A Simple Approach to Prepare Carboxycellulose Nanofibers from

Jun 23, 2017 - Citation data is made available by participants in Crossref's Cited-by Linking service. For a more comprehensive list of citations to t...
0 downloads 0 Views 8MB Size
Download PDF
Article pubs.acs.org/Biomac

A Simple Approach to Prepare Carboxycellulose Nanofibers from Untreated Biomass Priyanka R. Sharma, Ritika Joshi, Sunil K. Sharma, and Benjamin S. Hsiao* Department of Chemistry, Stony Brook University, Stony Brook, New York11794-3400, United States S Supporting Information *

ABSTRACT: A simple approach was developed to prepare carboxycellulose nanofibers directly from untreated biomass using nitric acid or nitric acid-sodium nitrite mixtures. Experiments indicated that this approach greatly reduced the need for multichemicals, and offered significant benefits in lowering the consumption of water and electric energy, when compared with conventional multiple-step processes at bench scale (e.g., TEMPO oxidation). Additionally, the effluent produced by this approach could be efficaciously neutralized using base to produce nitrogen-rich salts as fertilizers. TEM measurements of resulting nanofibers from different biomasses, possessed dimensions in the range of 190−370 and 4−5 nm, having PDI = 0.29− 0.38. These nanofibers exhibited lower crystallinity than untreated jute fibers as determined by TEM diffraction, WAXD and 13C CPMAS NMR (e.g., WAXD crystallinity index was ∼35% for nanofibers vs 62% for jute). Nanofibers with low crystallinity were found to be effective for removal of heavy metal ions for drinking water purification.

■

INTRODUCTION Carboxycelluloses are important derivatives of natural cellulose polymers, and they have been widely used in many biomedical applications, such as hemostatic materials and surgical sutures.1−3 Recently, the developments of different methods to produce carboxycelluloses in nanoscale, such as nanofibers or nanospheres, have further expanded their usage in existing and emerging applications, such as water purification,4−7 nanocomposites,8 nanopaper,9 drug delivery,10,11 ultraporous lightweight foams and aerogels,12,13 gas barrier films,14 biomaterials,15 stability enhancers for carbon nanotube dispersions,16 etc. Many other forms of nanocelluloses without carboxyl groups, such as cellulose nanocrystals, microfibrillated cellulose, bacterial nanocellulose, and cellulose nanofibers, have also been extensively studied in the literature.10,17−28 The major features of carboxycellulose nanofibers, which can be referred to as functional nanocelluloses, are two: (i) the nanoscale format results from the existence of building blockscellulose microfibrils29in the cell walls of biomass, rather than by regeneration of dissolved cellulose polymer chains30 requiring energy-intensive processes; (ii) the modification, such as by TEMPO oxidation,31−35 carboxymethylation,36 phosphorylation,37 acetylation,38 and silylation,39 on the © XXXX American Chemical Society

nanocellulose surface introduces negative charges, which not only facilitate nanofiber dispersion in suspensions, but also provide functional sites for utilization (e.g., adsorption) and further chemical reaction. In addition, nanocelluloses can be extracted from any biomass,10,17−28 including underutilized sources, such as grasses, weeds, shrubs, and agricultural waste. Thus, the development of environmentally friendly and low energy means to extract carboxycellulose nanofibers, from low valued biomass, has untapped potential to replace synthetic polymers in many applications, especially water purification.4−7 This is because carboxycellulose nanofibers can offer very large surface area and functional groups, ideally suited as filtration membranes or/and adsorption media for water treatments.4−6 The TEMPO-mediated oxidation method has been demonstrated as one of the most effective and reliable ways to extract carboxycellulose nanofibers from the cellulose component of biomass (i.e., free of lignin and hemicellulose).31−35 The mild reaction conditions in the TEMPO method selectively convert the hydroxyl groups at the C6 position of the anhydroglucose Received: April 17, 2017 Revised: June 19, 2017 Published: June 23, 2017 A

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Table 1. Effects of Reaction Conditions, Concentration of Nitric Acid, Concentration of Sodium Nitrite, Yield, Lignin Content, and Carboxylate Content on Carboxycellulose Nanofibers Extracted from Jute, Bamboo Cellulose, and Soft Spinnifex Samples S.N.

sample

1 2 3

jute fibersb jute fibers 1N treated jute fibers 2N treated jute fibers 3N treated jute fibers jute fibers bamboo cellulosec soft spinifexd

4 5 6 7 8

reaction condition (hr/°C)

conc. of nitric acid (mmol)

conc. of sodium nitrite (mmol)

yield wt (%)a

acid soluble lignin (mg/g)

−COONa (conductometric titration) mmol/g

10/50 12/50 12/50

22.2 22.2 22.2

28 14 14

24/15 63/38 67/40

0.91 0.65

0.91 1.09 1.15

12/50

22.2

14

70/42

0.45

1.07

12/50

22.2

14

67/40

0.38

1.03

24/50 12/50

22.2 22.2

28

33/20 38/35

ND -

0.75 1.28

12/50

22.2

28

36/20

0.38

0.16

a

Yield based on the cellulose content in biomass/yield based on the original weight of untreated biomass. bJute (cellulose content about 60%)87 c Bamboo cellulose (cellulose content about 92%). dSoft spinifex (cellulose content about 55%)75

unit into carboxyl groups, promoting the fibrillation of large cellulose aggregates into nanofibers (cross-sectional dimensions of only a few nanometers), while maintaining long fiber length (submicrons to microns).8,10,29,40 Many studies have been carried out to illustrate the applicability of this method on different biomass systems.41−43 However, the disadvantage of this method is that it consists of multiple-step processes as well as multiple radical generating chemicals (e.g., sodium hypochlorite, sodium bromide and TEMPO reagents) to convert biomass into carboxycellulose nanofibers, thus limiting the sustainability of the approach in large production scale. There are other chemical modification methods that can also produce carboxycellulose nanofibers from cellulose extracted from different biomass. These methods include etherification,44 oxidation,45,46 esterification,47−49 and carboxymethylation.47,48 Similar to the TEMPO approach, these methods are only effective for the cellulose components without much presence of lignin and hemicellulose, and therefore require various pretreatment procedures to extract cellulose from untreated biomass. The typical pretreatment procedures involve solvent treatment,50,51 alkali pretreatment,52,53 bleaching,54,55 and steam/ammonia extraction.56 These procedures are individually different and have their own virtues, depending on the source of biomass.57,19 In some instances, certain extraction procedures, such as bleaching, have to be repeated several times to remove lignin, hemicellulose, and other components from biomass.58 Additionally, subsequent mechanical treatments are sometimes required to fully fibrillate cellulose macroscale fibers into nanofibers. These treatments include homogenization,31 microfluidization,49 and sonication.59 Thus, the typical process of converting untreated biomass into carboxycellulose nanofibers often comprises multiple steps with multiple chemicals, extensive use of water, and electrical energy.31,47,52,53 Oxidation of cellulose at the C6 position using sodium nitrite with combination of nitric acid, sulfuric acid, and phosphoric acid plus sodium nitrate are well known to produce oxidized cellulose fibers, having length and width in the micrometer range.60−64 However, there has been no report until this study that has dealt with the preparation of carboxycellulose nanofibers using the nitric acid/sodium nitrite method. In the current study, a simple approach to prepare carboxycellulose nanofibers directly from untreated biomass (nonwood), such as jute, spinifex grass, and bamboo cellulose, was demonstrated by

