Comparison of Hydrogels Prepared with Ionic-Liquid-Isolated vs

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Comparison of Hydrogels Prepared with Ionic Liquid-Isolated vs. Commercial Chitin and Cellulose Xiaoping Shen, Julia L. Shamshina, Paula Berton, Jenny Bandomir, Hui Wang, Gabriela Gurau, and Robin D. Rogers ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.5b01400 • Publication Date (Web): 15 Dec 2015 Downloaded from http://pubs.acs.org on December 21, 2015

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ACS Sustainable Chemistry & Engineering

Comparison of Hydrogels Prepared with Ionic Liquid-Isolated vs. Commercial Chitin and Cellulose Xiaoping Shen,†,‡ Julia L. Shamshina,§ Paula Berton,†,║ Jenny Bandomir,† Hui Wang,† Gabriela Gurau§,║ Robin D. Rogers*,†,║ †

Department of Chemistry, The University of Alabama, Tuscaloosa, AL 35487, USA

‡

Key Laboratory of Bio-based Material Science and Technology (Ministry of Education),

Northeast Forestry University, 26 Hexing Road, Harbin 150040, China §

525 Solutions, Inc., 720 2nd Street, Tuscaloosa, AL 35401, USA

║

Department of Chemistry, McGill University, 801 Sherbrooke St. West, Montreal, QC H3A

0B8, Canada

ACS Paragon Plus Environment


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ABSTRACT

Physical and/or covalently-linked (chemical) hydrogels were prepared from chitin and cellulose extracted with ionic liquid from shrimp shells and wood biomass, respectively, and compared with hydrogels prepared from commercially available biopolymers, practical grade chitin, and microcrystalline cellulose. The highly porous aerogels were formed by initial dissolution of the biopolymers in NaOH/urea aqueous systems using freeze/thaw cycles, followed by thermal treatment (with or without epichlorohydrin as a cross-linker) and supercritical CO2 drying. The ionic liquid-extracted cellulose pulp and chitin, as well as practical grade chitin could form both stable physical and chemical hydrogels, whereas biopolymers of lower apparent molecular weight such as microcrystalline cellulose required a covalent cross-linker for hydrogel formation and commercially available pure chitin was not suitable for the preparation of hydrogels of either type. Hydrogels prepared from the ionic liquid-extracted biopolymers exhibited properties substantially different from those made from the commercially available biopolymers. Loading of an active ingredient into the hydrogel and its subsequent release was demonstrated using indigo carmine and revealed that the release rate was controlled mainly by the biopolymer concentration of the gel network.

KEYWORDS: Hydrogel, aerogel, chitin, cellulose, ionic liquid, release


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INTRODUCTION Hydrogels are highly swollen, hydrophilic, three-dimensional (3D) polymeric networks capable of absorbing large amounts of water or biological fluids and swelling.1 Currently, the majority of the hydrogels are manufactured using (meth)acrylic acid or acrylamide, compounds which raise public health concerns. 2 , 3 Considering not only safety, but also biocompatibility, tissuemimicking consistency, and biodegradability, biopolymer-based hydrogels have been drawing increasing interest and attention. 4 Cellulose and chitin, the two most abundant renewable biopolymers on earth, consist of straight chain of β-(1→4)-linked D-glucose, and β-(1→4)linked N-acetyl-D-glucosamine, respectively. Because of the hydroxyl and/or acetamido groups present in their structures, fused or amorphous cellulose and chitin chains are highly hydrophilic and potentially suitable to form superabsorbent hydrogels.5,6 In addition to their biodegradability,7,8 swollen hydrogels from cellulose and chitin are elastic but soft and possess diffusive properties identical to those of a liquid, and thus can absorb and release water or biological fluids in a reversible non-irritating manner, making them compatible with living tissues. 9, 10 Consequently, these hydrogels have attracted tremendous attention as excellent candidates for biomedical applications, such as drug delivery,11 wound dressings,12 and tissue engineering matrices.13 Chitin and cellulose hydrogels are usually prepared through a two-step process involving biopolymer dissolution and chain cross-linking. To date, there are only a few specific solvent systems reported to dissolve chitin or cellulose and form hydrogels, such as ionic liquids (ILs), 14 , 15 alkali or alkali/urea (thiourea) aqueous systems, 16 - 18 and polar solvent systems including LiCl/N-methyl-2-pyrolidone (NMP),8,

