Accepted Manuscript Title: Surface esterification of cellulose nanofibers by simple organocatalytic methodology ´ Author: Jhon Alejandro Avila Ram´ırez Camila Juan Suriano Patricia Cerrutti Mar´ıa L. Foresti PII: DOI: Reference:

S0144-8617(14)00789-9 http://dx.doi.org/doi:10.1016/j.carbpol.2014.08.020 CARP 9157

To appear in: Received date: Revised date: Accepted date:

12-6-2014 11-8-2014 17-8-2014

´ Suriano, C. J., Cerrutti, P., and Foresti, M. Please cite this article as: Ram´irez, J. A. A., L.,Surface esterification of cellulose nanofibers by simple organocatalytic methodology, Carbohydrate Polymers (2014), http://dx.doi.org/10.1016/j.carbpol.2014.08.020 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

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Highlights Surface esterified cellulose nanofibers were obtained by an organocatalytic route.

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Reaction involved a non-toxic natural-origin catalyst and no cosolvent was used.

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The substitution degree was tailored by the esterification time interval used.

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Esterified BNC showed increased thermal stability and reduced polar character.

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NMR, FTIR, XRD and TGA results indicated surface-only reaction.

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Surface esterification of cellulose nanofibers by simple organocatalytic

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methodology

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Jhon Alejandro Ávila Ramírez1,2, Camila Juan Suriano1, Patricia Cerrutti1,3, María L. Foresti1,2*

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Grupo de Biotecnología y Biosíntesis - Instituto de Tecnología en Polímeros y Nanotecnología

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(ITPN) – Facultad de Ingeniería, Universidad de Buenos Aires, Las Heras 2214 (CP 1127AAR), Buenos Aires, Argentina. Tel/fax: 0054 11 45143010. *[email protected] 2.

Departamento de Ingeniería Química, Facultad de Ingeniería, Universidad de Buenos Aires.

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Consejo Nacional de Investigaciones Científicas y Técnicas (CONICET), Argentina

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26 Abstract

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Bacterial cellulose nanofibers were esterified with two short carboxylic acids by means

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of a simple and novel organic acid-catalyzed route. The methodology proposed relayed on the

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use of a non-toxic biobased α-hydroxycarboxylic acid as catalyst, and proceeded under

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moderate reaction conditions in solventless medium. By varying the esterification interval,

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acetylated and propionized bacterial cellulose nanofibers with degree of substitution (DS) in the

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0.02-0.45 range could be obtained. Esterified bacterial cellulose samples were characterized by

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means of Solid-State CP/MAS

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NMR), Fourier Transform Infrared spectroscopy (FTIR), X-ray Diffraction (XRD),

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Thermogravimetric Analysis (TGA), and chosen hydrophobicity test assays. TGA results

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showed that the esterified nanofibers had increased thermal stability, whereas XRD data

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evidenced that the organocatalytic esterification protocol did not alter their crystallinity. The

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analysis of the ensuing modified nanofibers by NMR, FTIR, XRD and TGA demonstrated that

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esterification occurred essentially at the surface of bacterial cellulose microfibrils, something

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highly desirable for changing their surface hydrophilicity while not affecting their ultrastructure.

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C Nuclear Magnetic Resonance spectroscopy (CP/MAS

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Keywords: bacterial cellulose; surface esterification; organocatalysis; characterization

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1. Introduction

46 Cellulose is the main component of wood and most natural fibers, and it is also

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produced by tunicates and certain bacteria. In the last decade, cellulose microfibrils and

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cellulose nanocrystals obtained from the mentioned sources have received increasing attention

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based in their unique properties such as high water holding capacity, high crystallinity, low

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thermal expansion coefficient in the axial direction, and high tensile strength and Young

52

modulus. Among nanocellulose applications, its use as reinforcement of nanocomposites is a

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continuously growing field (Hubbe, Rojas, Lucia, & Sain, 2008; Siro & Plackett, 2010;

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Siqueira, Bras, & Dufresne, 2010). The relative low cost, low density, availability,

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renewability and environmentally benign character of cellulose, as well as the proved high

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mechanical properties of nanocellulosics, have encouraged their use as matrix reinforcement.

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However, the highly polar surface of native cellulose nanoparticles associated with their OH-

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rich structure leads to low interfacial compatibility with non-polar matrices which results in

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inadequate mechanical performance of the corresponding composites; high moisture uptake

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which leads to loss of fiber strength and composite deformation; and inter-fiber aggregation by

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hydrogen bonding which results in poor fiber dispersion in the composite (Ifuku et al., 2007;

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Tomé et al., 2011; Yuan, Nishiyama, Wada, & Kuga, 2006; Berlioz, Molina-Boisseau,

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Nishiyama, & Heux, 2009).

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To overcome the mentioned difficulties, chemical modification of cellulose nanofibers

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can be attempted. In particular, esterification reactions in which an acylant reacts with the

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hydroxyl groups of cellulose nanofibers to produce less hydrophilic ester groups is a common

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technique used to turn the cellulose surface into a more hydrophobic one. Esterification of

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cellulose nanofibers can be accomplished by use of homogeneous or heterogeneous processes.

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In the homogeneous mechanism, (mainly applied in cellulose acetylation processes), the

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esterified cellulose microfibrils are soluble in the medium, and thus the chains located at the

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cellulose surface diffuse into the surrounding solvent as soon as they are substantially

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derivatized (Sassi & Chanzy, 1995). On the other hand, in the heterogeneous or “fibrous”

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process, esterification is carried out in a medium that is a nonsolvent for both native and

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esterified cellulose; and cellulose nanofibers morphology is preserved (Berlioz et al., 2009). In

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the last decade, different heterogeneous methodologies for the esterification of cellulose

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nanoparticles have been proposed, including acetylation of compressed bacterial cellulose

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pellicles catalyzed by perchloric acid in toluene medium (Ifuku et al., 2007), surface acylation

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of cellulose whiskers with alkyenyl succinic anhydride aqueous emulsion followed by heat

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annealing (Yuan et al., 2006), solvent-free gas-phase esterification of cellulose whiskers and

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bacterial cellulose microfibrils with palmitoyl chloride (Berlioz et al., 2009), simultaneous

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isolation and Fischer esterification of hydroxyl groups of cellulose nanowhiskers in aqueous

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medium using acetic and butyric acids (Braun & Dorgan, 2009), heterogeneous esterification

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of cellulose whiskers with long-chain fatty acids in p-tosyl chloride/pyridine medium

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(Uschanov, Johansson, Maunu, Laine, 2011), solvent-free acetylation of bacterial cellulose

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using acetic anhydride in the presence of iodine as a catalyst (Hu, Chen, Xu, & Wang, 2011),

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and heterogeneous esterification of bacterial cellulose using ionic liquids as solvent media and

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catalysts (Tomé et al., 2011), among others.