using only nitric acid or nitric acid-sodium nitrite mixtures. This approach efficiently reduced the need to consume multiple chemicals, and thus greatly improved the recycling possibility of the used chemicals. The transition from multiple-step to singlestep process also greatly decreased the electrical energy and water consumption needs. Furthermore, the effluent obtained in this approach could be efficaciously neutralized using base, such as sodium hydroxide or potassium hydroxide, to produce nitrogen rich salts that could be used as plant fertilizer. The resulting carboxycellulose nanofibers were found to have excellent adsorption capability to remove heavy metal ions from water. As there is a vast amount of biomass sources currently considered no value or low-valued, these materials will be an ideal resource for extracting effective water purification materials for a wide range of applications. The demonstrated approach thus represents a cost-effective and sustainable pathway to tackle the drinking water challenges for many regions around the world. The carboxylcellulose nanofibers prepared in the current study have exhibited some features notably different from those of cellulose nanocrystals. For example, the prepared nanofibers exhibited relatively lower crystallnity (∼35% measured by WAXD, and confirmed by 13C CPMAS NMR and TEM diffracton measurements) than that of cellulose nanocrystals (crystallinity >60%). In terms of morphology, the extracted carboxylcellulose nanofibers exhibited substantially longer fiber length and higher aspect ratio than those of cellulose nanocrystals. In addition, these carboxylcellulose nanofibers did not exhibit tapering ends as typically observed in cellulose nanocrystals.10

■

EXPERIMENTAL SECTION

Untreated jute fibers (DP of extracted cellulose = 516) were provided by Toptrans Bangladesh Ltd. In Bangladesh; spinifex grass samples were provided by Dr. Darren Martin from the University of Queensland, Australia; bamboo cellulose samples were obtained from Crown Marina Co. Ltd., Thailand. All samples were cut into small pieces having 3−5 cm in length and subsequently washed, but without further treatment. Analytical grade nitric acid (ACS reagent, 65%) and sodium nitrite (ACS reagent ≥97%) were purchased from Sigma-Aldrich; sodium bicarbonate was purchased from Fisher Scientific. All chemicals were used without further purification. Preparation of Carboxycellulose Nanofibers. Carboxycellulose nanofibers were prepared using the following procedures. One gram of untreated biomass sample (i.e., jute, spinifex or bamboo) was placed in a three-neck round-bottom flask, in which 14 mL (22.2 mmol) of B

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

10°min−1). In this measurement, we also measured untreated jute fibers as a reference. Crystallinity index (CI) was calculated by using the XRD amorphous subtraction method, which was outlined by Ruland.66 In this method, CI was calculated as the ratio of the area of crystalline domain to the total area as expressed by the following equation:

nitric acid was subsequently added. When the samples became completely soaked in the acid, 0.96 or 1.96 g of sodium nitrite (14 or 22 mmol) was added to the reaction mixture under continuous stirring (the actual amount of sodium nitrite per experiment is listed in Table 1). It is imperative to note that upon the addition of sodium nitrite, red fumes formed inside the flask. To prevent the visible fumes from escaping, the mouths of the round-bottom flask were sealed with stoppers. The reaction was performed at 50 °C for 12 h and was then quenched by adding 250 mL of distilled water to the beaker. Once the final product equilibrated, the supernatant liquid was discarded to remove the excess acid. This decantation step was performed 2−3 times, depending on the source of biomass, until fibers started to suspend in water. After that, the recovered product was washed with water and centrifuged (5000 × g for 10 min) until the pH value of the upper suspension was above 2.5. The nanofiber suspension was then transferred to a dialysis bag (Spectral/Por, MWCO: 6−8 kDa) and equilibrated for 4−5 days until the conductivity of water reached below 5 μS. As the pH value of the resulting cellulose nanofibers was between 2.8 and 3.0 due to the presence of carboxyl groups (−COOH), the agglomeration of fibers could be clearly seen during dialysis. The second step of bicarbonate treatment was optional. It was carried out to obtain good dispersion of nanofibers in water. In this step, the fiber suspension was treated with 4 wt % sodium bicarbonate (1:10 wt/v%), until pH reached to 7.5, to generate carboxylate groups (−COO−) with ionic charges. The effect of the pH value on the dispersion of nanofibers with carboxyl groups is illustrated in Figure S1 in Supporting Information, which indicates that these nanofibers have a high tendency to aggregate at pH values below 3. Conductometric Titration Method. The carboxyl content in carboxycellulose nanofibers (the ionic form) having carboxylate (COONa) groups was determined by using the conductivity titration method.65 In this method, 0.3 g of dried nanofiber sample was dispersed in 55 mL of distilled water. Subsequently, 5 mL NaCl (0.01 M) was added to the above suspension and stirred for 15 min. The suspension was then set to a pH value in the range of 2.5−3 by adding 0.1 M HCl. A 0.04 M NaOH was added to the suspension at a rate of 0.1 mL/min until pH reached 11 (monitored by a pH meter). The carboxylate content of nanofibers was calculated from the conductivity and pH curves. Fourier Transform Infra-Red Spectrometry (FTIR). A PerkinElmer Spectrum One instrument was used to record the FTIR curves in the transmission mode, between 450 and 4000 cm−1. A total of 6 scans were taken per sample with a resolution of 4 cm−1. The solid samples were recorded in the attenuated total reflectance (ATR) mode. Transmission Electron Microscopy (TEM). TEM studies of carboxycellulose nanofibers obtained from biomass fibers were carried out by a FEI Tecnai G2 Spirit BioTWIN instrument, operated at an accelerating voltage of 120 kV and equipped with a digital camera. The instrument also possessed photographic film capability with goniometer and tilt stage accessories, as well as electron diffraction capability. In typical sample preparation, a 10 μL aliquot sample of 1 mg of 6-carboxycellulose in 10 mL distilled water was deposited on freshly glow discharged carbon coated Cu grids (300 mesh, Ted Pella Inc.), followed by staining with 2 wt % aqueous uranyl acetate solution. Atomic Force Microscopy (AFM). AFM measurements of carboxycellulose nanofibers obtained from untreated biomass samples (e.g., jute fibers) were performed using a Bruker Dimension ICON scanning probe microscope (Bruker Corporation, U.S.A.) equipped with a Bruker OTESPA tip (tip radius (max.) = 10 nm). A 10 μL of 0.005 wt % nanofibers suspension was deposited on the surface of a silica plate, where the air-dried sample was measured in tapping mode. Wide-Angle X-ray Diffraction (WAXD). X-ray diffraction measurements were carried out using a Benchtop Rigaku MiniFlex 600 instrument. The samples were prepared by coating nanofibers on sample holders made of glass. The Cu Kα radiation was generated at 40 kV and 40 mA (λ = 0.154 nm) using a Ni filter. Data collection was carried out using a flat holder in Bragg−Brentano geometry (5−50°;

