Forming Nanospherical Cellulose Containers - Industrial

Aug 19, 2014 - Greenspan , P.; Fowler , D. S. Spectrofluorometric studies of the lipid probe, Nile Red J. Lipid Res. 1985, 26, 781– 789. [PubMed], [...
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Forming Nanospherical Cellulose Containers Oshrat Tzhayik, Indra Neel Pulidindi, and Aharon Gedanken* Department of Chemistry, Bar-Ilan University Center for Advanced Materials and Nanotechnology, Bar-Ilan University, Ramat-Gan 52900, Israel ABSTRACT: Nanospherical cellulose containers (SCC) (average size, ∼50 nm) are prepared by the sonochemical method. Organic and water-soluble materials are encapsulated within the SCC using only microcrystalline cellulose (MCC), an organic solvent, and water. Using silicotungstic acid (H4SiW12O40), the prepared SCCs can be catalytically converted (78 wt % conversion) to glucose (30 wt % glucose yield) by applying microwave irradiation for only 3 min. These results are attributed to the large increase of surface area of the cellulose sphere relative to MCC, and a better contact with the solid catalyst. Sealed in a vessel, SCCs were found stable for more than 6 months when stored at 4 °C or at room temperature. The encapsulation efficiency of the organic phase was measured and found to be approximately 90%. The creation of the sphere involves the degradation of the MCC to smaller fragments by high-intensity ultrasound irradiation, and the organization of these fragments in nanospheres involves the formation of bonds of the same type as those in regular cellulose (MCC). The structure and properties of SCCs were analyzed using high resolution scanning electron microscopy, Fourier transform infrared microscopy, 13C magic angle spinning NMR, X-ray diffraction, and fluorescence microscopy.

1. INTRODUCTION Cellulose, the most abundant organic, natural polymer on earth,1−4 is generally considered to be a plant material, but is also produced by some bacteria. It is a nontoxic, renewable, biocompatible,1,5 and biodegradable polysaccharide, consisting of stiff long linear chains, of repeating β(1 → 4) linked units of 3 D-glucose. Because of the strong hydrogen bonds between the cellulose chains, cellulose is insoluble in water and in most organic solvents, and this lack of solubility is a main problem facing cellulose science. Over the past decades, interest in sustainability and green chemistry has led to a renewed interest in novel cellulosic materials6 and composites7,8 derived from a variety of cellulosic components including cellulose whiskers (sometimes referred as cellulose crystals).9,10 Cellulose is the material of choice for the preparation of nanoparticles suitable for nanomedical applications especially in targeted drug delivery systems or contrasting agents in imaging techniques.5,11 The use of cellulose in medical devices has recently been intensified, mainly owing to its lack of immunostimulatory properties in vivo.3 Furthermore, pure cellulose nanoparticles are favored over synthetic polymers in many applications, such as nanopatterning in nanoreactions, for diverse immobilization methods and as separation materials.11 Numerous researchers have practiced acid hydrolysis of cellulose, and have successfully prepared spherical nanocrystalline cellulose (SNCC) from waste cotton fabrics,12,13 cotton linter and bleached linen,1 and cellulose fibers.2 Others have prepared spheres made of modified cellulose such as methyl cellulose14 and ethyl cellulose3,14,15 and have used them for drug delivery systems. Because of the distinct advantages of cellulose, there have been several attempts to create pure cellulose capsules. Nelson and Deng have shown that cellulose aggregates can uniformly encapsulate inorganic colloidal templates (TiO2), giving unique core−shell-structured materials.4 These core−shell particles can be further converted to hollow cellulose nanoparticles (20−150 nm) by dissolving the inorganic core.4 Libert et al. prepared © XXXX American Chemical Society