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In the current manuscript, bacterial nanocellulose (BNC) was heterogeneously esterified

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in absence of solvents using an operationally simple eco-friendly methodology. Esterification

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with acetic and propionic acids as acylants was accomplished by use of an organic-acid-

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catalyzed heterogeneous route developed by Hafrén & Córdova, (2005) for the

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organocatalytic ring-opening polymerization of ε-caprolactone from cotton and paper cellulose.

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The mentioned authors also applied this route for the esterification of cotton fibers with

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hexadecanoic and pentynoic acids catalyzed by tartaric acid. The organocatalytic esterification

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methodology described has recently been applied to the synthesis of starch acetates and

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butyrates with varying acylation levels (Tupa, Maldonado, Vazquez, & Foresti, 2013).

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However, at the authors’ best knowledge the esterification of cellulose nanofibers by use of the

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described organocatalytic route has not been reported before. Acetylated and propionized

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cellulose nanofibers obtained herein were characterized by means CP/MAS

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DRX, TGA and chosen hydrophobicity test assays, in order to demonstrate the suitability of the

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organic acid-catalyzed route for BNC esterification, and study the effect of derivatization on

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particular properties of BNC.

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2. Materials and Methods 2.1. Microorganism

The bacterial strain of Gluconacetobacter xylinus (syn. Acetobacter aceti subsp. xylinus,

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Acetobacter xylinum) NRRL B-42 used in this work was gently provided by Dr. Luis Ielpi

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(Fundación Instituto Leloir, Buenos Aires, Argentina).

114 115

2.2. Materials

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Acetic acid (99.5 %) and propionic acid (99.5%) were purchased from Dorwil.

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Hydrochloric acid (36.5 – 38 %) and sodium hydroxide were reagent grade chemicals purchased

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from Anedra and BioPack, respectively. Potassium hydrogen phthalate and sodium carbonate

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were bought from Laboratorios Cicarelli and Mallinckrodt, respectively. For BNC production,

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anhydrous dextrose (Biopack), meat peptone and yeast extract (Britania, Laboratorios Britania

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S.A.), disodium phosphate (Anhedra), citric acid (Merck), glycerol (Sintorgan) and corn steep

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liquor (Ingredion) were used.

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Inocula were cultured for 48 h in Erlenmeyers flasks containing Hestrin and Schramm

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(HS) medium (%, w/v): glucose, 2.0; peptone, 0.5; yeast extract, 0.5; disodium phosphate, 0.27;

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citric acid, 0.115. The pH was adjusted to 6.0 with dil. HCl or NaOH. Agitation (200 rpm) was

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provided by an orbital shaker. For BNC production, static incubations at 28ºC were performed

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in Erlenmeyers flasks containing 4.0% w/v glycerol and corn steep liquor respectively, and

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keeping a flask/autoclaved medium v/v ratio of 5:1. After 14 days bacterial nanocellulose

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pellicles were harvested, rinsed with water to remove the culture medium, and homogenized in a

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blender for 5 min. The microfibrils thus obtained were treated for 14 h in a 5 % w/v KOH

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solution in order to eliminate the bacterial cells from the cellulose matrix, and then rinsed with

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water till neutralization.

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2.4. Organocatalytic esterification of BNC

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Homogenized bacterial nanocellulose pellicles (0.5 g, dry weight) were solvent

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exchanged from water to the acylant by soaking and centrifugation (three times, 20 mL). The

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acylant (0.53 mol, i.e. 30 mL of acetic acid or 40 mL of propionic acid), tartaric acid (0.47 g)

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and soaked BNC were mixed in an oven-dried 100 mL glass flask equipped with a magnetic stir

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bar, and a reflux condenser to prevent the loss of the acylant. The mixture was heated to 120 °C

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under continuous agitation in a thermostatized oil bath. When the mixture achieved the target

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temperature, all tartaric acid dissolved and this was considered as the beginning of reaction.

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Esterification was run for different reaction times (1, 3, 4, 5, 6 and 8 h). After the chosen

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reaction time, the mixture was allowed to cool down to room temperature, and the solid product

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was separated by vacuum filtration in a Buchner funnel. Several washings of the recovered solid

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with distilled water were performed in order to guarantee the removal of the catalyst and the

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unreacted acid.

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The acyl content of the esterified nanofibers was determined by heterogeneous

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saponification and back titration with HCl, as an adaptation of the approach used for

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determining the acyl content in cellulose acetate (ASTM D871 – 96, Standard Test Methods

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of Testing Cellulose Acetate). With this purpose, esterified BNC samples were made into

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pellicles by drying, grinded under liquid nitrogen flux, and carefully dried again in order to have

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a precise measure of the mass of sample used in saponification. With the described sample

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preparation protocol the method proved simple and accurate to quantitatively assay the bulk

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level of substitution achieved in BNC without specific instrumentation. Dried grinded samples

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(0,10 g, 105 ºC, 2 h) were thus transferred to 100 mL Erlenmeyers to which 10 mL of ethyl

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alcohol (75 %) were added. Flasks were heated loosely stoppered for 30 min at 50 °C.