CI =

(Itotal − Iam) Itotal

(1)

where Itotal is the integrated intensity of the diffraction spectrum, and Iam is the integrated intensity of the amorphous background. In addition, CI was also estimated by using a different method67 with the following equation:

CI =

(I200 − Iam) I200

(2)

where I200 is the intensity of the dominant (200) diffraction peak, and Iam is the intensity of the amorphous peak evaluated as the minimum peak arise between the dominant (200) peak and the secondary (110) peak. 13 C CPMAS NMR. Solid state 13C CPMAS NMR of biomass fibers and resulting carboxycellulose nanofibers were carried out by a Bruker Utrashield 500WB plus (500 MHz) NMR instrument, equipped with a 2.5 mm triple resonance magic angle spinning (MAS) NMR probe, capable of spinning samples up to 35 kHz. The resonance frequency for 13C was 10 000 Hz, and the samples were spun at the magic angle with a speed of 10 kHz. The CI values of the untreated jute fibers and carboxycellulose nanofibers were measured using the following equation:68

CI =

Area between 87 and 93 ppm × 100 Area between 80 and 93 ppm

(3)

where the area between 87 to 93 ppm and 80 to 93 ppm represent the areas of crystalline and amorphous C4 carbon signal of cellulose. Thermogravimetric Analysis (TGA). The thermal stability of untreated jute and resulting carboxycellulose nanofibers was studied by a PerkinElmer STA-6000 (Simultaneous Thermal Analyzer) instrument. Both TGA and differential thermogravimetry (DTG) curves were measured. The samples were run at a heating rate of 10 °C/min in the range of 30−850 °C under continuous nitrogen flow. Degree of Polymerization (DP) Measurements. The prepared carboxycellulose nanofibers were insoluble in organic solvent due to low degrees of substitution. However, it was possible to measure the degree of polymerization (DP) of nanofibers having nonionic functionality (i.e., COOH or carboxyl group) by the viscosity measurement having cupriethlenediamine as solvent (TAPPI T230 om-99 method).69 In this case, we assumed that the TAPPI method could apply to cellulose nanofibers with low degrees of substitution. Specifically, the graph of viscosity (centipoise, cP) versus DP (as illustrated in page 99 of “Wood and Cellulose Science”70) was used to determine the degree of polymerization. Determination of Lignin Content. The acid soluble lignin content in the resulting carboxycellulose nanofibers was measured by the acetyl bromide method.71 In this method, a 20 mg sample was first placed in a screwed-tight centrifuge tube, containing 0.5 mL of 25 wt % acetyl bromide (v/v in glacial acetic acid), and was subsequently incubated at 70 °C for 30 min until the sample was completely digested. After this step, the centrifuge tube was quickly cooled in an ice bath, and 0.9 mL of 2 M NaOH and 0.1 mL of 5 M hydroxylamine hydrochloride were added to the digested sample. Proper precautionary procedures were taken to contain toxic fumes during chemical addition. A desired amount of acetic acid was then added to the sample for complete dissolution of lignin (e.g., 6 mL of 17.4 M acetic acid was added to the above mixture containing untreated jute fiber). The final products were centrifuged at 1400 × g for 5 min, where the supernatant was analyzed for UV absorbance at 280 nm (Figure 2(ii)). In this test, a standard curve was generated using lignin procured from Sigma-Aldrich (lot no. 370957, formula wt. 505.01) to create a C

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

Figure 1. Mechanism of oxidation of cellulose using nitric acid/sodium nitrite. reference UV absorbance spectrum at 280 nm. The chosen molar extinction coefficient (ε) to calculate the content of lignin was 1.1 g−1Lcm−1. A TAPPI method (T222 om-08, “Acid-insoluble lignin in wood and pulp”)72 was used to measure the insoluble lignin content in nanofibers.

acid, concentration of sodium nitrite, yield, lignin content, and the resulting carboxylate content for jute, soft spinnifex and bamboo samples are summarized in Table 1. All recovered nanofibers produced by this approach were in the nonionized form (−COOH), which was subsequently converted into the ionized form (−COONa) by using 4 wt % sodium bicarbonate (NaHCO3). The ionization of the fiber surface allowed these nanofibers to be uniformly dispersed in an aqueous media. It was found that, when the reaction time was 10 h (Table 1, S.N. 1) under the chosen reaction condition, the yield of nanofibers was 15 wt % (whereby other 15 wt % were microfibers) based on original weight of untreated biomass. On the other hand, the yield of nanofibers increased to 38 wt %, when the reaction time increased to 12 h (Table 1, S.N. 2). It is important to note that, based on the starting cellulose content in biomass, the yield obtained for carboxycellulose nanofibers from untreated jute fibers for S.N. 2 was reasonably high (i.e., 63 wt %). The increase in reaction time also had a notable impact on the carboxylate content of carboxycellulose nanofibers, which increased from 0.91 mmol/g to 1.09 mmol/g (S.N. 1, and S.N. 2). In addition, the acid-soluble lignin content in this treatment was found to be 0.91 ± 0.5 mg/g and the insoluble lignin content (“klason lignin”) was 1.2 ± 0.5 mg/g. The elemental analysis of the product indicated that the presence of nitrogen was 0.043%. This might be due to the presence of traces amount of nitrite in the product. We have also checked the effect of pretreatment on the reactivity of jute fibers. The pretreatment involved the use of sodium hydroxide, as a base with different concentration (1N, 2N, and 3N, respectively), to treat the sample before the acid reaction. The base treatment was carried out at room temperature for 24 h. The treated fibers were first washed properly until the pH value of the filtrate became neutral, and then dried in an oven at 60 °C for 24 h. As seen in Table 1, three base-treated fibers (S.N. 3, 4, 5) were treated with a nitric acid and sodium nitrite mixture, under the same reaction conditions (12 h at 50 °C). The yield for carboxycellulose nanofibers obtained from these base-treated jute fibers were all in the range of 40−42 wt % (or 67−70 wt % based on the cellulose content in starting material), which was only slightly higher than the yield from untreated jute fibers. However, a significant decrease in the lignin content in cellulose nanofibers