pure, perfectly spherical cellulose nanoparticles with sizes of ca. 80−260 nm by dialysis, starting from trimethylsilylcellulose (TMSC).5,11 In contrast to cellulose nanocrystals, fast cellular uptake is found for nanospheres, thus it seems that there is an influence of the geometry of the nanoparticle on endocytosis.5 Carrick et al. showed that it is possible to create a hollow cellulose capsule, by a precipitation method, using only native cellulose fibers, a cellulose solvent, and an inert gas to saturate the cellulose solution. This technique makes it possible to prepare hollow cellulose capsules with volumes of 10 μL.3 In the current study, using ultrasonic waves, we created new, pure cellulose nanosphere containers. Our aspiration was to use these nanospherical cellulose containers (SCC) in a catalytic process of converting cellulose to glucose, and to investigate the efficiency of this process expressed mainly in the glucose yield. Developing technologies that can convert cellulosic materials (often referred to as biomass) into motor fuels (ethanol) has been a worldwide goal of governments and private industries for the last three decades. While first generation ethanol technology converts effectively the starch portion of grain to sugar and then the sugar to ethanol, cellulose offers an economically preferable starting material. In every conversion process, the depolymerization of cellulose to glucose is a first step on the path to transportation fuels and chemical products.9 Recently, Zhang et al.9 showed that ultrasound technology is a powerful tool for enhancing the reactivity of microcrystalline cellulose (MCC) in the presence of a solid acid catalyst (AC− SO3H). Ultrasound pretreatment decreased dramatically the MCC particle size, leading to a better interaction with solid catalyst surfaces. As a result, the subsequent hydrolysis of MCC was found to be highly selective in producing water-soluble Received: July 1, 2014 Revised: August 6, 2014 Accepted: August 19, 2014

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inside the reaction cell was not higher than room temperature (measured by a thermocouple). 2.3. Synthesis of Spherical Cellulose Container (SCC). Dodecane-encapsulated spherical cellulose containers (SCC-D) were synthesized by the sonochemical method, as described elsewhere.23 In a reaction cell 20 mL of dodecane were layered over 30 mL of 1% (w/v) of sonicated-MCC. The volume ratio between the aqueous phase (cellulose) and the organic solvent (dodecane) was kept at 3:2 in all the experiments. The cellulose spheres were synthesized with a high-intensity ultrasonic probe (Sonics and Materials, VC-600, 20 kHz, 0.5in Ti horn). The bottom of the high-intensity ultrasonic horn was positioned at the aqueous−organic interface, employing an intensity of about 150 W cm−2, with an initial temperature of 20 °C in the reaction beaker. The reaction cell was immersed in an ice− water cooling bath during the sonication. The sonication lasted 3 min. At the end of the process the temperature in the reaction beaker reached ∼30 °C (measured by a thermocouple). The product was separated from the mother solution. The separation between the aqueous phase (at the bottom), the organic phase (the upper layer), and the SCC-D (intermediate layer) was accomplished within a few minutes due to the density differences between the spheres and the mother solution. To obtain a more complete separation of the spheres from the mother solution, the flask was placed in a refrigerator (4 °C) for ∼3 h. After the separation the product was stored at 4 °C. Moreover, dodecane was chosen only as a model organic compound. However, organic compounds, such as canola oil or perfluorohexane were also used and SCC was obtained. Using canole oil makes the SCC human compatible. Fluorescent-SCC-D (Nile Red) was produced by the same method, for this purpose 20 mL of dodecane stained with Nile Red was layered over 30 mL of 1% (w/v) sonicated-MCC. These SCCs were then dyed with Calcofluor White M2R (which binds to cellulose) in order to visualize the cellulose shell of the spheres. Finally the product was washed with deionized water. Another type of fluorescent-SCC-D (rhodamine) was also prepared: dodecane (20 mL) was layered over 30 mL of 1% (w/v) sonicated-MCC stained with rhodamine 6G. The obtained SCCs were than washed with deionized water. 2.4. Catalytic Hydrolysis of SCC under Microwave Irradiation. The hydrolysis of SCC-D containers was performed using a modified domestic microwave oven. A distillation column was inserted through the upper wall of the microwave oven for enhanced operation safety, and a magnetic stirrer that was added at the bottom of the microwave oven could be turned on or off during the reaction.24−26 The microwave oven operated at 2.45 GHz in a batch mode under air at atmospheric pressure. The output of the applied microwave reactor was 1100 W. In a typical procedure, 20 mL of SCC-D (∼0.2 g MCC), H4[Si(W3O10)4]·H2O (2 g) and a magnetic stirring bar were introduced into the reactor. The reaction was conducted with stirring for 3 min. In a control experiment, under the same hydrolysis conditions, the same amounts of the catalyst H4[Si(W3O10)4]·H2O (2 g), deionized water (20 mL), and MCC (0.2 g) were used. Control experiments were also carried out under conventional reflux conditions with stirring for 3 min. At the end of each experiment the reaction vessel was cooled to room temperature, the reaction mixture was filtered (filter paper), and the recovered aqueous solution (after separation from the organic