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Afterwards, the suspensions were brought to slightly basic pH by addition of a few drops of

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0.1N NaOH using phenolphthalein as indicator. Aliquots of 10 mL of 0.1N NaOH were then

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added to each flask, and heated again at 50 °C for 15 min. Erlenmeyers were finally allowed to

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stand tightly stoppered at room temperature for 72 h. At the end of this time, the excess NaOH

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present in the flasks was back titrated with 0.1N HCl, using phenolphthalein as the end-point

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indicator. A blank determination (grinded native BNC) was carried through the complete

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procedure. NaOH and HCl solutions were standardized using previously dried standard

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potassium hydrogen phthalate and sodium carbonate, respectively.

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Acyl (%) = [(VB − VS) × NHCl × Macyl × 10−3 × 100] / W

(1)

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The acyl content was then calculated by:

Where VB (mL) is the volume of HCl required for titration of the blank; VS (mL) is the

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volume of HCl required to titrate the sample; NHCl is the normality of the HCl solution; Macyl is

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the molecular weight of the acyl group (43 for acetyl and 57 for propionyl), and W (g) is the

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mass of sample used.

181 182 183 184 185 186 187 188

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Using the Acyl % value, the substitution degree of esterified BNC samples was

calculated by:

DS = (162 × Acyl %) / [Macyl × 100 − ((Macyl − 1) × Acyl %)]

(2)

where 162 is the molecular weight of the anhydroglucose units.

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2.5. Characterization of esterified BNC

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Solid-State 13C NMR measurements: High-resolution 13C solid-state spectra of grinded

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samples were recorded using the ramp {1H} → {13C} CP/MAS pulse sequence (cross-

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polarization and magic angle spinning) with proton decoupling during acquisition. All

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experiments were performed at room temperature in a Bruker Avance II-300 spectrometer

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equipped with a 4-mm MAS probe. The operating frequency for protons and carbons was

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300.13 and 75.46 MHz, respectively. Glycine was used as an external reference for the

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spectra and to set the Hartmann-Hahn matching condition in the cross-polarization experiments.

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The recycling time varied from 5 to 6 s according to the sample. The contact time during CP

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was 2ms for all of them. The SPINAL64 sequence (small phase incremental alternation with 64

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steps) was used for heteronuclear decoupling during acquisition with a proton field H1H

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satisfying ω1H/2π = γHH1H = 62 kHz. The spinning rate for all the samples was 10 kHz. The

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crystallinity index (CI) of chosen samples was determined by separating the C4 region of the

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spectrum into crystalline and amorphous peaks, and calculated by dividing the area of the

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crystalline peak (87 to 93 ppm) by the total area assigned to the C4 peak (80 to 93 ppm) as

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described elsewhere (Newman, 2004; Park, Baker, Himmel, Parrilla, & Johnson, 2010).

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native and esterified grinded samples were acquired on an IR Affinity-1 Shimadzu Fourier

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Transform Infrared Spectrophotometer in transmission mode. Carefully dried (10 mg, 105 ºC, 1

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h) grinded samples of native and acetylated BNC were mixed with previously dried (130ºC,

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overnight) KBr in the ratio 1:25. The prepared pellets were further dried overnight at 105ºC and

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spectra were finally collected with 40 scans in the range of 4000 to 700 cm-1 with a resolution of

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4 cm-1. Samples were normalized against the band found at 1165 cm-1 assigned to the (C–O–C)

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link of cellulose (Ilharco, Gracia, daSilva, & Ferreira, 1997; Lee et al., 2011).

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Fourier Transform Infrared Spectroscopy (FTIR): Fourier transform infrared spectra of

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X-ray diffraction (XRD): The structure of esterified BNC grinded samples was analysed

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with a Rigaku D/Max-C Wide Angle automated X-ray diffractometer with vertical goniometer.

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The X-ray diffraction pattern was recorded in a 2θ angle range of 10 to 40° at a step size of

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0.02º. The wavelength of the Cu/Ka radiation source used was 0.154 nm, generated at

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accelerating voltage of 40kV and a filament emission of 30 mA. In order to calculate the

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crystallinity index of native and esterified BNC samples Segal’s method was applied (Segal,

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Creely, Martin, & Conrad, 1959):

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CI = (I002 – Iam)/ I002 × 100

(3)

Where I002 corresponds to the maximum intensity of the 002 lattice diffraction, and Iam corresponds to the intensity at 2θ=18º.

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Thermogravimetric Analysis (TGA): Thermogravimetric analysis of dried grinded

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samples (2.5 - 3 mg, 105 ºC, 1 h) was conducted in a TGA-50 Shimadzu instrument.

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Temperature programs were run from 25°C to 550°C at a heating rate of 10°C/min, under

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nitrogen atmosphere (30 mL/min) in order to prevent thermoxidative degradation. Analysis of Hydrophobicity: Native and esterified grinded BNC samples were

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qualitatively tested for hydrophobicity by water flotation in water filled vials, as well as by

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distribution in test tubes containing two immiscible polar/non polar liquid phases (i.e. distilled

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water/petroleum ether). Neat and esterified pellicles were also subjected to water drop tests, for

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which 10 μL water droplets were manually placed on the pellicles surface using a syringe, and

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the time required for complete absorption was determined.

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242 3. Results and Discussion

244 3.1. Organocatalytic esterification of BNC

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Figure 1 illustrates the evolution of the degree of substitution (DS) determined for

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acetylated and propionized BNC samples as a function of time. As it is shown, using both acids

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the substitution degree increased with reaction time, reaching a DS value of 0.45 for the

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acetylation reaction after 8 hours of reaction. Using propionic acid as acylant, a lower

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substitution degree was achieved, reaching a DS of 0.23 in 8 hours of reaction. The lower

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acylation level observed with propionic acid is explained in terms of a lower reactivity of this

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acylant due to a bulkier fatty acylium ion with higher steric hindrance. Similar tendencies have

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been reported previously by other authors who studied the heterogeneous esterification of

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cellulose nanofibers with acylants of different chain length (Uschanov et al., 2011; Tomé et

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al., 2011; Lee et al., 2011). The contribution of the uncatalyzed reaction was in all cases lower

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than 10% of the measured DS.