19

LiCl/N,N-dimethylacetamide (DMAc),

20

LiCl/dimethyl sulfoxide (DMSO), 21 and saturated CaCl2•2H2O/methanol. 22 Among these


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solvents, alkali/urea aqueous systems are attractive due to their low cost, energy consumption, and toxicity, and the relatively low viscosity of the resulting solutions. After dissolution, physical or chemical cross-links are needed to provide structural integrity and thus form hydrogels. Physical cross-links involve various interactions such as chain entanglements, strong van der Waals forces, hydrogen bonds, crystallite associations,23 and/or ionic interactions. 24 Chemical cross-links are covalent bonds between polymeric chains themselves or between the polymer(s) and a cross-linker added to gelate the solution.25 Physical hydrogels can be obtained from solution either by curing at a certain temperature,16,18 or by coagulating with an anti-solvent in a specific fashion (e.g., molding, casting, or dropping).19,26 Chemical hydrogels can be obtained via cross-linking using irradiation (e.g., electron beam,27 γray, 28 or UV 29 ) or using a chemical cross-linker (e.g., epichlorohydrin,9, 30 , 31 1,2,3,4butanetetracarboxylic dianhydride,8,32 or succinic anhydride7,33). In our previous work, we have developed a series of methods for the isolation of cellulose-rich materials (CRMs)34-36 and chitin (IL-chitin)37,38 by direct dissolution and reconstitution of the biopolymers from their corresponding biomass using ILs. It was found that these IL-isolated biopolymers had particularly higher molecular weights (MWs) than most commercial sources. We were curious whether the unique forms of the biopolymers we extracted from biomass using ILs would provide hydrogels with properties that might differ from the currently available materials. To test this, we utilized these extracted biopolymers for the formation of both physical and chemical hydrogels in NaOH/urea as the solvent system and compared their properties to analogous gels prepared from commercial cellulose and chitin and to hydrogels published in the literature. The formation, structure and morphology, water absorbency (rehydration), and active ingredient loading and release of these hydrogels are described here.


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RESULTS AND DISCUSSION Biopolymer Extraction and Dissolution in NaOH/urea. Chitin and cellulosic biopolymers used for this study were respectively extracted from shrimp shells (SS) and poplar, using [C2mim][OAc] and procedures we have previously described (See ESI for complete experimental details and composition of the biopolymers, Table S1).35,36,38 The materials studied here include IL-chitin (isolated from dried/processed SS using microwave-assisted dissolution38), CRM-175 (extracted from poplar at 175 °C for 30 min followed by regeneration with 50% v/v acetone/H2O35), and CRM-POM (extracted from poplar with polyoxometalate (POM) at 110 °C for 2 h followed by regeneration using 50% v/v acetone/H2O36). Commercial microcrystalline cellulose (MCC), pure chitin, and practical grade (PG)-chitin, were purchased from commercial suppliers and used for comparison. The literature has reported that commercial cellulose and chitin biopolymers can be dissolved in aqueous NaOH/urea solvent systems utilizing freeze/thaw (F/T) cycles.16,17,39,40 In NaOH/urea systems, the concentration of NaOH is reported to greatly affect biopolymer dissolution due to the ability of NaOH to break hydrogen bonds, while urea (generally above 4 wt%) is suggested to act as hydrogen bond donor, creating cellulose inclusion complex and preventing reaggregation of the biopolymer chains. Here, we verified that F/T treatment is also a suitable approach for the dissolution of IL-extracted biopolymers in NaOH/urea. To accelerate dissolution, we adopted an ethanol/dry ice bath for freezing instead of a refrigerator as noted in the literature. We first attempted to replicate this dissolution procedure for the IL-extracted biopolymers using 8 wt% NaOH/4 wt% urea and 7 wt% NaOH/12 wt% urea aqueous solutions; systems reported to dissolve chitin and cellulose, respectively. 41,42 We found (Table 1) that different