reducing sugars. Glucose was produced in yields similar (20 wt %) to those obtained using conventional pretreatment methods such as ball-milling and ionic liquids (and up to 42 wt % overall glucose yield in five catalytic cycles). The aim of the current study was to prepare nanometric pure cellulose containers from sonicated-MCC and investigate their conversion to glucose via a solid acid (H4SiW12O40) catalytic process. The idea was that spherical-shaped containers increase the available cellulose surfaces that can interact with the solid catalyst surface, thus increasing the conversion yield of cellulose to glucose. We encapsulated organic oils in pure, nonmodified cellulose nanospheres (∼50 nm diameter) using a very simple, inexpensive, and rapid, two-step ultrasonic method. In the first step we used ultrasound pretreatment on MCC (Avicel), and in the second step we have employed the well-known sonication emulsification methods using an aqueous solution of the sonicated-MCC coupled with organic oils to form cellulose spheres containing the oil in their inner volume. Ultrasonic emulsification is a well-known process that occurs in a biphasic system. Emulsification is necessary for the sonochemical capsule formation. Using this process proteinaceous microspheres16−18 as well as starch,19 DNA,20 RNA,21 and chitosan22 microspheres were prepared. However, if vortex mixing emulsification is used, no-long-lived capsules are formed. Consequently, emulsification by itself is not sufficient for sphere formation. Furthermore, the vortex emulsions are not stable and phase separation occurs. In contrast, when sonochemcial methods are applied the cellulose spheres obtained were stable for months either at room temperature or at 4 °C. Unlike previous work, in the current research, we did not use any cellulose solvent, modified cellulose, or inorganic materials in order to prepare the cellulose nanospheres containers. To the best of our knowledge, no one has, so far, used spherical nanocellulose containers for the conversion of cellulose to glucose, and this is the first time that nanospherical cellulose containers were formed solely by the ultrasonic method. From the viewpoint of green chemistry, using ultrasonic irradiation has noble advantages such as (1) the use of water as a solvent in the pretreatment stage, (2) short reaction time (≤2 h), (3) no need for an external source of heating, and (4) no polluting effluent is produced in the process.9 Thus, this research presents a new pathway for activating cellulose in a highly eco-efficient manner.

2. EXPERIMENTAL SECTION 2.1. Materials. Microcrystalline cellulose (MCC), Avicel PH-101; dodecane; Nile Red, P98.0% (HPLC); Calcofluor White M2R; poly-L-lysine solution; D-(+)-glucose; urea, 99.0− 100.5%; NaOH; rhodamine 6G, 95%; and tungstosilicic acid hydrate (H4[Si(W3O10)4]*H2O) were obtained from Sigma− Aldrich. All chemicals were used without any further purification. Deionized H2O was used in all the experiments. 2.2. Ultrasound Treatment of Microcrystalline Cellulose (Sonicated-MCC). A 1.5 g sample of MCC and 148.5 mL of deionized water (148.5 mL), as well as a magnetic stirrer, were placed in the sonication cell. This suspension was subjected to sonication employing an intensity of about 150 W cm−2 with a booster. The sonication lasted for 1.5 h yielding a highly stable, 1% (w/v) cellulose suspension (sonicated-MCC). During the sonication the reaction cell was kept in an ice− water cooling bath, in an attempt to keep the temperature constant at 25 °C. During the sonication, the temperature B