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Literature dealing with cellulose nanofibers esterification by other methodologies has

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demonstrated that for a number of applications low acyl contents have succeeded in altering

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cellulose nanofibers properties in the desired way. Nogi et al. (2006), prepared a nanocomposite

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consisting of an acrylic resin reinforced with acetylated BNC nanofibers with a DS of 0.17.

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Authors observed that the DS achieved effectively reduced the hygroscopicity of BNC

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nanocomposites and improved thermal degradation resistance regarding transparency. In the

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surface acetylation of bacterial cellulose nanofibers using perchloric acid as catalyst, by varying

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the volume of acylant used Ifuku et al. (2007) obtained acetylated nanofibers with DS in the 0-

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1.76 interval, which were also used as acrylic resins reinforcement. Authors demonstrated that

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the moisture content of the nanocomposites was minimized at intermediate DS, i.e. 0.45-0.56,

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which they explained in terms of two opposite effects: change in nanofibers surface

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hydrophobicity and water penetration into the internal spaces of the esterified nanofibers upon

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bulky ester groups introduction.

272 Although it has not been completely elucidated, the catalytic mechanism of

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esterifications catalyzed by α-hydroxy acids such as tartaric acid, has been proposed to proceed

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analogously to that of lipase/esterase-catalyzed reactions (Domínguez de María, 2010).

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According to the mechanism proposed, the acetylation and propionization reactions described

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herein (Scheme 1) initiate with a nucleophilic attack of the acylant by the hydroxyl group of

278

tartaric acid. The intermediate formed is then further nucleophilically attacked by the hydroxyl

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groups of bacterial cellulose nanofibers, resulting in transesterification (Dominguez de María,

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2010; Hafrén & Córdova, 2005).

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3.2. CP/MAS 13C NMR spectroscopy

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Esterification of BNC by the proposed organocatalytic methodology was confirmed by

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use of CP/MAS

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obtained for native, acetylated (DS=0.22), and propionized (DS=0.18) BNC. Native BNC

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spectrum showed carbon peaks typical of cellulose I, i.e. C1: 105 ppm, C4: 89 ppm, C4′: 84

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ppm, cluster C2-C3-C5: 70-80 ppm, C6: 65.3 ppm, and C6′: 62.7 ppm (Earl & VanderHart,

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1980). As the esterification reaction proceeded, several new resonances appeared on the spectra

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(Figure 2 b,c). In the spectrum of the acetylated BNC a new peak was observed at 174 ppm,

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assignable to the resonance of the carbonyl ester peak. For BNC esterification with propionic

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acid this signal was observed at 176.2 ppm. Both samples also showed the appearance of new

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signals between 9 and 30 ppm, which arose from the terminal methyl carbons of acetic (21.3

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ppm) and propionic acid (9.6 ppm) (Yamamoto, Horrii, & Hirai, 2005). The signal observed

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at 27 ppm in the propionized sample corresponds to CH2. Signals attributable to tartaric acid

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acting as grafting molecule were not observed.

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C NMR spectroscopy. Figure 2 shows the CP/MAS

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Crystalline and disordered components of cellulose are detected in solid-state 13C NMR

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spectra as downfield and upfield lines for the C4 or C6 carbons, respectively (Yamamoto et al.,

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2005; Jandura, Kokta, & Riedl, 2000). As shown in Figure 2, native BNC exhibited broad

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non-crystalline resonances centred at 84 and 62.7 ppm (upfield lines, i.e. C4` and C6`,

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respectively). Downfield signals centred at 89 and 65.3 ppm (C4 and C6, respectively) have

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been assigned to ordered cellulose structures (Atalla & Vanderhart, 1999; Park et al. 2010).

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The crystallinity index of native, acetylated (DS= 0.22) and propionized BNC (DS= 0.18) was

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calculated using the peak separation method of Newman (2004), by dividing the area of the

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crystalline peak by the total area of assigned to the C4 peaks. The CI values obtained suggested

308

that no significant change in the crystallinity of BNC took place during esterification (Table 1,

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CI NMR).

310 3.3. FTIR spectroscopy

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BNC esterification was further qualitatively confirmed by FTIR spectroscopy of

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acylated samples. Figure 3 shows the FTIR spectra of acetylated and propionized BNC with

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chosen DS. The spectrum of native BNC was also included for comparison. Native BNC

316

spectrum showed bands typical of cellulose I, showing the presence of hydroxyl groups (3360

317

cm-1); C-H stretching (2895 cm-1); H-O-H bending vibration of absorbed water molecules (1647

318

cm-1); CH2 symmetrical bending (1427 cm-1); cellulose C-O-C bridges (1165 cm-1); C-O bond

319

stretching (1118 cm-1); ether C-O-C functionalities (1061 cm-1); and the band at 897 cm-1 which

320

is typical of β-linked glucose polymers (Castro et al., 2011; Moosavi-Nasab, & Yousefi,

321

2011; Rani, Navin, & Appaiah, 2011; Ashori, Babaee, Jonoobi, & Hamzed, 2014).

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Spectra of acetylated and propionized BNC provided evidence of esterification by

324

appearance of a new band at 1730 cm-1 which is assigned to the stretching of the ester carbonyl

325

groups (C=O) introduced (Lee et al., 2011; Tomé et al., 2011; Ashori et al., 2014; Yuan et al.,

326

2006; Braun & Dorgan, 2009). As shown in Figure 3, in the spectra of samples with lower DS

327

the intensity of the C=O ester was found to increase with the DS measured by saponification.

328

On the other hand, although esterification implied substitution of a fraction of hydroxyl groups

329

by less polar ester groups, no clear reduction of the intensity of the band associated with the

330

vibration of OH groups located at 3300 cm-1 was observed. This behaviour has previously been

331

attributed to BNC derivatization occurring essentially on the accessible hydroxyl groups of the

332

surface of the nanofibers or in the amorphous fraction of the cellulose (Lee et al., 2011). The

333

absence of absorption at 1700 cm−1 attributed to carboxylic groups indicated that no unreacted

334

acylant remained in the esterified BNC.

