Towards Biomimicking Wood: Fabricated Free-standing Films of Nanocellulose, Lignin, and a Synthetic Polycation

Summary

The objective of this research was to form synthetic plant cell wall tissue using layer-by-layer assembly of nanocellulose fibrils and isolated lignin assembled from dilute aqueous suspensions.  Surface measurement techniques of quartz crystal microbalance and atomic force microscopy were used to monitor the formation of the polymer-polymer nanocomposite material.

Abstract

Woody materials are comprised of plant cell walls that contain a layered secondary cell wall composed of structural polymers of polysaccharides and lignin. Layer-by-layer (LbL) assembly process which relies on the assembly of oppositely charged molecules from aqueous solutions was used to build a freestanding composite film of isolated wood polymers of lignin and oxidized nanofibril cellulose (NFC). To facilitate the assembly of these negatively charged polymers, a positively charged polyelectrolyte, poly(diallyldimethylammomium chloride) (PDDA), was used as a linking layer to create this simplified model cell wall. The layered adsorption process was studied quantitatively using quartz crystal microbalance with dissipation monitoring (QCM-D) and ellipsometry. The results showed that layer mass/thickness per adsorbed layer increased as a function of total number of layers. The surface coverage of the adsorbed layers was studied with atomic force microscopy (AFM). Complete coverage of the surface with lignin in all the deposition cycles was found for the system, however, surface coverage by NFC increased with the number of layers. The adsorption process was carried out for 250 cycles (500 bilayers) on a cellulose acetate (CA) substrate. Transparent free-standing LBL assembled nanocomposite films were obtained when the CA substrate was later dissolved in acetone. Scanning electron microscopy (SEM) of the fractured cross-sections showed a lamellar structure, and the thickness per adsorption cycle (PDDA-Lignin-PDDA-NC) was estimated to be 17 nm for two different lignin types used in the study. The data indicates a film with highly controlled architecture where nanocellulose and lignin are spatially deposited on the nanoscale (a polymer-polymer nanocomposites), similar to what is observed in the native cell wall.

Introduction

There is great interest to derive additional chemicals and fuels from biomass, as carbon sequestered by plants during photosynthesis is part of the current CO₂ cycle. The majority of sequestered carbon (42-44%) is in the form of cellulose, a polymer composed of β 1-4-linked glucopyranose units; when hydrolyzed, glucose can be used as the primary reactant for fermentation into alcohol based fuels. However, cell wall architecture of woody plants has evolved for millennia creating a material that is resistant to degradation in the natural environment¹. This stability carries over into the industrial processing of woody materials such as energy crops making cellulose difficult to access, isolate, and breakdown into glucose. A closer look at the ultrastructure of the secondary cell wall reveals that it is a polymer nanocomposite composed of layered paracrystalline cellulose microfibrils embedded in an amorphous matrix of lignin and hemicelluloses^2-4. The longitudinally oriented cellulose microfibrils have a diameter of approximately 2-5 nm, which are aggregated together with other hetero-polysaccharides to form larger units of fibril bundles⁵. The fibril bundles are embedded in a lignin-hemicellulose complex composed of an amorphous polymer of phenylpropanol units with some linkages to other hetero-polysaccharides like glucoronoxylan⁴. Furthermore, this structure is further organized into layers, or lamellae, throughout the lignified secondary cell wall^6-8. Enzymes, like cellulases, have a very difficult time accessing cellulose within the cell wall as it is found in its fibril form and embedded in lignin. The crux of truly making biobased fuels and renewable chemical platforms a reality is to develop processes that economically allow the saccharification of cellulose in its native form.

New chemical and imaging technologies are aiding in the study of the mechanisms involved in the saccharification of cellulose^9,10. Much work has centered on Raman confocal imaging¹¹ and atomic force microscopy¹² to study the cell wall chemical composition and morphology. Being able to closely follow mechanisms of delignification and saccharification is a significant step forward, impacting the conversion of cellulose to glucose. Saccharification of model cellulose surfaces was analyzed by measuring enzyme kinetic rates with a quartz crystal microbalance with dissipation monitoring (QCM-D)¹³. However, native cell walls are highly complex as indicated above, and this creates ambiguity of how different conversion processes change the structure of the plant cell wall (polymer molecular weight, chemical linkages, porosity). Free-standing models of the cell wall substances with known structural composition would address this concern and allow the integration of samples into state-of-art chemical and imaging equipment.