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NaOH concentrations and numbers of F/T cycles were required to dissolve the biopolymers. For chitin, a minimum of 7 wt% NaOH in NaOH/urea was required for its dissolution, while for cellulose, only the 4‒8 wt% NaOH concentrations were able to dissolve MCC and CRM-POM, and 4‒6 wt% to dissolve CRM-175. The optimized conditions were chosen as those in which the biopolymer was dissolved using the least number of F/T cycles: 8 wt% NaOH/4 wt% urea for chitin, 6 wt% NaOH/4 wt% urea for MCC and CRM-POM; and 4 wt% NaOH/4 wt% urea for CRM-175. Table 1. Alkali Solvent Selection, Viscosity Values of the Resulting Solutions, and the Maximum Concentration of the Biopolymer in the Optimum Solvent System.

Chitin

Biopolymer

Cellulose

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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NaOH/urea x wt%/y wt% (F/Ta cycles) 2/2 4/0

4/4

6/4

7/12

8/4 +(2); √d +(3); √ +(3); √ +(4); √

Purechitin PGchitin Rec. PGe ILchitin

‒c

‒

‒

Swc

+(4)c

‒

‒

‒

Sw

+(8)

‒

‒

‒

Sw

+(6)

‒

‒

‒

Sw

+(8)

MCC

‒

‒

+(6)

+(2); √ +(3); √

+(2)

+(4)

Max. conc.i (wt%)

12/6b

20/0b

Viscosityf (cP)

+(2)

+(2)

5.0g

12.5(3)

+(3)

+(3)

14.1(8)h

4.3(2)

+(3)

+(3)

8.4(2)

6.5(2)

+(4)

+(4)

18.8(2)

2.4(2)

‒

‒

8.9(1)

5.2(2)

CRM‒ ‒ +(8) +(6) +(6) ‒ ‒ 9.4(1) 5.4(2) POM CRM+(4); ‒ ‒ +(10) Sw Sw ‒ ‒ 31.0(4) 1.4(2) 175 √ a F/T: Freeze/thaw; b These conditions for chitin might result in chitosan formation; c ‒: insoluble; +: soluble; Sw: insoluble but can swell; d The optimized solvent for each biopolymer; e Rec. PG: Reconstituted PG-chitin from the [C2mim][OAc] solution to remove impurities (100 °C, 1 h; ESI); f Viscosity of 1 wt% solution in NaOH/urea at 20 °C (Fig. S2); g The error was lower than 0.1; h The large error was caused by insoluble impurities in the PG-chitin solution; i Max. conc.: maximum concentration in the optimized NaOH/urea system.


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It is noteworthy that using any given NaOH/urea system, the number of F/T cycles required depended on the biopolymer, e.g., IL-chitin needed more F/T cycles than pure chitin in the same NaOH/urea solution. In addition, the viscosities of the 1 wt% chitin solutions in NaOH/urea at 20 °C increased from 5 cP for pure chitin to 18.8 cP for IL-chitin, while for cellulose solutions the viscosities of the 1 wt% solutions increased from 8.9 cP for MCC to 31 cP for CRM-175. Both of these observations suggest that the biopolymers obtained by IL extraction from raw biomass have higher MWs than the commercial biopolymers. A reconstitution process of PG-chitin with IL (Rec. PG) was performed to remove insoluble impurities. It was found that although the purity of Rec. PG was improved and the errors in viscosity were decreased, the viscosity of the Rec. PG solution, 8.4 cP, was substantially lower than that of PG-chitin (14.1 cP), suggesting that depolymerization might occur during IL dissolution. To determine the biopolymer concentrations that can be dissolved in a corresponding NaOH/urea system, the concentration above which the solution solidified within 2 min after cooling to room temperature was determined, i.e., the maximum concentration. Solutions of each biopolymer in the optimum NaOH/urea at 1 wt% were prepared using F/T treatment and then the biopolymer concentration in the solution was increased step-wise in 0.1 wt% increments, followed by several F/T cycles. The additions were continued until immediate gelation occurred at room temperature. It should be noted here that increasing the biopolymer concentration above the maximum concentration led to gelation of the solution, however, the resulting hydrogel was too weak to transfer from the mold (vial) and we considered these to be weak “pseudohydrogels.” Among all the biopolymers tested, CRM-175 and IL-chitin demonstrated the lowest maximum concentrations of 1.4 and 2.4 wt%, respectively (Table 1), again a likely indicator of a higher