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3.4. X-ray diffraction (XRD)

337 338

Cellulose has several crystalline polymorphisms (I, II, III, IV). Cellulose I is the

339

crystalline polymorphism that is naturally produced by a variety of organisms, i.e. trees, plants,

340

tunicates, algae and bacteria; and it is the crystal structure with the highest axial elastic modulus

341

(Moon, Martini, Nairn, Simonsen, & Youngblood, 2011). Diffraction patterns obtained for

342

native BNC, acetylated BNC (DS=0.14 and DS=0.45) and propionized BNC (DS=0.09 and

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DS=0.23) are shown in Figure 4. Native BNC showed four diffraction peaks at 2θ= 14.4° (101),

344

16.7° (10-1), 22.5° (002), and 34.0º (040) which confirmed that only cellulose I was present in

345

all samples (Vazquez, Foresti, Cerrutti, & Galvagno, 2013; Johnson Ford, Mendon,

346

Thames, & Rawlins, 2010; Terinte, Ibbett, & Schuster, 2011; Gea et al., 2011). For the DS

347

attained in this contribution, esterified cellulose nanofibers maintained the diffraction pattern of

348

native BNC, with no evidence of peak shifting or appearance of new peaks. This is further

349

evidence that the organocatalytic reaction involved essentially the OH groups on the surface or

350

in the amorphous regions of BNC, not affecting in a great extent its ultrastructure (Tomé et al.,

351

2011; Lee et al., 2011). The degree of crystallinity of native and modified BNC was determined

352

using the Segal equation (Segal et al., 1959). Results included in Table 1 (CI

353

confirmed that no significant change in BNC crystallinity took place during organic-acid-

354

catalyzed esterification of BNC. As expected, absolute CI values differed from those obtained

355

by NMR peak separation methods, since CI values are known to vary significantly depending

356

on the measurement method used to determine them (Park et al., 2010; Terinte et al., 2011).

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3.5. Thermogravimetric analysis (TGA)

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Esterified BNC samples were characterized by thermogravimetric analysis to study the

362

impact of organocatalytic esterification on the thermal properties of BNC. Figure 5 shows the

363

TG and DTG curves obtained for (i) acetylated BNC and (ii) propionized BNC samples with

364

increasing DS. DTG data have been normalized with respect to the initial sample mass, and

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results have been moved up in the y axes for clarity purposes. The TG and DTG curves of

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native BNC have also been included for comparison.

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Neat BNC pyrolysis showed a single weight loss stage, with an extrapolated onset

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temperature (Tonset) of 313ºC. Tonset was calculated as the intersection of the extrapolated pre-

370

decomposition ordinate value and a tangential line drawn to the point of steepest slope of the

371

weight loss curve in the decomposition region. According to DTG data, the maximum weight

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loss rate of neat BNC (Tmax) took place at 329 ºC. Thermogravimetric data of neat BNC showed

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a minimal weight loss assignable to BNC dehydration (2.3%), due to sample preconditioning at

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105ºC during 1 h. The measured weight loss (%) during samples’ preconditioning decreased as

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the DS of the samples increased, which was a consequence of the effect of esterification on

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BNC hydrophilicity.

377 378

As it is shown in Figure 5-i, acetylated bacterial cellulose nanofibers descomposed in a

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single step process. Irrespectively of acylation level achieved, acetylated nanofibers started to

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decompose at substantially higher temperatures than neat BNC, with Tonset values that were

11 Page 11 of 26

found to increase with the DS of the sample. Tmax values of acetylated samples also showed a

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positive correlation with DS. Similar tendencies were found for propionized BNC (Figure 5-ii).

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The increase of Tonset and Tmax with the DS of acetylated and propionized samples is illustrated

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in Figure 6. Results included in Figure 6 also suggested that the length of the ester group

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introduced showed no significant effect on the Tonset and Tmax values registered for acetylated

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and propionized samples with similar DS. Higher thermal stability of modified celluloses for

387

increasing substitution degrees has previously been reported (Jandura, Riedl, & Kokta, 2006;

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Morgado & Frollini, 2011; Uschanov et al., 2011). Increased thermal stability of esterified

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samples has been attributed to the replacement of hydroxyl groups by more stable ester groups

390

(Ashori et al., 2014; Lin, Huang, Chang, Feng, & Yu, 2011).

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Among the methodologies used in the literature to heterogeneously esterify BNC, a

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number of authors have reported a reduction in the thermal stability of cellulose nanoparticles

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upon modification, characterized by onset and maximum temperature decomposition peaks

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found at lower temperatures than those of neat BNC (Tomé et al., 2011; Uschanov et al., 2011;

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Lee et al., 2011). The mentioned behaviour was attributed to the reduction of cellulose

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crystallinity induced by esterification, and, less frequently, to the high liability of the ester

398

linkages introduced in the cellulose structure, or to the reduced number of effective hydrogen

399

bonds remaining among BNC nanofibers upon esterification (Tomé et al., 2011; Uschanov et

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al., 2011; Lee et al., 2011). In this context, the increase in thermal stability of BNC shown in

401

Figure 5 is in agreement with previous characterization data which evidenced that the

402

organocatalytic esterification of cellulose nanofibers took place in their outmost layers, while

403

keeping the ultrastructure unaffected. Specially when used as reinforcement for polymers whose

404

melting processing require elevated temperatures, it is important that alterations of the surface

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chemistry of cellulose nanofibers do not decrease the onset temperature of thermal degradation

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(Braun & Dorgan, 2009).