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Figure 1. (a) HRSEM image of MCC; (b) HRSEM image of SCC; (c) combined image of white light mode and fluorescence images of SCC-D synthesized in the presence of Nile Red (red signal); (d) DLS graph depicting the size distribution of the sonicated SCC-D.

dodecane phase) was analyzed using 13C NMR (Bruker Avance DPX 300 instrument using D2O as solvent). The conversion (wt %) was determined by the change in the cellulose weight before and after the reaction. The amount of cellulose after the reaction was determined by weighing the residue after filtration and drying for at least 12 h at 80 °C. The yield of glucose was calculated by an analytical method developed and reported recently by our laboratory.27 This method relates to the detection of glucose in the range of 0.04− 5.2 mM, with the aid of in situ-generated carbon nanoparticles (NPs) as sensor. The glucose sensing is based on UV-lightabsorbing carbon NPs with a specific absorbance at 275 nm. The filtrates were neutralized with aqueous solutions of NaOH, 2 M and 0.1 M. To 0.2 mL of each neutralized filtrate we added 1 mL of urea (10%) and 13.5 mL of deionized water. The mixtures were heated in an autoclave at 120 °C for 20 min during which time the carbon NPs were generated. For calibration, the same urea−autoclave treatment was applied to seven analytes containing varied concentrations of glucose. The UV−visible spectra of the glucose−urea samples after autoclave treatment were recorded on a Cary 100 scan Varian UV−vis spectrophotometer, and the yield of glucose was calculated. 2.5. Characterization. For imaging and characterization, high resolution scanning electron microscopy (HRSEM) measurements were conducted using a FEI XHR-SEM Magellan 400L scanning electron microscope operating at 5 kV. HRSEM samples were prepared either by applying powder

of lyophilized SCC-D onto a carbon tape, or by applying a drop of SCC-D containers onto a glass wafer covered with a poly-Llysine layer, and drying at room temperature. The samples were then vacuum-coated for 3 min with a thin carbon film. The average size and size distribution of the SCC-D containers were evaluated using HRSEM measurements, NanoSight LM20 Instrument, and DLS (dynamic light scattering) measurements. The DLS measurements were carried out on a Zetasizer Nano ZS series (Malvern Instrument) under 633 nm He−Ne laser. The SCC-D were analyzed and characterized by X-ray diffraction (XRD), 13C NMR measurements, 13C magic angle spinning (MAS) NMR measurements, Fourier transform infrared (FTIR) spectra, and light and fluorescence microscopy images. X-ray diffraction (XRD) patterns were collected on a Bruker D8 Advance X-ray diffractometer using Cu Kα operating at 40 kV/30 mA with a 0.02 step size and 1 s per step. 13C NMR measurements were conducted for aqueous phase components’ identification after hydrolysis. Each 13C NMR analysis was performed on a Bruker Avance DPX 300 instrument using D 2O as a solvent. 13 C MAS NMR measurements were performed on a Bruker Avance III 500 MHz spectrometer using a 4 mm double-resonance MAS probe. The FTIR spectra were recorded using Avarter model FT-IR spectrometer in the range of 4000−400 cm−1. Light and fluorescence microscopy images were obtained using an Axio imager Z1 microscope, Zeiss optical microscope. For light microscopy measurements, samples were prepared by depositC

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Figure 2. HRSEM images of lyophilized SCCs, magnified by 6000 (a); 40 000 (b); and 200 000 (c).