There is a dearth of cell wall models and the few available can be categorized as blends of polymer materials and regenerated cellulose or bacterial cellulose¹⁴, enzymatically polymerized lignin-polysaccharide composites^15-17, or model surfaces^18-21. Some models that begin to resemble the cell wall are the samples that contain lignin precursors or analogs polymerized enzymatically in the presence of cellulose in its microfibrillar form. However, these materials suffer from the lack of organized layer architecture. A simple route for the creation of nanocomposite materials with organized architecture is the layer-by-layer (LbL) assembly technique, based on the sequential adsorption of polymers or nanoparticles with complementary charges or functional groups to form organized multilayered composite films^22-25. Free-standing hybrid nanocomposites of high strength, made by LbL deposition of polymer and nanoparticles, have been reported by Kotov et al.^26-30. Among many other applications, LbL films have also been investigated for their potential use in therapeutic delivery³¹, fuel cell membranes^32,33, batteries³⁴, and lignocellulosic fiber surface modification^35-37. The recent interest in nanoscale cellulose based composite materials have led to the preparation and characterization of LbL multilayers of cellulose nanocrystals (CNC) prepared by sulfuric acid hydrolysis of cellulose fibers, and positively charged polyelectrolytes^38-43. Similar studies have also been conducted with cellulose nanocrystals obtained from marine tunicin and cationic polyelectrolytes⁴⁴, CNC and xyloglucan⁴⁵, and CNC and chitosan⁴⁶. LbL multilayer formation of carboxylated nanofibrillated celluloses (NFCs), obtained by high-pressure homogenization of pulp fibers with cationic polyelectrolytes has also been studied^47-49. The preparation, properties, and application of CNCs and nanofibrillated cellulose have been reviewed in detail^50-53.

The present study involves the examination of LbL technique as a potential way to assemble isolated lignocellulosic polymers (such as nanocellulose and lignin) in an ordered fashion as the first step towards a biomimetic lignocellulosic composite with lamellar structure. The LbL technique was selected for its benign processing conditions such as, ambient temperature, pressure, and water as the solvent, which are conditions for natural composite formation⁵⁴. In this study we report on the multilayer build-up of constitutive wood components, namely cellulose microfibrils from the tetramethylpiperidine 1-oxyl (TEMPO) mediated oxidation of pulp and isolated lignin into free-standing lamellar films. Two different lignins are used from different extraction techniques, one a technical lignin from the organosolv pulping process, and the other a lignin isolated from ball-milling with less modification during isolation. These compounds are combined with a synthetic polyelectrolyte in this initial study to demonstrate the feasibility of making stable free-standing films with architecture similar to the native cell wall.

Protocol

1. Nanofibrillated Cellulose Preparation⁵⁵

1. Setup a 3 L three-neck flask with 2 L of deionized water, an overhead stirrer, and pH probe.
2. Add delignified kraft pulp, 88% brightness (20 g, 1% (w/v, dry weight basis)), 2,2,6,6-tetramethylpiperidine 1-oxyl (TEMPO) (0.313 g, 0.1 mmol/g cellulose), and sodium bromide (NaBr, 2.0 g, 1 mmol/g cellulose) to the flask.
   1. Mix the pulp fiber with the overhead stirrer until the fiber is dispersed and no aggregates can be seen in the reaction.
      Note: Dispersion can be aided by blending the slurry in water prior to adding the pulp to the 3 L flask.
3. Initiate the oxidation by slowly adding a 12% solution of sodium hypochlorite (NaClO, 51.4 ml, 5 mmol per g of cellulose) to the reaction mixture.
   Note: For consistency throughout the reaction, use a syringe pump to deliver the NaClO with an injection rate of 1.5 ml/min.
4. Fill a second syringe with sodium hydroxide (NaOH, 0.5 M) and manually meter the alkali solution into the flask drop-wise to maintain the pH at 10 ± 0.2.
5. Monitor the change in pH with time and once all the accessible hydroxyl groups on the cellulose are oxidized the pH will no longer decrease and the reaction is complete.
6. Add excess EtOH to consume remaining NaClO. Approximately 6 ml of 200 proof EtOH will consume all 100 mmol of the original NaClO.
7. Filter and wash the oxidized fiber thoroughly with purified water to remove the reagents until pH is neutral. Use a basket centrifuge or some filtration device like a Büchner funnel to recover the fiber. Store the fiber at 4 °C until further use.
   Note: At the completion of the experiment, the fiber should have a carboxylic acid content, as determined by conductometric titration, between 1.0 to 1.5 mmol per gram of fiber. There should be little difference in appearance of the fiber after the TEMPO oxidation.
8. Create a 3% (w/v, dry weight basis) slurry of the TEMPO oxidized pulp and blend in a Warring blender until the slurry becomes viscous and the blades start spinning in air because of gelling of the suspension.
   Note: Lower concentrations do not work as effectively to fibrillate the cellulose.
   1. Dilute the blended slurry to 0.1% (w/v) and continue blending until suspension becomes transparent.