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In the case of propionized BNC samples a second high temperature decomposition peak

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was observed in the 430-550 ºC range (Figure 5-ii). The DTG peak was the result of a second

410

thermal degradation step registered in TG curves of propionized samples, which led to weight

411

residues fractions much lower than those observed for neat BNC in the same temperature

412

interval. The behaviour observed suggested that organocatalytic propionization induced changes

413

that increased the volatility of the solid residue of BNC samples at high temperatures. Further

414

studies on the nature and reasons for the mentioned effects of organocatalytic propionization of

415

BNC are currently going on.

416 417 12 Page 12 of 26

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3.6. Surface hydrophobicity

419 Grinded acetylated and propionized BNC samples were qualitatively tested for

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hydrophobicity by water flotation, distribution in polar/non-polar biphasic systems, and water

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drop tests. All assays confirmed that acetate or propionate groups covalently attached to the

423

surface of BNC altered its polarity. Sample distribution in test tubes containing equal volumes

424

of distilled water and petroleum ether (0.67 g/mL) is illustrated in Figure 7 for chosen samples.

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When added to the biphasic liquid system, native BNC fell to the lower aqueous phase,

426

absorbed water and sank immediately to the bottom of the test tube. Contrarily, acetylated

427

(DS=0.45) and propionized (DS=0.23) BNC grinded samples fell across the non polar phase and

428

remained floating in the non-polar/polar interphase without crossing to the aqueous phase

429

(Figure 7c and 7f, respectively). The described sample distribution did not change upon

430

vertically shaking of test tubes. For both acylants, esterified samples with lower DS showed

431

intermediate behaviours (Figure 7b and 7e, respectively). In water flotation tests, samples were

432

placed on the surface of water-filled vials. In the case of native BNC the sample absorbed water

433

and sank immediately to the vial bottom. In contrast, esterified samples corresponding to

434

DS=0.45 (acetylated BNC) and DS=0.23 (propionized BNC), did not absorb water and

435

remained floating in the water surface (data not shown). In water drop tests 10 μL water

436

droplets were manually placed on the surfaces of native, acetylated (DS= 0.14) and propionized

437

(DS= 0.12) BNC pellicles. Figure 7g-i shows the appearance of the droplets after 10 seconds.

438

Whereas native BNC surface immediately absorbed the water droplet (Figure 7g), a significant

439

hydrophobicity of samples surface was observed in esterified pellicles even at relatively low

440

DS. In the case of the acetylated pellicle (DS=0.14, Figure 7h), it took 38-40 min to completely

441

absorb the drop, whereas in the case of propionized BNC (DS=0.12, Figure 7i) 37-39 min were

442

required.

444 445 446 447

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4. Conclusion

A simple environmental friendly esterification route to confer hydrophobicity to

448

cellulose nanofibers was applied to the acetylation and propionization of bacterial cellulose

449

nanofibers. The protocol applied was characterized by the use of a non-toxic natural-origin

450

catalyst and the absence of cosolvent. By controlling the esterification time interval acetylated

451

and propionized cellulose nanofibers with varying DS could be easily obtained. The

452

esterification reaction was further confirmed by solid-state CP/MAS

453

spectroscopy. XRD and TGA were used to monitor the effect of esterification on the

454

crystallinity and thermal properties of BNC. Esterified BNC samples showed increased thermal

13

C NMR and FTIR

13 Page 13 of 26

stability in terms of Tonset and Tmax, and no crystallinity reduction with respect to native BNC.

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Besides, hydrophobicity tests confirmed the change in cellulose microfibrils polarity. NMR,

457

FTIR, XRD and TGA results all contributed to the conclusion that the organocatalytic

458

acetylation and propionization of BNC occurred mainly in the surface of cellulose nanofibers.

459

Surface esterification of cellulose nanofibers is desirable for improving the compatibility of the

460

cellulosic substrate with apolar matrices, while preserving the crystalline ultrastructure of

461

cellulose. In this context, the organocatalytic route proposed appears as a promising ecofriendly

462

approach for tailoring the surface properties of BNC to be used for example in the development

463

of biodegradable polyester reinforced nanocomposites. Further assays, aiming to assert the

464

effect of reaction conditions to be used to tune the extent of esterification achieved in BNC and

465 466 467

other cellulose nanoparticles substrates are currently on progress.

468

5. Ackowledgements

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Authors acknowledge Consejo Nacional de Investigaciones Científicas y Técnicas

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(CONICET- PIP 11220110100608), University of Buenos Aires (UBACYT 20020090100065),

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and Agencia Nacional de Promoción Científica y Tecnológica for financial support (PICT 1957

473 474 475 476 477 478 479 480 481 482 483 484 485 486 487 488 489 490 491 492 493 494 495 496 497 498 499

2012 – PRESTAMO BID).

6. References

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Ashori, A., Babaee, M., Jonoobi, M., & Hamzed, Y. (2014). Solvent-free acetylation of cellulose nanofibers for improving compatibility and dispersion. Carbohydrate Polymers, 102, 369-375. Atalla, R.H., & VanderHart, D.L. (1999). The role of solid state 13C NMR spectroscopy in studies of the nature of native celluloses. Solid State Nuclear Magnetic Resonance, 15, 1-19. ASTM D871 – 96, Standard Test Methods of Testing Cellulose Acetate. Berlioz, S., Molina-Boisseau, S., Nishiyama, Y., & Heux, L. (2009). Gas-phase surface esterification of cellulose microfibrils and whiskers. Biomacromolecules, 10, 2144-2151. Braun, B., & Dorgan, J.R. (2009). Single-step method for the isolation and surface functionalization of cellulosic nanowhiskers. Biomacromolecules, 10, 334-341. Castro, C., Zuluaga, R., Putaux, J.L., Caroa, G., Mondragon, I., & Gañan, P. (2011). Structural characterization of bacterial cellulose produced by Gluconacetobacter swingsii sp. from Colombian agroindustrial wastes. Carbohydrate Polymers, 84, 96-102. Domínguez de María, P. (2010). Minimal hydrolases: Organocatalytic ring-opening polymerizations catalyzed by naturally occurring carboxylic acids. ChemCatChem, 2, 487-492.