Figure 3. XRD patterns of MCC (black line) and SCC (red line).

diameter of 47 ± 3 nm (Figure 1d)) and by the Nanosight instrument (20−100 nm particle distribution with an average size of 41 ± 18 nm). The DLS results are in good agreement with the size determined by the HRSEM measurements. It may be concluded that these cellulose microspheres are actually aggregates of cellulose nanospheres. Moreover, it was found that the cellulose microspheres were unstable under the vacuum formed in the SEM chamber, or when they self-dried by placing them on a glass or freshly cleaved mica at ambient temperature. In both cases they collapsed. In contrast, SCCs are stable for more than 6 months when stored in a sealed bottle in the refrigerator (4 °C) or at room temperature. When lyophilized, the cellulose containers reveal a globular web of interwoven fibers with an approximate width of 40 nm and undefined length, as illustrated in Figure 2a−c. X-ray diffraction was used to better understand the developed cellulose nanosphere structure and to estimate the influence of the sonication on the nanosphere’s composition. Figure 3 displays XRD measurements of both MCC and SCC. Although the obtained cellulose nanosphere and MCC have similar diffraction profiles, some changes in the peak intensity occurred during the creation of the cellulose containers. The intensities of the SCCs XRD peaks are weaker than the MCC peaks. The SCCs did not retain the exact crystallinity as the initial MCC. The SCCs are more amorphous than MCC, and the ultrasonic effect has an essential role in reducing the crystalline nature of the sample. However, as concluded by Zhang and co-workers,9 these XRD profile changes indicate

ing the SCC-D dispersions onto a glass slide. For fluorescence microscopy, samples were prepared by depositing SCC-D onto a glass slide covered with poly-L-lysine layer. Fluorescence imaging was performed at three different excitation and emission ranges, using the blue fluorescent filter, “DAPI” (blue fluorescent signal; λex = 359 ± 25 nm, λem = 457 ± 30 nm) and two types of red fluorescent filters, “Texas Red” (red fluorescent signal; λex = 560 ± 25 nm, λem = 615 ± 20 nm) and “Rhod” (λex = 546 ± 12 nm, λem = 575−640 nm).

3. RESULTS AND DISCUSSION 3.1. Characterization of Spherical Cellulose Containers (SCC). The morphology of MCC and SCC-D was characterized using high resolution scanning electron microscopy (HRSEM) (Figure 1a,b, respectively). HRSEM analysis shows that MCC (Avicel), our staring material, was irregular in shape with ∼50 μm particle size (Figure 1a). In contrast, the SCC-Ds were formed as smooth surface spherical particles, with an average diameter (from 50 random particles) of 52 ± 3 nm (Figure 1b). This observation was supported by the DLS measurement which indicated that the cellulose containers have an average diameter of 47 ± 3 nm. As shown in Figure 1b the cellulose nanospheres have a tendency to aggregate, and after several days they create cellulose microspheres with a diameter in the range of 1−10 μm, as shown in Figure 1c. It should be noted that when the obtained microspheres are sonicated for ∼10 min they regained their original nanometer size as measured by DLS (average D

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Figure 4. FTIR spectra of MCC (black line) and SCC (red line).

Figure 5. 13C MAS NMR spectra of (a) MCC (black line) (b) cellulose spheres (SCC) (red line). Labels show the assignment of peaks to the different carbon atoms of the glucopyranose repeat unit.

cellulose spheres does not eliminate or weaken these hydrogen bonds. It appears that the sonication enables both the breaking and the formation of new H-bonds during the spheres’ formation. The peak at 2902 cm−1 in the MCC curve belongs to the stretching vibration of the C−H bonds of glucose units, whereas the bending vibration of these bonds absorbs at 1428 cm −1. Characteristic peaks at 1638 cm−1 for the MCC and 1640 cm−1 for the SCCs presumably correspond to the bending mode of water molecules that are tightly bound.28 In the MCC fingerprint region, 800−1200 cm−1, the spectral characteristics of MCC and SCCs are similar. The presence of all these detailed peaks confirms that the basic structural units of cellulose were not changed significantly by the sonochemical

that the sonication does not significantly affect the supramolecular ordering of MCC and that the ultrasound affects mostly the particle size of the cellulose chains. To elucidate the interfacial cross-linkage of cellulose molecules, IR spectroscopic experiments were performed (Figure 4). Figure 4 depicts the comparative FTIR spectra of MCC and developed cellulose nanoshperes (SCC). Overall there is no significant difference between the spectra. Characteristic stretching vibration of the cellulose alcohol groups that are H-bonded at 3333 cm−1 and at 3341 cm−1 appeared both for MCC and SCC. This indicates that hydroxyl groups and H-bonds exist before and after the cross-linking of the cellulose nanospheres, and that the formation of the E