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MW. The maximum concentrations for MCC (5.2 wt%), CRM-POM (5.4 wt%), PG chitin (4.3 wt%), and Rec. PG (6.5 wt%) were all similar and low. The maximum concentration found for pure chitin was much higher (12.5 wt%), again suggesting the much lower MW of this polymer, which results from its multistep processing. Interestingly, the maximum concentrations observed were independent of the particle size (not shown in the tables). However, smaller particles did result in faster dissolution, i.e., fewer F/T cycles. For example, 1 F/T cycle was needed for IL-chitin dissolution when the particle size was ˂ 125 µm vs. 8 F/T cycles when the particle size was ˃ 250 µm. This can most likely be attributed to the higher surface area and easier access of the smaller particles, although we cannot rule out the possibility of a reduction of MW due to mechanical degradation during grinding. Formation of Hydrogels. Although coagulation in acid, salt, or organic anti-solvent is frequently employed for hydrogel formation from biopolymer/NaOH/urea solutions, 43 when attempting to form bulk hydrogels or hydrogels from solutions of low concentrations,44 thermally induced gelation is considered to be more proper. Therefore, to produce stable bulk hydrogels, two strategies were attempted here: a) curing, i.e., keeping the solution at a certain temperature for a given amount of time to afford firm physical cross-linking, and b) adding a chemical crosslinker, epichlorohydrin (ECH), to form chemical hydrogels. The flowchart for each process is shown in Fig. 1.


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Fig. 1. Processes for making physical and chemical hydrogels with IL-chitin (left) and CRM-175 (right) as examples. Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. Physical Hydrogels. The effect of curing temperature on the time required to form firm physical hydrogels from solutions at maximum concentrations was studied (Table S2). At 30 °C, longer times (usually > 12 h) were required to form firm hydrogels from either cellulose or


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chitin. Curing time significantly decreased at higher temperatures, i.e., 4 h at 60 °C for chitin or 50 °C for cellulose. We therefore selected heating regimes of 50 °C for cellulose and 60 °C for chitin, attempting to produce strong physical hydrogels in a shorter time (4 h). The effect of biopolymer concentration on hydrogel formation was also studied to determine the minimum biopolymer concentrations required to prepare physical hydrogels (Table 2). Only those biopolymers with higher apparent MWs, i.e., IL-chitin, PG-chitin, and CRM-175, were able to produce stable physical hydrogels from solutions of concentrations at and above 2 wt% (for chitin) or 1 wt% (for CRM-175). The remaining biopolymers studied, i.e. pure chitin, Rec. PG, MCC, and CRM-POM, could not form physical gels at lower concentrations or only formed pseudo gels at higher concentrations, regardless of the higher purity. For the comparative data below, we have designated the physical hydrogels of 2 wt% from PG- and IL-chitin as PG-2-0 and IL-2-0, respectively, and the cellulosic physical gel as Ph-CRM-175.

Table 2. Effect of Biopolymer Concentration on the Formation of Physical Hydrogels at 50 °C (for Cellulose) or 60 °C (for Chitin) for 4 h. NaOH/urea (x wt%/y wt%)

1.0 wt%

1.5 wt%

2.0 wt%

3.0 wt%

4.0f wt%

Pure chitin PG-chitin Rec. PGa IL-chitin MCC CRMPOM

8/4 8/4 8/4 8/4 6/4

—b — — — —

— Pseudoc — Pseudo —

— √, PG-2-0d Pseudo √, IL-2-0d —

— √ Pseudo N/Ae Pseudo

— √ Pseudo N/A Pseudo

6/4

—

—

—

Pseudo

Pseudo

CRM-175

4/4

Chitin

Biopolymer

Cellulose

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√, PhN/A N/A N/A N/A CRM-175 a b Rec. PG: Reconstituted PG-chitin from [C2mim][OAc] to remove impurities; No hydrogel was formed; c Pseudo: The resulting gel was not strong enough to maintain integrity during transfer; d Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin; e N/A: limited by the maximum concentration of the biopolymer; f The results were the same even at maximum biopolymer concentrations.