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Earl, W.L., & VanderHart, D.L. (1980). High resolution, magic angle sample spinning 13C NMR of solid cellulose I. Journal of the American Chemical Society, 102, 3251-3252. Gea, S., Reynolds, C.T., Roohpour, N., Wirjosentono, B., Soykeabkaew, N., Bilotti, E., & Peijs, T. (2011). Investigation into the structural, morphological, mechanical and thermal behaviour of bacterial cellulose after a two-step purification process. Bioresource Technology, 102, 91059110.

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Hafrén, J., & Córdova, A. (2005). Direct organocatalytic polymerization from cellulose fibers. Macromolecular Rapid Communications, 26, 82–86.

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Hu, W., Chen, S., Xu, Q., & Wang, H. (2011). Solvent-free acetylation of bacterial cellulose under moderate conditions. Carbohydrate Polymers, 83,1575–1581.

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Hubbe, M.A., Rojas, O.J., Lucia, L.A., & Sain M. (2008). Cellulosic nanocomposites: a review. BioResources, 3, 929-980.

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Ifuku, S., Nogi, M., Abe, K., Handa, K., Nakatsubo, F., & Yano, H. (2007). Surface modification of bacterial cellulose nanofibers for property enhancement of optically transparent composites: dependence on acetyl-group DS. Biomacromolecules, 8, 1973-1978.

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Ilharco, L.M., Gracia, R.R., daSilva, J.L., & Ferreira, L.F.V. (1997). Infrared approach to the study of adsorption on cellulose: influence of cellulose crystallinity on the adsorption of benzophenone. Langmuir, 13, 4126-4132.

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Jandura, P., Kokta, B.V., & Riedl, B. (2000). Fibrous long-chain organic acid cellulose esters and their characterization by diffuse reflectance FTIR spectroscopy, solid-state CP/MAS 13CNMR, and W-Ray Diffraction. Journal of Applied Polymer Science, 78, 1354-1365.

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Jandura, P., Riedl, B., & Kokta, B.V., (2000). Thermal degradation behaviour of cellulose fibers partially esterified with some long chain organic acids. Polymer Degradation and Stability, 70, 387-394. Johnson Ford, E.N., Mendon, S.K., Thames, S.F., & Rawlins, J.W. (2010). X-ray diffraction of cotton treated with neutralized vegetable oil-based macromolecular crosslinkers. Journal of Engineered Fibers and Fabrics 5, 10-20.

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500 501 502 503 504 505 506 507 508 509 510 511 512 513 514 515 516 517 518 519 520 521 522 523 524 525 526 527 528 529 530 531 532 533 534 535 536 537 538 539 540 541 542 543 544 545 546 547 548 549 550 551 552 553 554

Lee, K.-Y., Quero, F., Blaker, J.J., Hill, C.A.S., Eichhorn, S.J., Bismarck, A. (2011). Surface only modification of bacterial cellulose nanofibers with organic acids. Cellulose, 18, 595-605. Lin, N., Huang, J., Chang, P. R., Feng, J., & Yu, J. (2011). Surface acetylation of cellulose nanocrystal and its reinforcing function in poly(lactic acid). Carbohydrate Polymers, 83, 1834– 1842. Moon, R.J., Martini, A., Nairn, J., Simonsen, J., & Youngblood, J. (2011) Cellulose nanomaterials review: structure, properties and nanocomposites. Chemical Society Reviews, 40, 3941-94. Moosavi-Nasab, M., & Yousefi, A. (2011). Biotechnological production of cellulose by Gluconacetobacter xylinus from agricultural waste. Iranian Journal of Biotechnology, 9, 94101. Morgado, D.L., & Frollini, E. (2011). Thermal decomposition of mercerized linter cellulose and its acetates obtained from a homogeneous reaction. Polímeros, 21, 111-117.

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Newman, R.H. (2004). Homogeneity in cellulose crystallinity between samples of Pinus radiata wood. Holzforschung, 58, 91-96 Nogi, M., Abe, K., Handa, K., Nakatsubo, F., Ifuku, S., & Yano, H. (2006). Property enhancement of optically transparent bionanofiber composites by acetylation. Applied Physics Letters, 89, 233123-1-233123-3.

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Park, S., Baker, J.O., Himmel, M.E., Parilla, P.A., & Johnson, D.K. (2010). Cellulose crystallinity index: measurement techniques and their impact on interpreting cellulose performance. Biotechnology for Biofuels, 3, 1-10.

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Rani, M.U., Navin, K.R., & Appaiah, K.A.A. (2011). Statistical optimization of medium composition for bacterial cellulose production by Gluconacetobacter hansenii UAC09 using coffee cherry husk extract – an agro-industry waste. Journal of Microbiology and Biotechnology, 21, 739-745. Sassi, J.-F., & Chanzy, H. (1995). Ultrastructural aspects of the acetylation of cellulose, Cellulose, 2, 111–127.

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Segal, L., Creely, J.J., Martin, A.E., Conrad, C.M. (1959). An empirical method for estimating the degree of crystallinity of native cellulose using the X-ray diffractometer. Textile Research Journal, 29, 786-794.

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Siqueira, G., Bras, J., & Dufresne, A. (2010). Cellulosic bionanocomposites: a review of preparation, properties and applications. Polymers, 2, 728-765.

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Siro, I., & Plackett, D. (2010). Microfibrillated cellulose and new composite materials: a review. Cellulose, 17, 459-494.

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Terinte, N., Ibbett, R., & Schuster, K.C. (2011). Overview on native cellulose and microcrystalline cellulose I structure studied by X-ray diffraction (WAXD): comparison between measurement techniques. Lenzinger Berichte, 89, 118-131. Tomé, L.C., Freire, M.G., Rebelo, L.P.N., Silvestre, A.J.D., Neto, C.P., Marrucho, I.M., Freire, C.S.R. (2011). Surface hydrophobization of bacterial and vegetable cellulose fibers using ionic liquids as solvent media and catalysts. Green Chemistry, 13, 2464-2470.