■

RESULTS AND DISCUSSION Proposed Mechanism of Oxidation of Cellulose using Nitric Acid/Sodium Nitrite. The mechanism of oxidation of cellulose at the C6 position using nitric acid/sodium nitrite is presented in Figure 1. HNO2 is formed by the reaction of sodium nitrite (NaNO2) and nitric acid (HNO3, which acts as a true oxidant). The reaction liberates nitroxonium ions (NO+) in the presence of excess acid. The produced nitroxonum ion can attack the primary hydroxyl group of cellulose at the C6 position and produce aldehyde group via nitrite ester (R− CH2−O−NO), which is an intermediate. In other words, nitrite ester (R−CH2−O−NO) decomposes in acidic medium to generate HNO and aldehyde group at the C6 position on cellulose. The excess nitric acid oxidizes HNO to HNO2 and the oxidation cycle continues. Thus, nitric acid is consumed, as the reaction proceeds and HNO undergoes self-dissociation to N2O (nitrous oxide) and water molecule.73,74 The produced aldehyde group on cellulose, then oxidizes into carboxyl group in the presence of hydronium ion. The high reaction temperature (70 °C) and high acid concentration (22.2 mmol) can assist the complete oxidation of aldehyde group into the carboxyl group. We hypothesize that the presence of nitric acid initiates the fibrillation process of untreated biomass by eliminating noncellulosic components as well as removing some noncrystalline cellulose regions. Very likely, the further interaction of nitric acid and the crystalline cellulose component led to nanofibers of lower crystallinity. The exact mechanism of preparation of nanofibers using nitric acid/sodium nitrite from untreated biomass is still under investigations. Preparation of Carboxycellulose Nanofibers Directly from Untreated Biomass. Our first set of experiments involved the use of untreated jute fibers (10 g), which were either treated by pure nitric acid (22.2 mmol) or by a nitric acid (22.2 mmol) and sodium nitrite (14 or 28 mmol) mixture at 50 °C. The different reaction conditions, concentration of nitric D

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

oxidization method was simply used as a benchmark reference. There are other oxidation methods, possessing less steps than the TEMPO method, but they are still more complicated than the demonstrated method. It was also demonstrated that the nitric acid and sodium nitrite approach used substantially less chemicals (Figures S5 and S7) and water (Figures S7 and S8) than the TEMPO method. The major differences between the new approach and the TEMPO method for fabrication of carboxycellulose nanofibers, based on our bench experiments, are given in the Supporting Information (Table S1). A simple procedure was also used to neutralize the effluent, consisting of degraded cellulose (sugar molecules), byproducts of hemicellulose and lignin, as well as nitrites and nitrates, from the demonstrated process. This procedure is shown in Figure S9. The salt obtained from this treatment was analyzed using Thermo EA1112 elemetal analyzer, and the results indicated that the product contained N = 1.10%; NO3−/NO2− = 5 mg/L, which could be used directly as plant fertilizer. The FTIR spectra of untreated jute fiber in Figure 2(i) shows several distinctive peaks: 3328 cm−1 (OH streching) and 2900 cm−1 (C−H symmetrical streching) in the cellulose unit; 1515

was observed for nanofibers produced from the base-pretreated jute samples. These results confirmed that the base-pretreatment process removed some lignin content. However, the results also indicated that the method of using nitric acid and sodium nitrite was effective to extract carboxycellulose nanofibers from untreated jute, whereby the base-pretreatment step was unnecessary. Typical SEM images of untreated jute fibers (Figure S2) showed their widths are in the range of 20− 40 μm and their lengths are in the order of several micrometers. Very little difference in fiber width was observed after the base treatment. However, the EDS spectra of untreated jute fibers (Figure S3) depicted the presence of many impurities of nonmetal and metal ions, such as magnesium, aluminum, and silica. These impurities could be partially removed after treatment with base, such as sodium hydroxide, potassium hydroxide, etc., as presented in Supporting Information (Figure S4). To prove the practical application and durability of the demonstrated method, we used the untreated jute fibers without any further purification. The present study clearly indicates that nitric acid is as an effective fibrillating and oxidizing agent. However, the role of sodium nitrite was also examined. It was found that, although nitric acid alone could simultaneously fibrillate and oxidize the jute fibers, the yield to produce carboxylcellulose nanofibers was relatively low even after 24 h of reaction (e.g., the COONa contents were found to be 0.75 mmol/g, respectively). The addition of sodium nitrite appeared to play a synergistic role in enhancing the fibrillating and oxidizing ability of nitric acid, where the proposed mechanism is illustrated in Figure 1. Bamboo cellulose was also used to check the versatility of the combined nitric acid/sodium nitrite method to produce carboxycellulose nanofibers. The resulting nanofibers showed a decent yield (35 wt %) with 1.28 mmol/g of carboxylate group. These values were higher than those from untreated jute and similar to those from base-treated jute fibers due to the intrinsic property of bamboo cellulose structure. However, when compared with the initial cellulose content, the yield of bamboo cellulose was lower than those of some untreated jute fiber (S.N. 2) and all base treated jute fibers. The facile ability of the demonstrated method was also tested on soft spinifex grass,75 having a relatively low lignin content and high hemicellulose content. Because of the relatively low cellulose content in spinifex, the final yield obtained was very low (20 wt %). In addition, the degree of oxidation obtained for carboxycellulose from spinifex grass was quite low (the carboxylate content was 0.16 mmol/g). The low degree of oxidation in spinifex grass was probably due to the high hemicellulose content, which could also have reacted with the oxidation reagent. We made a simple calculation to evaluate the cost reduction in producing carboxycellulose nanofibers by the nitric acid and sodium nitrite method, based on bench experiments. The detailed scheme for the calculation is shown in Figure S5. This exercise indicated that the demonstrated approach could offer a very low-cost and sustainable pathway to produce carboxycellulose nanofibers directly from untreated biomass. Additionally, an estimate on the consumption of electrical energy (Figure S5) by the demonstrated approach was carried out and compared with that by the TEMPO method (Figure S6). The calculation on energy consumption based on bench scale experiment indicated that the demonstrated method could be significantly more energy efficient than the existing methods, such as the TEMPO approach. We note that the TEMPO

Figure 2. (i) FTIR spectra and (ii) UV spectra (after treatment with acetyl bromide) of untreated jute fibers and extracted carboxycellulose nanofibers. E