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Freshly-prepared chitin physical hydrogels appeared yellowish and non-transparent. The opacity is primarily caused by the architecture of the macromolecules or the hydrogel formation mechanism (phase separation vs. crystallite formation), which was previously reported as the key factor for the transparency of the hydrogel; when phase separation precedes crystallite formation (molecular association), an opaque gel is obtained.20 Further, the high gelation temperature (50 or 60 °C) increased the degree of phase separation and thereby heterogeneities (i.e., the number and size of polymer aggregates),45,46 resulting in the complete opacity of chitin hydrogels. After washing with DI water, the chitin hydrogels appeared ivory-colored possibly due to the removal of NaOH and urea molecules. Ph-CRM-175 had a yellow color before and after washing, probably caused by the presence of lignin. The physical chitin hydrogels appeared swollen after washing, whereas Ph-CRM-175 hardly swelled, suggesting over-entanglement with much stronger chain intertwining and hydrogen-boding developing during physical cross-linking of the cellulosic biopolymer chains. Chemical Hydrogels. The generalized mechanism of cross-linking of the biopolymers with ECH is shown in Scheme 1.47 Lower heating time was needed to form chemical hydrogels (1 h for cellulose, and 2 h for chitin) than that needed for physical hydrogels (4 h) at the same temperature, indicating the effectiveness of ECH in cross-linking. The real mass of ECH consumed for cross-linking was estimated taking into account the MW increase due to the incorporation of ‒CH2CH(OH)CH2‒ connecting units, and under the assumption that the mass of biopolymers in the chemical gel from the biopolymer solution (1 g) was equal to that in the physical gel of the same biopolymer concentration (see ESI for calculation). The consumption of ECH for gel formation was found to be low, i.e., 0.0033 g for 2 wt% chitin gels and 0.0015 g for


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1 wt% CRM-175 gels (Table 3), with others unable to be calculated due to the failure of physical gel formation.

Scheme 1. Chemical Cross-linking Reaction between ECH and Biopolymer (chitin is shown). Reproduced from Ref. 47 with permission from the Royal Society of Chemistry. Table 3. Preparation of Chemical Hydrogels at 50 °C/1 h for Cellulose and 60 °C/2 h for Chitin. Conc. (wt%)

Mass ratio of ECH/biopolymerf

Gel Codeg

MCC CRM-POM

1 2a 1 2 2c 1 2 4d 4

10 10 10 5 5 10 5 2.5 2.5

— — √, PG-1-10 √, PG-2-5 Too weak √, IL-1-10 √, IL-2-5 √, MCC √, CRM-POM

ECH consumptionh (g) N/A N/A N/A 0.0033(2) N/A N/A 0.0033(2) N/A N/A

CRM-175

1e

10

√, CRM-175

0.0015(2)

Biopolymer

Chitin

Pure chitin PG-chitin Rec. PGb IL-chitin Cellulose

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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a

Even increasing to maximum concentration, no firm gels from pure chitin were formed; b Rec. PG: Reconstituted PG-chitin from [C2mim][OAc] to remove impurities; c This concentration was used attempting to produce gels for comparison with PG-2-5; d Lower concentration led to very weak hydrogels; e The concentration is limited by the maximum concentration in NaOH/urea; f Mass ratio of ECH/biopolymer: e.g., 10 represents the mass of ECH added is 10 times that of the biopolymer in the solution; g Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin; h The real mass of ECH consumed for cross-linking.