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Tupa, M., Maldonado, L., Vázquez, A., & Foresti, M.L. (2013). Simple organocatalytic route for the synthesis of starch esters. Carbohydrate Polymers, 98, 349– 357. Uschanov, P., Johansson, L.-S., Maunu, S.L., & Laine, J. (2011). Heterogeneous modification of various celluloses with fatty acids. Cellulose, 18, 393-404. Vazquez, A., Foresti, M.L. Cerrutti, P., & Galvagno, M.A. (2013). bacterial cellulose from simple and low cost production media by Gluconacetobacter xylinus. Journal of Polymers and the Environment, 21, 545-554. Yamamoto, H., Horrii, F., & Hirai, A. (2005). Structural studies of bacterial cellulose trough the solid-phase nitration and acetylation by CP/MAS 13C NMR spectroscopy. Cellulose, 13, 327342. Yuan, H., Nishiyama, Y., Wada, M., & Kuga, S. (2006). Surface acylation of cellulose whiskers by drying aqueous emulsion. Biomacromolecules, 7, 696-700.

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Figure 2. CP/MAS 13C NMR spectra of neat and esterified BNC samples. (a) Neat BNC; (b) Acetylated BNC – DS=0.22; (c) Propionized BNC – DS=0.18.

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Figure 3. FTIR spectra of neat and esterified BNC samples. Acetylated BNC (left, i): (a) Neat BNC; (b) DS=0.05; (c) DS=0.12; (d) DS=0.14; (e) DS=0.39; (f) DS=0.45. Propionized BNC (right, ii): (a) Neat BNC; (b) DS=0.06; (c) DS=0.09; (d) DS=0.12; (e) DS=0.23.

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Figure 4. X-ray diffraction pattern of neat and esterified BNC. (a) Neat BNC; (b) Propionized BNC - DS=0.09; (b) Propionized BNC - DS=0.23; (c) Acetylated BNC – DS=0.14; (d) Acetylated BNC – DS=0.45. For clarity purposes, baselines of XRD patterns have been moved up in the y axes.

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Figure 5. TG and DTG curves of neat and esterified BNC samples. (i) Acetylated BNC: (a) Neat BNC; (b) DS=0.05; (c) DS=0.12; (d) DS=0.14; (e) DS=0.22; (f) DS=0.39; (g) DS=0.45. (ii) Propionized BNC: (a) Neat BNC; (b) DS=0.02; (c) DS=0.06; (d) DS=0.09; (e) DS=0.12; (f) DS=0.18; (g) DS=0.23.

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Figure 6. Tonset (ºC) and Tmax (ºC) values calculated for neat and esterified BNC samples with increasing DS. Figure 7. Surface hydrophobicity assays. (a, d, g) Neat BNC; (b) Acetylated BNC – DS=0.14; (c) Acetylated BNC – DS=0.45; (e) Propionized BNC - DS=0.09; (f) Propionized BNC DS=0.23; (i) Acetylated BNC – DS=0.14; (e) Propionized BNC - DS=0.12.

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638 639 640 641 642 643 644 645 646 647

Figure 1. Organocatalytic esterification of bacterial nanocellulose using acetic acid and propionic acid as acylants, 120ºC.

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635 636 637

Figure captions

Table and Scheme Captions

Ac ce p

610 611 612 613 614 615 616 617 618 619 620 621 622 623 624 625 626 627 628 629 630 631 632 633 634

Table 1. Crystallinity index of neat and esterified BNC calculated by Newman’s (NMR) and Segal’s (XRD) method. Scheme 1. Proposed mechanism for the esterification of bacterial cellulose nanofibers with acetic and propionic acids catalyzed by L-tartaric acid.

17 Page 17 of 26

Sample Native BNC Acetylated BNC, DS= 0.14 Acetylated BNC, DS= 0.22 Acetylated BNC, DS= 0.45 Propionized BNC, DS= 0.09 Propionized BNC, DS= 0.18 Propionized BNC, DS= 0.23

CI NMR (%) 71.8 72.3 69.0 -

CI XRD (%) 91.8 90.1 91.9 92.3 90.7

Ac ce p

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649 650

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647 648

18 Page 18 of 26

Figure 1

Acetic acid

0.50

Propionic acid 0.45 0.40 0.35

DS

0.30 0.25

0.15 0.10 0.05 0.00

0

1

2

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4

5

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8

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Page 19 of 26

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Sheme 1

Page 20 of 26

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Figure 2

Page 21 of 26

Figure 3

i)

ii) (f)

(e)

Absorbance

(d) (c)

(c)

(b)

(b)

(a)

(a)

3900

(d)

3600

3300

3000

2700

2400

2100

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3900

-1

3600

3300

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Absorbance

(e)

2100

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1500

1200

900

-1

Wavenumber [cm ]

Ac

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Wavenumber [cm ]

Page 22 of 26

Figure 4

(002)

(101)

(101) (040)

(e)

(d)

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(c)

(b)

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40

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Page 23 of 26

Figure 5

i) (a)

100

(b) 90

(g)

(c) (d)

80

(f)

(e) (f)

(e)

(g)

60

DTG

m/m

0

[%]

70

50

(d)

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(c)

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(b)

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(a)

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(b) 90

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(f)

(e) 70

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(f) (e)

(g)

DTG

60 50

(d)

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[%]

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ii)

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350

Temperature [°C]

Temperature [°C]

m/m

300

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(b)

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Temperature [°C]

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Page 24 of 26

Figure 6

370

360

340

330

320

Neat BNC - Tonset Acetylated BNC - Tonset Propionized BNC - Tonset

310

Neat BNC - Tmax Acetylated BNC - Tmax 300

0.00

0.05

0.10

0.15

0.20

0.25

0.30

0.35

0.40

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0.50

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Figure 7

a)

b)

g)

c)

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e)

f)

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Page 26 of 26

Surface esterification of cellulose nanofibers by a simple organocatalytic methodology.

Bacterial cellulose nanofibers were esterified with two short carboxylic acids by means of a simple and novel organic acid-catalyzed route. The method...
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