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules cm−1 (CC aromatic symmetrical streching) in the lignin unit; 1739, 1460, 1240, and 810 cm−1 in the xylan and glucomannan of hemicellulose units. It was found that the OH streching peak at 3328 cm−1 in carboxycellulose nanofibers became much broader than that in untreated jute. This is perhaps due to the disturbance in the hydrogen bonding from the modification of C6 primary hydroxy to carboxyl group in cellulose chains.76 In addition, the peak from the C−H stretching at 2900 cm−1 in carboxycellulose was found to decrease, while the COONa peak, at 1588 cm−1, notably increased. This further confirmed the oxidation of the anhydroglucose units at the C6 position. It was interesting to note that the peaks due to hemicellulsoe and lignin units at 1739, 1460, 1240, 810, and 1515 cm−1, all greatly reduced or completely disappeared in carboxycellulose nanofibers, indicating that the treatment of nitric acid and sodium nitrite was effective in removing impurities of hemicellulose and lignin in jute fibers. Figure 2(ii) shows the overlay UV spectra of untreated jute fibers and carboxycellulose nanofibers, after digestion with acetyl bromide, according to the procedure given in the Experimental Section. The UV absorbance measured for jute fibers and carboxycellulsoe nanofibers (0.2 wt %) at 280 nm was 1.6 and 1.1, respectively. These values were used to measure the lignin contents (mg/g) by the molar extinction coefficient (ε = 1.1 g−1Lcm−1 for standard lignin). The lignin contents of untreated jute and extracted carboxycellulose nanofibers using different treatments are illustrated in Table 1. Characterization of Carboxycellulose Nanofibers. Typical TEM images of carboxycellulose nanofibers obtained from untreated jute fiber and from base treated jute fiber are illustrated in Figure 3. Based on these images, the estimated morphological data, such as the average fiber length (L) and average fiber width (D), are shown in Table 2. It was found that all nanofibers possessed similar widths between 4 and 5 nm, but different average lengths (190−370 nm) and polydispersity indexes (PDI: 0.29−0.38). PDI was measured by using the NanoBrook 90 Plus Particle size analyzer (Brookhaven Instruments Corporation). For untreated samples, the nanofiber length had the following order: jute > spinnifex > bamboo. In addition, the base treatment was found to reduce the fiber length. In Figure 3A, the left inset represents the electron diffraction of nanofibers extracted from untreated jute fibers by TEM. The disappearance of discrete diffraction rings/spots depicted the noncrystalline nature (or low crystallinity) of nanofibers, in agreement with the data obtained in WAXD, NMR, and DTG. Figure 3B illustrates the nanofibers extracted from (1N NaOH) base-treated jute fibers, which indicates that the base treatment did not alter the width (4−5 nm) of the fibers, but did reduce the average length (290 ± 40) nm and thus the aspect ratio. Among the three samples (jute, bamboo and spinifex), the nanofibers produced from bamboo were the shortest in length (97 ± 33 nm, Table 3), followed by those produced from spinifex grass (220 ± 30 nm), with produced jute nanofibers being the longest. The typical TEM image of nanofibers from spinifex is shown in Figure S10. All extracted nanofibers exhibited similar width (4−5 nm) and low crystallinity. We believe that, although differing fiber lengths obtained from different biomasses might reflect the intrinsic cellulose structure differences in cell walls, they are more likely due to the chosen reaction conditions (e.g., the reagent concentration and temperature). AFM of carboxycellulose nanofibers obtained from (1N NaOH) base-treated jute fibers is presented in Figure

Figure 3. TEM image of (A) carboxycellulose nanofibers (containing carboxylate group) obtained from untreated jute fibers (inset left: electron diffraction diagram), obtained at magnification 395 000×, (B) carboxycellulose nanofibers (containing carboxylate group) obtained from base treated jute fibers, obtained at magnification 548 000× (ImageJ software was used to calculate average length and average diameter of 20 fibers: (A) average length = 350 ± 20 nm, average width = 4.20 ± 0.5 nm; and (B) average length = 290 ± 40 nm, average width = 4.47 ± 0.5 nm).

Table 2. Morphological Data of Carboxycellulose Nanofibers from Various Sources Based on TEM Measurementsa TEM (nm)

untreated jute fibers 1N treated jute fibers bamboo cellulose spinifex a

(L)

(D)

Aspect ratio (L/D)

PDI

350 ± 20 290 ± 40

4.20 ± 0.5 4.47 ± 0.5

91 77

0.29 0.31

97 ± 33 220 ± 30

4.27 ± 0.4 4.49 ± 0.5

24 56

0.38 0.35

L: average fiber length; D: average fiber width.

4, which further confirmed that the average thickness of obtained nanofibers was around 5 nm. The TEM images presented in Figure 2A and Figure 2B are of nanofibers having F

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules Table 3. Resonance Assignment for Main Peaks in 13C CPMAS NMR Spectra of Jute Fibers and Carboxycellulose Nanofibers chemical shift (ppm) carbon atom

jute fiber

cellulose nanofibers

crystalline C4 amorphous C4 crystalline C6 amorphous C6 reducing end group

89.15 84.47 65.27 62.83 -

89.18 (very small) 83.30 (major) 65.36 62.65 (smaller) 92.94

Figure 4. Non-contact-mode AFM image of carboxycellulose nanofibers (containing carboxylate group) obtained from base-treated jute fibers (corresponding 2D height image). Figure 5. (i) WAXD overlay of untreated jute fibers and carboxycellulose nanofibers; (ii) 13C CPMAS NMR of untreated jute fibers and carboxycellulose nanofibers.

the carboxylate group, while Figure S11 shows the nanofibers having the carboxyl group, which was formed without bicarbonate treatment from 1N NaOH-treated jute fibers. WAXD patterns of untreated jute fibers and extracted carboxycellulose naofibers are illustrated in Figure 5(i). These patterns indicate that both samples exhibited the characteristic of a cellulose I structure, with three peaks located at 2θ angles of 16.5°, 22.7°, 35.1°, corresponding to the (110), (200), and (004) lattice planes, respectively.77,78 However, the (11̅0) and (102) peaks were not visible in the WAXD pattern of untreated jute fibers, in agreement with the published results.78 Most probably, the presence of hemicellulose and lignin molecules in cellulose crystal structure weakens the intermolecular hydrogen bonding plane, leading to the disappearance of the (110̅ ) peak.79 The main peak for the (200) peak at 2θ (22.8°) is indicative of the distance between hydrogen-bonded sheets in cellulose I. The WAXD pattern of carboxycellulose nanofibers (Figure 5(i)B) shows a diffraction pattern having peak positions at 2θ angles of 14.9, 16.7, 22.6 and 34.5°, corresponding to (11̅0), (110), (200) and (004) lattice planes, respectively. The reappearance of the very weak (11̅0) peak in carboxycellulose nanofibers depicts the removal of some hemicellulose and lignin in the cellulose I structure.79 CI calculated from the WAXD data using the Ruland method was 35% for carboxycellulose nanofibers, and 61% for untreated jute fibers. Similarly, CI estimated by using the “Segal equation” was 38% for carboxycellulose nanofibers, and 62% for jute fiber. The estimated CI values for untreated jute fibers were similar to that reported previously,80 while carboxycellulose nanofibers clearly possessed a lower degree of crystallinity.

Solid state 13C CPMAS NMR spectra of untreated jute fibers and carboxycellulose nanofibers are shown in Figure 5(ii). The region between 60 and 70 ppm was assigned to C6 carbon of the primary alcohol group. The next cluster between 70 and 80 ppm could be attributed to the C2, C3, and C5 carbons. The region between 80 and 95 ppm was associated with C4 carbon, and that between 100 and 110 ppm was due to the anomeric carbon C1.81 The NMR spectra of untreated jute fibers (Figure 5(ii)A) exhibited distinct peaks at ∼21 and 56 ppm corresponding to glucoroxylans of hemicellulose. Also, the signals at ∼153 and 171−172 ppm belonged to C1 carbon of guaiacyl units and C4, C3 carbons of syringyl units of lignin.82 The disappearance of distinct lignin and hemicellulose signals in the 13C CPMAS NMR spectra of carboxy cellulose nanofibers confirmed the removal of hemicellulose, lignin, and other component impurities during oxidation by nitric acid/sodium nitrite. The main peaks appearing in 13C CPMAS NMR of carboxycellulose nanofibers represented the assembly of C6, C2,3,5, C4 and C1 carbons in the anhydroglucose units. The resonance assignment for the foremost distinct peaks in the 13C CPMAS NMR spectra of untreated jute fiber and carboxycellulose nanofibers is listed in Table 3. The crystalline and noncrystalline peaks for C4 carbon of untreated jute fiber appeared at ∼89 ppm and ∼84 ppm, respectively,79 with almost equal intensity. However, in carboxycellulose nanofibers, the noncrystalline peaks (∼83 ppm) became the dominant peak, adjacent to the residual small crystalline peak at ∼89 ppm. G