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Figure 6. Fluorescence images of SCC-D that were synthesized in the presence of Nile Red and then colored with the blue fluorophore, Calcofluor White (a−c). (a) Fluorescence image using Texas Red filter (red signal); (b) fluorescence image using Dapi filter (blue signal); (c) combined image of images a and b; (d) combined white light mode image and fluorescence image using Rhod filter of SCC-D that were synthesized in the presence of rhodamine 6G.

filters (Figure 6a−c). Figure 6a presents the image obtained when we applied only the Texas Red filter. It clearly shows that the strong fluorescence of the red color is distributed homogeneously across the sphere, indicating that the sphere is loaded with dodecane containing Nile Red. Figure 6b presents the image that was revealed when we applied only the Dapi filter on the same group of SCCs. As expected, the labeled cellulose with the blue fluorophore is seen to spread homogeneously all over the surface of the droplets, forming a dense spherical cellulose layer, and the presence of the cellulosic shell is thus proven. It is assumed that the high stability of the cellulose layer may be attributed to the formation of hydrogen bonds between the hydroxyl groups of the cellulose fragments. Furthermore, the image in Figure 6c, taken when both the Texas red and the Dapi filters were simultaneously applied, shows spheres with a purple signal, as a result of an exact overlapping of the red and the blue signals, indicating that these spheres are indeed cellulosic spheres that encapsulate organic nonpolar liquid (dodecane). Further study involved rhodamine 6G, which is often used as a tracer dye in water to determine the rate and direction of flow and transport. When the aqueous phase was dyed with rhodamine 6G before sonication, and the Rhod filter was used on the obtained SCCs, the characteristic rhodamine’s red fluorescence emitted from the sphere’s core was observed (Figure 6d), suggesting a polar environment, obviously due to the presence of water molecules. In fact, the OH of the cellulose chains enables hydrogen

formation of SCCs and that the container’s shell consists of pure cellulose. We have used 13C MAS NMR to further investigate the nature of the bonds that form the cellulose sphere. Figure 5 depicts the spectra of both MCC and SCCs, and the peaks which have been assigned to the different carbon atoms of the glucopyranose repeating unit in cellulose are labeled. As can be seen, the spectra of MCC and of SCC are quite similar. This further supports the conclusion that the bonds that form the cellulosic sphere are the same bonds that exist in MCC, and a new type of bonds was not formed during the spherization process. In conclusion the XRD, FTIR, and the 13C MAS NMR measurements do not indicate the formation of new covalent bonds in the creation of the cellulose nanospheres. Further experiments were carried out in order to visualize the creation of the cellulosic shell and to investigate the nature of the core, that is, the inner side of the sphere. For this purpose, the fluorescent dye, Nile Red, was dissolved in the dodecane phase before sonication. Nile Red is an uncharged, heterocyclic molecule that strongly fluoresces bright red in hydrophobic environments (its solubility in water is negligible29,30). The obtained SCCs were then further dyed with the blue fluorophore, Calcofluor White, which binds specifically to cellulose, in order to visualize the cellulose shell of the spheres (blue signal). Fluorescent microscope images of these colored cellulose spheres were photographed using Texas Red and Dapi F

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Figure 7. Schematic representation of the SCC’s formation mechanism by ultrasonic waves. The symbol ))) corresponds to the ultrasonic irradiation.

