 26, 116− 124. (36) Sadeghifar, H.; Cui, C.; Argyropoulos, D. S. Ind. Eng. Chem. Res. 2012, 51, 16713−16720. (37) Argyropoulos, D. S. J. Wood Chem. Technol. 1994, 14, 45−63. (38) Mörck, R.; Yoshida, H.; Kringstad, K. P. Holzforschung 1986, 40, 51−60. (39) Sevastyanova, O.; Helander, M.; Chowdhury, S.; Lange, H.; Wedin, H.; Zhang, L.; Ek, M.; Kadla, J. F.; Crestini, C.; Lindström, M. E. J. Appl. Polym. Sci. 2014, 131, 9505−9515. (40) Cui, C.; Sun, R.; Argyropoulos, D. S. ACS Sustainable Chem. Eng. 2014, 2, 959−968. (41) Faix, O. In Methods in Lignin Chemistry; Lin, S. Y., Dence, C. W., Eds.; Springer-Verlag: Berlin-Heidelberg, 1992; pp 83−109. (42) Hesse, M.; Meier, H.; Zeeh, B. Spectroscopic Methods in Organic Chemistry, 2nd ed.; Thieme: Stuttgart, Germany, 2007. (43) Ardeleanu, R.; Airinei, A.; Sacarescu, G.; Sacarescu, L. Eur. Polym. J. 2002, 38, 2265−2270. (44) Georgi, U.; Reichenbach, P.; Oertel, U.; Eng, L. M.; Voit, B. React. Funct. Polym. 2012, 72, 242−251. (45) Cobo, I.; Li, M.; Sumerlin, B. S.; Perrier, S. Nat. Mater. 2015, 14, 143−159. (46) Tang, C.; Amin, D.; Messersmith, P. B.; Anthony, J. E.; Prud’homme, R. K. Langmuir 2015, 31, 3612−3620. (47) Kawamura, A.; Miyata, T. In Encyclopedia of Polymeric Nanomaterials; Kobayashi, S., Müllen, K., Eds.; Springer-Verlag: Berlin-Heidelberg, 2015; pp 1619−1626. (48) Guo, L.; Li, X.; Guo, Z. J. Mater. Sci.: Mater. Electron. 2015, 26, 6837. (49) Cui, C.; Sun, S.; Argyropoulos, D. S. ACS Sustainable Chem. Eng. 2014, 2, 959−968. (50) Duval, A.; Vilaplana, F.; Crestini, C.; Lawoko, M. Holzforschung 2015, DOI: 10.1515/hf-2014-0346.

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]; Tel: +39 067259 4734. *E-mail: [email protected]; Tel: +46 8 7908047. Notes

The authors declare no competing financial interest.

■ ■

ACKNOWLEDGMENTS The Knut and Alice Wallenberg Foundation is acknowledged for funding. REFERENCES

(1) Boerjan, W.; Ralph, J.; Baucher, M. Annu. Rev. Plant Biol. 2003, 54, 519−546. (2) Ralph, J.; Lundquist, K.; Brunow, G.; Lu, F.; Kim, H.; Schatz, P. F.; Marita, J. M.; Hatfield, R. D.; Ralph, S. A.; Christensen, J. H.; Boerjan, W. Phytochem. Rev. 2004, 3, 29−60. (3) Vanholme, R.; Demedts, B.; Morreel, K.; Ralph, J.; Boerjan, W. Plant Physiol. 2010, 153, 895−905. (4) Sixta, H.; Ed. Handbook of Pulp; Wiley-VCH Verlag GmbH: Weinheim, Germany, 2008. (5) Lora, J. In Monomers, Polymers and Composites from Renewable Resources; Belgacem, M. N., Gandini, A., Eds.; Elsevier: Amsterdam, 2008; pp 225−241. (6) Strassberger, Z.; Tanase, S.; Rothenberg, G. RSC Adv. 2014, 4, 25310−25318. (7) Tomani, P. Cellulose Chem. Technol. 2010, 44, 53−58. (8) Tomani, P.; Axegard, P.; Berglin, N.; Lovell, A.; Nordgren, D. Cellulose Chem. Technol. 2011, 45, 533−540. (9) Ragauskas, A. J.; Williams, C. K.; Davison, B. H.; Britovsek, G.; Cairney, J.; Eckert, C. A.; Frederick, W. J.; Hallett, J. P.; Leak, D. J.; Liotta, C. L.; Mielenz, J. R.; Murphy, R.; Templer, R.; Tschaplinski, T. Science 2006, 311, 484−489. (10) Ragauskas, A. J.; Beckham, G. T.; Biddy, M. J.; Chandra, R.; Chen, F.; Davis, M. F.; Davison, B. H.; Dixon, R. A.; Gilna, P.; Keller, M.; Langan, P.; Naskar, A. K.; Saddler, J. N.; Tschaplinski, T. J.; Tuskan, G. A.; Wyman, C. E. Science 2014, 344, 1246843. (11) Pandey, M. P.; Kim, C. S. Chem. Eng. Technol. 2011, 34, 29−41. (12) Chatel, G.; Rogers, R. D. ACS Sustainable Chem. Eng. 2014, 2, 322−339. (13) Azadi, P.; Inderwildi, O. R.; Farnood, R.; King, D. A. Renewable Sustainable Energy Rev. 2013, 21, 506−523. (14) Baker, D. A.; Rials, T. G. J. Appl. Polym. Sci. 2013, 130, 713− 728. (15) Duval, A.; Lawoko, M. React. Funct. Polym. 2014, 85, 78−96. (16) Crestini, C.; Melone, F.; Sette, M.; Saladino, R. Biomacromolecules 2011, 12, 3928−3935. (17) Sette, M.; Wechselberger, R.; Crestini, C. Chem. - Eur. J. 2011, 17, 9529−9535. (18) Crestini, C. Lignin Structure: A revisitation of current paradigms through NMR analysis; 13th European Workshop on Lignocellulosics and Pulp, Conference Proceedings, 2014; pp 59−62. (19) Crestini, C.; Lange, H.; Sette, M. Lignin Structure: reexamination of current paradigms through NMR analysis. NWBC Nordic Wood Biorefinery Conference, Stockholm, 2014; pp 29−32. (20) Feng, Q.; Chen, F.; Wu, H. BioResources 2011, 6, 4942−4952. (21) Kim, Y. S.; Kadla, J. F. Biomacromolecules 2010, 11, 981−988. (22) Gao, G.; Dallmeyer, J. I.; Kadla, J. F. Biomacromolecules 2012, 13, 3602−3610. (23) Gao, G.; Xu, W. Z.; Kadla, J. F. J. Wood Chem. Technol. 2014, 35, 73−90. (24) Duval, A.; Lange, H.; Lawoko, M.; Crestini, C. Green Chem. 2015, DOI: 10.1039/C5GC01319D. (25) Escudero, D.; Trupp, S.; Bussemer, B.; Mohr, G. J.; González, L. J. Chem. Theory Comput. 2011, 7, 1062−1072. (26) Kumar, G. S.; Neckers, D. C. Chem. Rev. 1989, 89, 1915−1925. (27) Natansohn, A.; Rochon, P. Chem. Rev. 2002, 102, 4139−4175. 2989


Modification of Kraft Lignin to Expose Diazobenzene Groups: Toward

Recommend Documents