2. Layer-by-layer Film Deposition for QCM-D Experiments

1. Prepare the following aqueous solutions and adjust each solution with 0.1 M NaOH to a pH of 10.5: aqueous buffer solution (water and NaOH); 0.5% (w/v) aqueous solution of polydiallyldimethylammonium chloride (PDDA); and 0.01% (w/v) lignin. Adjust the pH of the 0.1% NFC suspension to 8.0.
   Note: The pH for these experiments was elevated because it was previously shown that lignin adsorbs in a less aggregated state in alkaline pH⁵⁶.
2. Clean a gold-coated quartz crystal following manufacturers recommendation of using a base piranha solution [CAUTION] (3:1 concentrated NH₄OH:H₂O₂ at 60 °C) for 10 min.
   1. Rinse the crystals with purified water, blow dry in a stream of N₂, and immediately insert into the quartz crystal microbalance flow cell to avoid contamination from the air.
3. Pass the buffer through the flow cell to obtain a baseline response of the resonating crystal exposed to the liquid.
   1. Deposit a layer of PDDA onto the quartz crystal by exposing the quartz crystal to the PDDA solution for 5 min.
   2. After 5 min switch back to the buffer solution.
      Note: This process in step 2.3 creates a single layer response where the amount of polymer deposited can be determined without the effect of the polymer solution viscosity.
   3. Repeat the adsorption of other polymers in the following sequence with a buffer rinse between each step: PDDA (+) (step 2.3.1); lignin (-); PDDA(+); and NFC(-). Repeat the cycle 4x to deposit 16 total layers of polymers and nanoparticles.

3. Layer-by-layer Film Deposition for AFM and Ellipsometry Experiments

1. Glue a circular disc of mica to a glass microscope slide using a quick epoxy adhesive. After the adhesive cures, attach a piece of tape to the mica disc. Peel the tape away causing the mica surface to cleave.
   1. Clean a silicon wafer with acid piranha [CAUTION] (3:1 H₂SO₄:H₂O₂) for 20 min followed by significant rinsing in water prior to layer deposition.
2. With solutions prepared in 2.1, dip either the freshly cleaved mica that is attached to a glass slide or a freshly cleaned silicon wafer in each solution following the same sequence of protocol outlined in 2.3.3.
   Note: This technique will create layers of polymers on each of these surfaces that can be inserted into the AFM or ellipsometer, respectively.
3. Image the deposited layers with an atomic force microscope. Use the intermittent contact mode and cantilevers with 10 nm radius silicon tips (spring constant 42 N/m) when collecting images of the sample. Set scan size as 2.5 x 2.5 μm, scan point as 512 and integral gain of 10 to collect specific sample images.
4. For thickness measurement of the layers with AFM of the dried LbL films, use a soft plastic pipette tip and scar a line across the surface of the prepared LbL films on the mica surface.
5. Deposit LBL films for ellipsometry measurement onto silicon wafers. Measure the dry film thickness with a phase modulated ellipsometer at a wavelength of 632.8 nm using the multiple angle of incidence mode. Vary the angles between 85° and 65° at 1° intervals.

4. Preparation of Free-standing LBL Film

1. Cut a 25.4 x 7.6 mm rectangle of cellulose acetate (CA) film (DS 2.5) that is 0.13 mm thick and attach to automated dipper arm.
   Note: Cellulose acetate of DS 3.0 is not soluble in acetone so DS 2.5 is preferred to recover the layered films.
2. Fill each 500 ml beaker with solutions of PDDA, lignin, and nanocellulose according to concentration and pH in step 2.1.
   1. Fill three additional beakers with aqueous buffer to use as a rinse solution for each deposition cycle.
   2. Program the dipper arm to proceed in same sequence as reported in 2.3.3.
      Note: It is important to use a different rinse solution after each respective polymer solution because in the layer-by-layer process, some polymer that is not tightly bound to the surface will desorb. Cross contamination of the rinse solutions quickly causes precipitation of polyelectrolyte complexes, which can adsorb as "defects" to the film surface.
3. Change the solutions in the beaker during 250 cycles periodically as they begin to appear cloudy because of colloidal complexes. An option is to automate the renewal of the solution by using peristaltic pump to deliver fresh solution or buffer to custom made polyvinylchloride (PVC) containers with inlets and outlets.
   Note: Agitated solutions in the containers help enhance the diffusion of the polyelectrolytes to the surface.
4. Carefully trim the edges of the dried sample with scissors exposing the CA edge and place into a covered glass Petri dish filled with acetone to dissolve the CA.
   Note: Two films are isolated after this experiment from the front and backside of the CA.
5. Soak isolated films in acetone for 24 hr and rinse films repeatedly with acetone to maximize the removal of residual CA.