ultrasound.31 In the first stage a stable radical at the anomeric carbon will be formed, as H· will leave this position. In the second stage, the hydroxyl radical (OH·) will replace the H· at the anomeric carbon, and finally this replacement will consequentially cause a break in the chain (Figure 7b). After this process, the obtained smaller fragments are again organized to the more thermodynamically stable conformations and spherical cellulose containers (SCCs) are formed (Figure 7c). 3.3. Catalytic Hydrolysis of SCC under Microwave Irradiation. Since ultrasound is capable of producing stable spherical cellulose containers, the catalytic hydrolysis of these spheres was then checked with the aim of evaluating the efficiency of such a pretreatment. In particular, the large increase of surface area of the cellulose sphere relative to MCC was expected to favor a better contact with the solid catalyst and thus to enhance the reactivity. Solid acid catalysts capable of promoting the hydrolysis of cellulose have recently received considerable attention because of their potential use at an industrial level.9 Among the developed solid acid catalysts, it was demonstrated that the keggin-type heteropolyacids (HPAs) such as H4SiW12O40 and H3PW12040 exhibit catalytic performances similar to dilute H2SO4 for cellulose transformation into methyl glucosides in methanol.33 These HPAs are promising solid acids and can replace environmentally harmful liquid acid catalysts such as H2SO4 with respect to corrosiveness, safety, quantity of waste generated, and separability.34 H3PW12040 was first employed to decompose cellulose under microwave irradiation (3 h). It was found that concentrated H3PW12040 (50%−80% w/w) could convert the cellulose and obtain a considerably high selectivity to glucose at low temperatures (80−100 °C), and with a glucose yield of 75.6%35 (see the

bonding with the water molecules. Thus, the presence of water molecules located at the core and on the surface of the SCCs is to be expected. Overall, these observations imply that the core of the cellulose containers is composed of a mixture of dodecane, water, and possibly some fragments of cellulose. Furthermore, the sonochemical method produce cellulosic spheres (SCCs) that encapsulate dodecane with a good loading efficiency. Typically, at the end of the sonication-encapsulation process a residue of not more than 2 mL of dodecane was obtained. Hence, the encapsulation, or loading efficiency, (defined as the weight percent of encapsulated dodecane related to the initial dodecane weight), is ∼90%. This number includes dodecane molecules present on and inside the spheres. 3.2. Proposed Mechanism for the Sonochemical Formation of SCC. Cellulose, like starch, is a polysaccharide consisting of glucose units that are held together by glycosidic bonds. The mechanism we propose for the sonochemical creation of SCCs is similar to that described for the sonochemical formation of starch microspheres by Grinberg and Gedanken.19 Upon sonication, a degradation of the MCC to smaller fragments occurs first, and then these fragments are organized around the collapsing bubble. Their low vapor pressure avoids their presence inside the bubble, and they arrange around the bubble in a spherical structure. During ultrasound irradiation of an aqueous solution the thermolysis of water molecules occurs due to the high temperatures created upon the collapse of the acoustic bubbles, and free radicals are formed31,32 (Figure 7a). The hydroxyl radicals that are produced recombine into H2O2, and the formed H2O2 and its sonolytic decomposition products have an effect on the degradation of organic compounds in the presence of G

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Figure 8. 13C NMR spectra of hydralyzate from (a) SCC and (b) MCC upon microwave irradiation for 3 min and in the presence of 2 g of H4SiW12O40.