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The influence of biopolymer concentration on chemical gel formation was tested, and it was found that chemical hydrogels were formed from the three biopolymer solutions at concentrations that successfully formed physical hydrogels (2 wt% PG- and IL-chitin, and 1 wt% CRM-175). Additionally, 1 wt% chitin (PG- and IL-chitin) solutions and 4 wt% CRM-POM and MCC solutions that could not form firm physical gels did produce chemical hydrogels, whereas pure chitin of the lowest MW was not able to form either type of hydrogel. After washing with DI water, swelling was observed in both chitin and cellulose hydrogels. The chitin chemical hydrogels appeared similar (nontransparent) to the physical ones except for the much higher swollen volumes after washing (Table S3, ESI). Although the preparation conditions were similar, cellulosic hydrogels exhibited higher transparency than chitin ones, indicating a lower degree of phase separation. In particular, MCC hydrogels had the highest transmittance among all hydrogels before and after washing, while CRM hydrogels of lower purity became translucent after dramatic swelling during washing. The transparency of the CRM hydrogels increased after swelling compared with the as-prepared gels, which may suggest that transparency of cellulosic hydrogels is also related to the degree of swelling. As expected, the firmness of the 1 wt% swollen chitin gels were lower compared with analogous 2 wt% gels, while the highly absorbent cellulosic gels were so soft and sticky that extra care had to be taken when transferring these from washing bath to the ethanol container (for the following solvent exchange and supercritical drying). Structural confirmation of chemical cross-linking was assessed through Fourier transform infrared (FTIR) spectroscopy (taking chitin as a benchmark, Fig. S3, ESI). A series of narrow absorption bands, typical for chitin, including O-H stretch (3445 cm-1), different types of C-H stretches (3099, 2918, 2870 cm-1), amide I (C=O stretch at 1653 and 1620 cm-1 due to the


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occurrence of hydrogen bonding C=O···HN and C=O···HOCH2, respectively48), amide II (N–H bend and C-N stretch at 1550 cm-1), amide III (complex vibrations of NHCO at 1305 cm-1), 49 and C–O–C and C–O stretches (four intensive bands at 1153, 1110, 1065 and 1022 cm-1) were detected. While physical gels exhibit the typical FTIR signature of chitin, the FTIR spectra of chemically cross-linked gels looks slightly different. If we compare, for example, the band corresponding to the stretch of the C–O bond in chemical (1028 cm-1) and physical (1022 cm-1) gels, the latter looks broader and more pronounced, an indication that there are more OH groups bonded to the carbon backbone. There is also a noticeable difference in the amide I and II bands. In native chitin, the amide I band at 1620 cm-1 is higher in intensity than the one at 1653 cm-1, and the amide II band is higher than the amide I bands. In chemically cross-linked chitin, the relative intensities of the two amide I bands are reversed and amide II becomes lower than amide I, suggesting that a) chemical cross-linking results in shortening the distances between biopolymeric molecules thus C=O and NH become involved in the hydrogen bond interactions to a larger extent; b) chemical cross-linking may occur at the CH2OH sites. Due to the very low amount of cross-linking, other changes were not detected in the FTIR spectra. Preparation of Aerogels. Physical and chemical hydrogels were oven-dried, freeze-dried, or supercritically CO2 dried (ScCO2-dried). Oven-drying totally destroyed the structure of the hydrogel network, leading to collapse of the pores due to the recrystallization of the biopolymers (Fig. S4). Additionally, powder X-ray diffraction (PXRD) analysis showed that the (020) crystalline plane of α-chitin at 9.2° or the ( 110 ) plane of cellulose II at 11.8° shifted (partially) to 7.6° after chemical cross-linking (Fig. S5). (While these observations suggest a change in crystalline form induced by the chemical cross-linking, additional characterization techniques are