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

curve obtained for carboxycellulose nanofibers exhibited a Tonset degradation temperature at 164 °C with 13 wt % weight loss. The initialization of thermal degradation of nanofibers was probably due to the presence of anhydroglucuronic acid units, consisting of thermally unstable carboxyl groups. However, the Toffset for nanofibers appeared at 531 °C with a weight loss of 99.8 wt %. In Figure 6(ii), the DTG decomposition curve of untreated jute fiber showed the appearance of four peaks, in which the lowest peak appeared at 292 °C, indicating the presence of hemicellulose, while the largest peak appeared at 367 °C indicating the presence of the crystalline region of cellulose.85,86 The other peak of DTG at 500 °C appeared to be due to the degradation of high molecular weight lignin, consisting of highly cross-linked aromatic polymer units.8 The DTG curve of cellulose nanofibers showed four distinct peaks, in which the lowest peak was at 181 °C, corresponding to anhydroglucoronic units. The carboxyl group in anhydroglucoronic units is sensitive to thermal treatment due to facile decarbonization upon heating. Another peak adjacent to the anhydroglucoronic unit ascended to 241 °C, resembling noncrystalline cellulose with a small residual peak of crystalline cellulose at 322 °C, indicating that the prepared nanofibers possessed predominant noncrystalline domains with low crystallinity. The results obtained here are in agreement with the results obtained in a previous study,86 where the crystalline peak shifted and lowered because of a decrease in the degree of polymerization and carboxylation. The presence of DTG curve at 464 °C, confirmed the availability of trace amounts of lignin in nanofibers. Quantitative measurements were carried out using the acetyl bromide method, as depicted in the previous section. Carboxycellulose nanofibers with low crystallinity were found to be effective media for removal of heavy metal ions. A simple experiment demonstrated by addition of uranium acetate solution to nanofibers suspension, lead to an immediate coagulation (Figure S12 in the Supporting Information), enabling pure water to be easily extracted by decanting or gravity-driven microfiltration. This study demonstrated the great potential of using carboxycellulose nanofibers for heavy metal ion removal in varying applications of water purification. The detailed results from this study will be published elsewhere.

Similarly, in the crystalline and amorphous regions, the peak intensity for C6 carbon of untreated jute fiber appeared at ∼65 ppm and ∼62 ppm, respectively. The crystalline peak for C6 carbon of cellulose significantly decreased in nanofibers at ∼62 ppm. Correspondingly, the peak at about ∼92 ppm in nanofibers was assigned to the C1 anomeric carbon at the reducing end of cellulose.83 The above observation indicated a decrease in the degree of polymerization (DP) in nanofibers. The DP value measured by using the method TAPPI T230 om99 method69 for the nanofibers was 176. The solid state 13C CPMAS NMR analysis showed that the prepared carboxycellulose nanofibers possessed mostly noncrystalline region of both C4 and C6 carbon atoms. The CI of untreated jute fiber and carboxycellulose nanofibers were further confirmed by using 13C CPMAS NMR as elucidated in the Experimental Section. CI measured for jute fibers and nanofibers were 62% and 32%, respectively, in agreement with the CI values obtained from WAXD experiments. The thermogravimetric and derivative thermogravimetric analysis of jute fibers and carboxycellulose nanofibers is shown in Figure 6(i). The thermal degradation curve of jute fibers occurred in three major steps at 281 °C, 376 and 520 °C. The initial onset temperature (Tonset) for degradation was observed at 281 °C with 8 wt % weight loss and the offset temperature (Toffset) was about 520 °C. These results were similar to the data in the literature.84 In contrast; the thermal degradation

■

CONCLUSIONS In this study, a simple approach was demonstrated to prepare carboxycellulose nanofibers directly from different untreated biomass sources, such as jute, spinifex grass, and bamboo, using only nitric acid or nitric acid-sodium nitrite mixtures. The results indicate that this approach offers substantial reduction in the consumption of chemicals, water and electrical energy, when compared to conventional multiple-step approaches, such as TEMPO oxidation, to extract carboxylated nanocelluloses from the same source. In addition, the effluent produced with this method can be efficaciously neutralized by base (e.g., sodium hydroxide) to produce nitrogen-rich salts that can be utilized as fertilizers. TEM measurements of carboxycellulose nanofibers extracted from different biomasses all possessed an average length in the range of 190−370 nm, width of 4−5 nm and polydispersity index (PDI) of 0.29−0.38. These nanofibers possessed lower crystallinity than the untreated biomass as determined by TEM diffraction pattern, WAXD and 13C CPMAS NMR analysis. For example, the crystallinity index (CI) measured by WAXD for jute-derived nanofibers was only 35%, when compared with 62% in untreated jute. Produced by

Figure 6. (i) TGA of untreated jute fibers and carboxycellulose nanofibers (ii) DTG of untreated jute fibers and carboxycellulose nanofibers. H

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules

(University High School, Irvine, CA) for testing uranium remediation with carboxycellulose nanofiber suspension.

this simple and low-cost approach from sustainable resources, carboxycellulose nanofibers with low degree crystallinity were found to be particularly effective to remove toxic metal ions, thus they may provide a new pathway to deal with various water pollution challenges.