experimental section for details). In this study we used H4SiW12O40 (10% w/w) as a solid acid catalyst under microwave irradiation for only 3 min in order to decompose cellulose spheres containers to glucose. 3.3.1. Characterization of the Hydrolysis Products. The 13 C NMR spectrum of the reaction product is depicted in Figure 8a. It can be seen that upon microwave irradiation for 3 min in the presence of H4SiW12O40, the characteristic peaks of glucose are observed, 61 ppm (C6), 70 ppm (C4), 72 ppm (C2β), 73 ppm (C2α), 74 ppm (C3), 76 ppm (C5), 92 ppm (C1α), and 96 ppm (C1β), indicating the complete conversion of cellulose to glucose (two stereoisomer). No other sugars, such as xylose, were detected in the products.36,37 In contrast, in the control experiments (done under exactly the same conditions as the cellulose sphere) apart from glucose, levulinic, and formic acid were also formed; these last two are decomposition products of glucose. As illustrated in the control experiments spectrum (Figure 8b), the peaks observed at 28 and 38 ppm, are typical of the methylene attached to the carboxyl group and of the methylene attached to the carbonyl group of levulinic acid, CH3COCH2CH2COOH. The peak at 29 ppm corresponds to the methyl group, and the peak at 178 ppm corresponds to the carboxyl group of levulinic acid. The peak observed at 166 ppm is typical of formic acid. The appearance of levulinic acid and formic acid is associated with the formation of hydroxymethyl furfural (HMF) from glucose, and its subsequent decomposition to levulinic acid and formic acid. No typical signals of HMF were seen in all the hydrolysate, implying that the decomposition of HMF to levulinic and formic acid is complete and fast under microwave irradiation. It has been shown clearly that upon solid acid hydrolysis (H4SiW12O40) in a time period as short as 3 min, cellulose spheres yielded exclusively glucose, and no other C5 or C6 sugars. The later are inevitably formed when cellulosic biomass is employed as a feedstock for the generation of sugars. 3.3.2. Evaluation of the Cellulose Conversion and the Glucose Yield. In the first set of experiments, MCC was tested in order to highlight the contribution of the sphere’s creation via sonication in the activation of MCC. As shown in Table 1, without any activation the conversion of MCC is 70% while the glucose yield did not exceed 15.3%. Moreover, along with glucose, the production of leuvlinic and formic acid was also detected by C13 NMR, as depicted in Figure 8b. Then, the

Table 1. Catalytic Hydrolysis of SCC and MCC over H4SiW12O40 Catalysta sample

conversion (wt %)

yield of glucose (wt %)

cellulose sphere (SCC) MCC

78 70

30 15.3

a The experimental error of these values is ±10%. The initial weight of cellulose is 0.2 g.

catalytic hydrolysis of SCCs was investigated under identical conditions. Starting from SCC-D (Figure 9c), the conversion

Figure 9. Pictures of the hydralyzate from (a) MCC and (b) SCC; (c) picture of SCC-D.

was 8% higher than from MCC, indicating that the spheres formation led to a better contact between cellulose and the solid catalyst. The catalytic hydrolysis of SCCs also led to a significant increase in glucose yield, and in fact the glucose yield in the case of SCC (30 wt %) was nearly twice as high as the glucose yield from MCC (15.3 wt %). Interestingly, for MCC the aqueous phase color turned dark-brown (Figure 9a), whereas for SCC it turned clear bright-yellowish (Figure 9b,). Glucose formation from MCC was always accompanied by a considerable formation of furan derivatives or soluble humins. The latter give a dark-brown color to the reaction medium.9 In our case, the recovery of a bright-yellowish aqueous phase may suggest that unsaturated products are not produced. This observation is supported by the 13C NMR spectra analysis of the aqueous phase, which did not reveal the presence of any unsaturated products, and detected only glucose (Figure 8a).

4. CONCLUSIONS In this study, it was shown that it is possible to create a completely new material, nanospherical cellulose containers H

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(∼50 nm), by the sonochemical method, using only microcrystalline cellulose (MCC), an organic solvent (such as dodecane), and water. This makes sonication an efficient and easy way to encapsulate organic materials and water-soluble materials within pure cellulose nanocapsules. We have shown that using H4SiW12O40, these SCCs can be catalytically converted (78 wt % conversion) to glucose (30 wt % glucose yield) by applying microwave irradiation for only 3 min. To the best of our knowledge, this is one of the highest yields of glucose ever obtained from MCC, in one catalytic cycle, in the presence of a solid catalyst in water. Glucose can be fermented readily by Saccharomyces cerevisiae to ethanol, and thus the present findings may be of great importance in the motor-fuelethanol industry. More work is needed to establish a solid relationship between the SCC concentration and the microwave irradiation period, the catalyst type, the conversion, and the glucose yield. In addition, sonochemically prepared starch microspheres, should be investigated for the catalytic hydrolysis process.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS A.G. is thankful to the ISF (Israel Science Foundation) for supporting this research through Grant No. 598/12 and to the Ministry of Science for supporting this research through a Korea−Israel Grant No. 3-9802.

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