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needed to completely understand these changes.) Although freeze-dried gels didn’t collapse, drying gels using this process required several days. ScCO2-drying was employed to provide porous aerogels. The hydrogels were first immersed in ethanol forming alcogels, which were much more compact and firmer than the original hydrogels. Chitin and CRM-175 alcogels appeared glassy and very regular, whereas MCC and CRM-POM alcogels were rough and lumpy (Fig. 2). After ScCO2-drying, all of the resulting aerogels were lightweight, and the PXRD analysis showed that the crystalline peak at 19.2° (for 110 plane of α-chitin) or 20.0° (for 110 plane of cellulose II) almost disappeared and an amorphous peak at about 11.8° occurred (Fig. S5). The amorphous scattering for the aerogels is different from that of amorphous cellulose or chitin (ca. 16°,50,51) which may be an important clue for understanding the molecular mechanisms of aerogel structure. Most of the aerogels appeared white or nearly white except for the CRM-POM aerogel that exhibited a yellow color. Additionally, cellulosic aerogels were much harder and stronger (by pinch testing) than the chitin aerogels, which might be associated with both the nature of the biopolymer and the more compact aerogel microstructure.

Fig. 2. Photographs of (a) IL-chitin and (b) cellulose gels (original gels ‒ left, alcogels ‒ middle,


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and aerogels ‒ right) during preparation. Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. Morphology of Aerogels. The surfaces and cross-sections of both the chitin and cellulosic aerogels were studied using scanning electron microscopy (SEM, Fig. 3). As the cellulosic aerogels were much more compact, higher magnification was required. Each aerogel had a characteristic channel-like structure of phase separation (so-called spinodal decomposition 52 ) resulting from the thermodynamic instability of the biopolymer/NaOH/urea solutions. The chitin aerogels consisted of a discontinuous hierarchical porous network, which entailed a welldeveloped macro-porous system with pore diameters up to 40 µm and a nanoscale substructure. Contrarily, the cellulosic aerogels were much more compact and homogeneous, consistent with the lower phase separation and higher transparency. Most of the pores in the cellulosic aerogels were unobservable and the maximum pore size was less than 1 µm. The CRM aerogels also included a few uneven fibrils, possibly due to the presence of lignin or hemicellulose impurities.

Fig. 3. SEM images of surfaces and cross-sections of IL-chitin (IL-2-5, top) and cellulose (CRM175, bottom) aerogels at magnifications in the range of 500‒2500, and 2000‒4000, respectively.


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Shrinkage and Porosity. Air-drying led to collapse of the gels with up to 98% shrinkage compared with the original hydrogels, while ScCO2-drying maintained the pore structure and provided much lower shrinkage. During both solvent exchange and ScCO2-drying, the volumes of the physical gels hardly changed, whereas all chemical hydrogels suffered relatively severe volume reduction (Table 4). Such differences in shrinkage could be explained by the proposed mechanism shown in Fig. 4. Specifically, in chemical hydrogels, the polymer chains are likely to be less intertwined than those in physical gels due to the presence of the cross-linker and the shorter time required for preparation. When immersed in ethanol and then ScCO2-dried, the stretched biopolymer chains in chemical hydrogels can be re-entangled, resulting in significant shrinking. Less entanglement might also be the reason for the higher shrinkage of the chemical chitin hydrogels at 1 wt% concentration than those at 2 wt%. In other words, the bigger pores in the more swollen original hydrogels, even with thinner walls in the 1 wt% gels, are assumed to be easier to collapse during solvent exchange and ScCO2-drying. Table 4. Shrinkage, Density, and Porosity of Chitinous and Cellulosic Gels. Density Porosity (%) (×10-2 g/cm3) PG-2-0 -0.3c -1.7c 2.6(1) 96(1) IL-2-0 -0.5c -20(3) 2.6(2) 95(1) PG-1-10 -34(3) -57(2) 2.4(1) 95(1) PG-2-5 -9.0(3) -38(2) 3.3(1) 94(1) IL-1-10 -37(3) -65(1) 2.7(1) 94(1) IL-2-5 -24(3) -48(2) 3.4(1) 94(1) MCC -76(2) -88(3) 18(3) 62(3) CRM-POM -71(3) -84(3) 7.7(5) 87(2) CRM-175 -58(2) -87(2) 5.8(4) 91(2) Ph-CRM-175 0 0 5.9(1) 90(2) a Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. b Salco and Saero: Volume shrinkage of alcogels and aerogels compared with the original gels, respectively; c The deviation is higher than the average value. Salcob (%)