■

■

(1) Spangler, D.; Rothenburger, S.; Nguyen, K.; Jampani, H.; Weiss, S.; Bhende, S. Surg. Infect. 2003, 4, 255−262. (2) Jeschke, M. G.; Sandmann, G.; Schubert, T.; Klein, D. Wound Repair Regen. 2005, 13, 324−331. (3) Dineen, P. Surg. Gynecol. Obstet. 1976, 142, 481−486. (4) Ma, H. Y.; Burger, C.; Hsiao, B. S.; Chu, B. Biomacromolecules 2012, 13, 180−186. (5) Ma, H. Y.; Hsiao, B. S.; Chu, B. ACS Macro Lett. 2012, 1, 213− 216. (6) Ma, H. Y.; Burger, C.; Hsiao, B. S.; Chu, B. J. Mater. Chem. 2011, 21, 7507−7510. (7) Ma, H. Y.; Burger, C.; Hsiao, B. S.; Chu, B. ACS Macro Lett. 2012, 1, 723−726. (8) Johnson, R. K.; Zink-Sharp, A.; Renneckar, S. H.; Glasser, W. G. Cellulose 2009, 16, 227−238. (9) Olsson, R. T.; Azizi Samir, M. A. S.; Salazar-Alvarez, G.; Belova, L.; Strom, V.; Berglund, L. A.; Ikkala, O.; Nogues, J.; Gedde, U. W. Nat. Nanotechnol. 2010, 5, 584−588. (10) Lin, N.; Huang, J.; Dufresne, A. Nanoscale 2012, 4, 3274−3294. (11) Huang, L.; Chen, X.; Nguyen, T. X.; Tang, H.; Zhang, L.; Yang, G. J. Mater. Chem. B 2013, 1, 2976−2984. (12) Sakai, K.; Kobayashi, Y.; Saito, T.; Isogai, A. Sci. Rep. 2016, 6, 20434. (13) Aulin, C.; Gallstedt, M.; Lindstrom, T. Cellulose 2010, 17, 559− 574. (14) Fukuzumi, H.; Saito, T.; Iwata, T.; Kumamoto, Y.; Isogai, A. Biomacromolecules 2009, 10, 162−165. (15) Koga, H.; Tokunaga, E.; Hidaka, M.; Umemura, Y.; Saito, T.; Isogai, A.; Kitaoka, T. Chem. Commun. 2010, 46, 8567−8569. (16) Sharma, P. R.; Varma, A. J. Chem. Commun. 2013, 49, 8818− 8820. (17) Trache, D.; Hussin, M. H.; Haafiz, M. K.; Thakur, V. K. Nanoscale 2017, 9, 1763−1786. (18) Moon, R. J.; Martini, A.; Nairn, J.; Simonsen, J.; Youngblood, J. Chem. Soc. Rev. 2011, 40, 3941−94. (19) Brinchi, L.; Cotana, F.; Fortunati, E.; Kenny, J. M. Carbohydr. Polym. 2013, 94, 154−69. (20) Lee, H. V.; Hamid, S. B.; Zain, S. K. Sci. World J. 2014, 2014, 631013. (21) Reid, M. S.; Villalobos, M.; Cranston, E. D. Langmuir 2017, 33, 1583−1598. (22) Roy, D.; Semsarilar, M.; Guthrie, J. T.; Perrier, S. Chem. Soc. Rev. 2009, 38, 2046−2064. (23) Isogai, A.; Saito, T.; Fukuzumi, H. Nanoscale 2011, 3, 71−85. (24) Habibi, Y. Chem. Soc. Rev. 2014, 43, 1519−42. (25) Van de Ven, T. G.; Sheikhi, A. Nanoscale 2016, 8, 15101−14. (26) Abdul Khalil, H. P. S.; Bhat, A. H.; Ireana Yusra, A. F. Carbohydr. Polym. 2012, 87, 963−979. (27) Kim, J.-H.; Shim, B. S.; Kim, H. S.; Lee, Y.-J.; Min, S.-K.; Jang, D.; Abas, Z.; Kim, J. Int. J. Pr. Eng. Man.-GT 2015, 2, 197−213. (28) Klemm, D.; Kramer, F.; Moritz, S.; Lindstrom, T.; Ankerfors, M.; Gray, D.; Dorris, A. Angew. Chem., Int. Ed. 2011, 50, 5438−66. (29) Su, Y.; Burger, C.; Ma, H.; Chu, B.; Hsiao, B. S. Biomacromolecules 2015, 16, 1201−1209. (30) Medronho, B.; Lindman, B. Adv. Colloid Interface Sci. 2015, 222, 502−508. (31) Saito, T.; Nishiyama, Y.; Putaux, J. L.; Vignon, M.; Isogai, A. Biomacromolecules 2006, 7, 1687−1691. (32) Fan, Y. M.; Saito, T.; Isogai, A. Biomacromolecules 2008, 9, 192− 198. (33) Saito, T.; Okita, Y.; Nge, T. T.; Sugiyama, J.; Isogai, A. Carbohydr. Polym. 2006, 65, 435−440. (34) Shinoda, R.; Saito, T.; Okita, Y.; Isogai, A. Biomacromolecules 2012, 13, 842−849.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.7b00544. Photographs showing carboxycellulose nanofibers in suspension (0.02 wt %) at different pH; SEM images of untreated jute fibers and base treated jute fibers; EDS spectra of untreated jute fibers and base treated jute fibers; estimates on the materials cost and consumption of energy for fabrication of jute carboxycellulose nanofibers using nitric acid−based on bench experiments; estimates on consumption of energy for fabrication of jute carboxycellulose nanofibers using TEMPO oxidation; estimates on consumption of chemicals for fabrication of jute carboxycellulose nanofibers using nitric acid and sodium nitrite method based on bench experiments; estimates on consumption of chemicals for fabrication of jute carboxycellulose nanofibers using TEMPO oxidation based on bench experiments; estimate on salt obtained from the effluent produced during preparation of carboxycellulose nanofibers using nitric acid and sodium nitrite method; typical TEM Image of carboxycellulose nanofibers (carboxylate form) extracted from untreated spinifex fibers using the nitric acid and nitrite method; typical TEM Image of carboxycellulose nanofibers (carboxyl form-before bicarbonate treatment) extracted from base treated jute fibers using the nitric acid and nitrite method; a table illustrating notable differences between TEMPO and nitric acid−based methods; and a photograph showing remediation efficiency of carboxycellulose nanofibers produced from jute fibers using the nitric acid-sodium nitrite method. (PDF)

■

REFERENCES

AUTHOR INFORMATION

Corresponding Author

*Tel: +16316327793; E-mail: [email protected]. ORCID

Benjamin S. Hsiao: 0000-0002-3180-1826 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors would like to thank the SusChEM Program of the National Science Foundation (DMR-1409507) for financial support. The authors would like to thank Prof. Darren Martin and Dr. Nasim Amiralian (University of Queensland, Australia) for providing spinifex grass samples, Ms. Susan Von Horn (iLab at Stony Brook University) for conducting TEM analysis, Dr. Jim Quinn (Materials Science and Engineering at Stony Brook University) for SEM analysis, and Dr. Martin Ziliox (NMR facility at Stony Brook University) for NMR analysis. Authors would also like to thank Mr. Aurnov Chattopadhyay I