Results

QCM-D Analysis of Structured Woody Polymer Film Fabrication

The LbL adsorption of lignin, NFC and PDDA was monitored in real-time with QCM-D in two different experiments involving two types of lignins. This analysis method is very sensitive to detect changes in frequency when molecules adsorb to the surface of the quartz crystal. Figure 1 contains a detailed description of the QCM-D response in one deposition cycle, which involves two bilayers (PDDA:HMWL and PDDA:NC). The data re...

Discussion

Fabrication of Nanocellulose

For nanocellulose fabrication the successful oxidation of the pulp fiber is necessary for facile fibrillation. Oxidation is controlled by available sodium hypochlorite, which should be slowly added at known quantities based on the amount of cellulose. One reason for limited oxidation arises from the storage of the sodium hypochlorite solution for extended periods. This reduced oxidation efficiency can be noted during the reaction; the pulp slurry should turn a pale-ye...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Sulfate pulp	Weyerhaeuser	donated	brightness level of 88%
Organosolv lignin	Sigma Aldrich	371017	discontinued
Hardwood milled wood lignin	see reference in paper
Polydiallyldimethylammonium chloride	Sigma Aldrich	409022	Mn = 7.2 x 104, Mw = 2.4 x 105
2,2,6,6-Tetramethylpiperidine 1-oxyl (TEMPO)	Sigma Aldrich	214000	catalytic oxidation of primary alcohols to aldehydes with a purity of 98%, molecular weight is 156.25 g/mol
Sodium bromide	Sigma Aldrich	S4547	purity ≥99.0%, molecular weight 102.89
Sodium hypochlorite	Sigma Aldrich	425044	reagent grade, available chlorine 10~15%, molecular weight 74.44 g/mol
Sodium hydroxide	VWR	BDH7221-4	0.5 N aqueous solution, density 1.02 g/ml, molecular weight 40 g/mol
Sodium hydroxide	Acros Organics	AC12419-0010	0.1 N aquesous solution, specific gravity 1.0 g/ml, molecular weight 40 g/mol
Ammonium hydroxide	Acros Organics	AC39003-0025	25% solution in water, pH 13.6, density 0.89, molecular weight 35.04 g/mol
Hydrogen peroxide	Fisher Scientific	H325-100	30.0~32.0% certified ACS, pH 3.3, density 1.11
Mica sheets	TED Pella	NC9655733	Pelco, grade V5, 10 x 40 mm, 23 mm T, minimum air and bubbles, very clean
Sulfuric acid	Fisher Scientific	A300-212	95.0~98.0 w/w%, certified ACS plus, molecular weight 98.08 g/mol
Cellulose acetate	McMaster Carr	8564K44	degree of substitution 2.5
Ethanol	Decon Laboratories	04-355-223	200 proof (100%), USP
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Acetone	Fisher Scientific	A18-4	purity ≥99.5%, certified ACS reagent grade, density 0.79 g/ml, molecular weight 58.08 g/mol
Syringe pump	Harvard Apparatus	552226	pump 22 infusion/withdraw with standard syringe holder, flow rate 0.002 μl/hr~55.1 ml/min
Mill-Q water purification system	EMD Millipore	D3-UV	Direct-Q, UV, water conductivity 18.5 MΩ·cm with 20 L reservoir
pH meter	Mettler Toledo		SeverMulti
Balance	Mettler Toledo	AB135-S	accuracy 0.1 mg
Atomic force microscope	Asylum Research		MFP-3D, Olympic fluorescent microscope stage
Ellipsometer	Beaglehole Instruments
Fiber centrifuge	unknown		basket style centrifuge
Waring blender	Waring		Commercial
Ultrasonic processor	Sonics		Sonics 750 W, sound enclosure
Quartz crystal microbalance with dissipation monitoring (QCM-D)	Q-Sense Inc.	E4	measure fundamental frequency of 5 MHz, and monitor odd number overtones/harmonics from 3~13, use gold-coated piezoelectric quartz crystals
Automatted dipper arm	Lynxmotion