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX

Article

Biomacromolecules (35) da Silva Perez, D.; Montanari, S.; Vignon, M. R. Biomacromolecules 2003, 4, 1417−1425. (36) Naderi, A.; Lindstrom, T.; Pettersson, P. Cellulose 2014, 21, 2357−2368. (37) Kokol, V.; Bozic, M.; Vogrincic; Mathew, A. P. Carbohydr. Polym. 2015, 125, 301−313. (38) Abraham, E.; Kam, D.; Nevo, Y.; Slattegard, R.; Rivkin, A.; Lapidot, S.; Shoseyov, O. ACS Appl. Mater. Interfaces 2016, 8, 28086− 28095. (39) Zhang, Z.; Sebe, G.; Rentsch, D.; Zimmermann, T.; Tingaut, P. Chem. Mater. 2014, 26, 2659−2668. (40) Siro, I.; Plackett, D. Cellulose 2010, 17, 459−494. (41) Saito, T.; Kimura, S.; Nishiyama, Y.; Isogai, A. Biomacromolecules 2007, 8, 2485−2491. (42) Missoum, K.; Belgacem, N.; Bras, J. Materials 2013, 6, 1745− 1766. (43) Abdul Khalil, H. P. S.; Davoudpour, Y.; Islam, Md N.; Mustapha, A.; Sudesh, K.; Dungani, R.; Jawaid, M. Carbohydr. Polym. 2014, 99, 649−665. (44) Knaus, S.; Bauer-Heim, B. Carbohydr. Polym. 2003, 53, 383− 394. (45) Kim, U. J.; Kuga, S.; Wada, M.; Okano, M.; Kondo, T. Biomacromolecules 2000, 1, 488−492. (46) Varma, A. J.; Kulkarni, M. P. Polym. Degrad. Stab. 2002, 77, 25− 27. (47) Naderi, A.; Lindstrom, T.; Sundstrom, J.; Pettersson, T.; Flodberg, G.; Erlandsson, J. Cellulose 2015, 22, 1159−1173. (48) Naderi, A.; Lindstrom, T.; Erlandsson, J.; Sundstrom, J.; Flodberg, G. Nord. Pulp Pap. Res. J. 2016, 31, 364−371. (49) Siró, I.; Plackett, D.; Hedenqvist, M.; Ankerfors, M.; Lindström, T. J. Appl. Polym. Sci. 2011, 119, 2652−2660. (50) Das, S.; Saha, A. K.; Choudhury, P. K.; Basak, R. K.; Mitra, B. C.; Todd, T.; Lang, S.; Rowell, R. M. J. Appl. Polym. Sci. 2000, 76, 1652−1661. (51) Allen, S. G.; Schulman, D.; Lichwa, J.; Antal, M. J.; et al. Ind. Eng. Chem. Res. 2001, 40, 2934−2941. (52) Gassan, J.; Bledzki, A. K. J. Appl. Polym. Sci. 1999, 71, 623−629. (53) Bjerre, A. B.; Olesen, A. B.; Fernqvist, T.; Plöger, A.; Schmidt, A. S. Biotechnol. Bioeng. 1996, 49, 568−577. (54) Mannan, Kh. M.; Talukder, M. A. I. Polymer 1997, 38, 2493− 2500. (55) Salam, A.; Reddy, N.; Yang, Y. Ind. Eng. Chem. Res. 2007, 46, 1452−1458. (56) Kumar, P.; Barrett, D. M.; Delwiche, M. J.; Stroeve, P. Ind. Eng. Chem. Res. 2009, 48, 3713−3729. (57) Moran, J. I.; Alvarez, V. A.; Cyras, V. P.; Vazquez, A. Cellulose 2008, 15, 149−59. (58) Watkins, D.; Nuruddin, Md.; Hosur, M.; Tcherbi-Narteh, A.; Jeelani, S. J. Mater. Res. Technol. 2015, 4, 26−32. (59) Frone, A. N.; Panaitescu, D. M.; Donescu, D.; Spataru, C. I.; Radovici, C.; Trusca, R.; Somoghi, R. BioResorces 2011, 6, 487−512. (60) Gert, E. V.; Shishonok, M. V.; Zubets, O. V.; Torgashov, V. I.; Kaputskii, E. N. Polym. Sci. Series 1995, A37, 670−675. (61) Kumar, V.; Yang, T. Carbohydr. Polym. 2002, 48, 403−412. (62) Kumar, V.; Yang, D. J. J. Biomater. Sci., Polym. Ed. 2002, 13, 273−286. (63) Painter, T. J. Carbohydr. Res. 1977, 55, 95−103. (64) Wanleg, H. U.S. Patent 2,758,112, 1956. (65) Saito, T.; Isogai, A. Biomacromolecules 2004, 5, 1983−1989. (66) Ruland, W. Acta Crystallogr. 1961, 14, 1180−1185. (67) Segal, L.; Creely, J.; Martin, A. E. J.; Conrad, C. M. Text. Res. J. 1959, 29, 786−794. (68) Park, S.; Baker, J. O.; Himmel, M. E.; Parilla, P. A.; Johnson, D. K. Biotechnol. Biofuels 2010, 3, 10. (69) TAPPI; T230 om-99 1999. (70) Stamm, A. J. Wood and Cellulose Science; Ronald Press Co.: New York, 1964.

(71) Moreira-Vilar, F. C.; Siqueira-Soares, Rd. C.; Finger-Teixeira, A.; de Oliveira, D. M.; Ferro, A. P.; da Rocha, G. J.; Ferrarese, M. d. L. L.; dos Santos, W. D.; Ferrarese-Filho, O. PLoS One 2014, 9, e110000. (72) TAPPI; T222 om-08 1988. (73) Joshi, S. R.; Kataria, K. L.; Sawant, S. B.; Joshi, J. B. Ind. Eng. Chem. Res. 2005, 44, 325−333. (74) Arendt, J. H.; Carriere, J. P.; Bouchez, P.; Sachetto, J. P. J. Polym. Sci., Polym. Symp. 1973, 42, 1521−1529. (75) Amiralian, N.; Annamalai, P. K.; Memmott, P.; Taran, E.; Schmidt, S.; Martin, D. J. RSC Adv. 2015, 5, 32124−32132. (76) Hinterstoisser, B.; Åkerholm, M.; Salmén, L. Biomacromolecules 2003, 4, 1232−1237. (77) Isogai, A.; Usuda, M.; Kato, T.; Uryu, T.; Atalla, R. H. Macromolecules 1989, 22, 3168−1393. (78) Chidambareswaran, P. K.; Sreenivasan, S.; Patil, N. B.; Sundaram, V.; Srinathan, B. J. Appl. Polym. Sci. 1976, 20, 3443−3448. (79) Sharma, P. R.; Rajamohanan, P. R.; Varma, A. J. Carbohydr. Polym. 2014, 113, 615−623. (80) Sathitsuksanoh, N.; Zhu, Z.; Wi, S.; Percival Zhang, Y. H. Biotechnol. Bioeng. 2011, 108, 521−529. (81) Ranby, B. G.; et al. Acta Chem. Scand. 1952, 6, 101−115. (82) Vane, C. H.; Drage, T. C.; Snape, C. E.; Stephenson, M. H.; Foster, C. Int. Biodeterior. Biodegrad. 2005, 55, 175−185. (83) Sebe, G.; Ham-Pichavant, F.; Ibarboure, E.; Koffi, A. K. C.; Tingaut, P. Biomacromolecules 2012, 13, 570−578. (84) Ornaghi Junior, H. L.; Zattera, A. J.; Amico, S. C. Cellulose 2014, 21, 189−201. (85) Zhang, X.; Zhao, Z.; Ran, G.; Liu, Y.; Liu, S.; Zhou, B.; Wang, Z. Powder Technol. 2013, 246, 664−668. (86) Sharma, P. R.; Varma, A. J. Carbohydr. Polym. 2014, 104, 135− 142. (87) Jahan, M. S.; Saeed, A.; He, Z.; Ni, Y. Cellulose 2011, 18, 451− 459.

J

DOI: 10.1021/acs.biomac.7b00544 Biomacromolecules XXXX, XXX, XXX−XXX