Saerob (%)

Chitin

Gel Codea

Cellulose

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Fig. 4. The proposed structural change of chemical and physical gels before and after shrinking caused by solvent exchange and ScCO2-drying. The type of biopolymer also had an effect on shrinkage. Chemical hydrogels of cellulose sharply shrank after drying, forming both denser and harder aerogels than chitin hydrogels. The MW of biopolymers seemed to have little impact on the total shrinking of the gels, which can be deduced from the CRM-175 and CRM-POM chemical gels (Table 4). Additionally, the purity of the biopolymers might play a role. PG-2-5 (made from PG-chitin as received) showed less shrinkage than purer IL-2-5 (made from IL-chitin isolated from SS with [C2mim][OAc]), although we cannot completely eliminate the influence of the lower MW of PG-chitin. All of the chitin aerogels in this work had densities an order of magnitude lower (2.4‒3.4 ×10-2 g/cm3) than those reported in the literature (12‒27 ×10-2 g/cm3),45,53 which may be related to the different solvent systems, chitin concentrations, or gelation methods (heating or coagulation) used. Moreover, the densities we obtained approached those of chitin nanowhisker-based aerogels (0.5‒2.1 ×10-2 g/cm3).54-56 These low densities correlate well with the observed high porosities (94‒96%).


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The cellulosic aerogels were denser than the chitin gels, in agreement with their appearances in the SEM images. Their density and porosity were roughly in agreement with the literature data.57,58 Of all the cellulosic aerogels, CRM-175 gels (physical and chemical) had the lowest densities and thus the highest porosities, while the MCC chemical gel possessed the highest density and thus the lowest porosity. Rehydration Properties. The water rehydration ability of the hydrogels was tested by placing the aerogels in water for 24 h to evaluate both the rehydration ratio and rate. Immediately after placing the hydrogels into DI water, all aerogels would first shrink before swelling due to the gas venting from the aerogels. Rehydrated cellulosic hydrogels were quite tough, whereas the chitin gels were relatively fragile, which may affect their service life and recycling use. The presence of lignin did not inhibit the water uptake of CRM hydrogels. Out of all gels, the CRM-175 chemical hydrogel showed the highest water uptake (9300% or 93 times the weight of the aerogel), followed by the higher MW chitin gels IL-1-10 (6800%), IL-2-5 (4900%), and finally IL-2-0 (4350%; Fig. 5). These results suggest that higher biopolymer MW, lower biopolymer concentration, and chemical gel type are favorable parameters in achieving higher ultimate aerogel water uptake capacity. These three conditions favor the extension of the polymer chains and thus the formation of bigger pores in the rehydrated hydrogels, which might explain the higher rehydration ratios.


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Fig. 5. Rehydration ratio of chitin and cellulosic hydrogels after soaking the aerogels in water at room temperature for 24 h. Chitin gel code XX-m-n, where XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. The rehydration rate was evaluated by continuously monitoring hydrogel mass increase during water uptake (Fig. 6a). The rehydration rate depended on the biopolymer concentration, i.e., 1 wt% chitin hydrogels rehydrated faster than the 2 wt% gels, presumably due to faster water diffusion in the bigger pores of the former. The pore structure of the aerogels also affected the rate of water uptake, with cellulose gels of denser structure absorbing water much slower than chitin gels because their microporous network is harder to unfold. MCC and CRM-POM chemical hydrogels absorbed water particularly slowly since both were the most compact aerogel structures and contained the highest biopolymer concentrations. Although the aerogels could be well rehydrated, the volumes and masses of the fully rehydrated hydrogels were lower than observed for the original gels. Mass recoveries (calculated using Eq. 7, ESI) for the chitin hydrogels depended on chitin concentration, with higher concentrations resulting in higher recoveries (Fig. 6a), which was likely to be related to the lower


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shrinkage (collapse) of these hydrogels during drying (Table 4, Saero column). For this same reason, the chemical cellulose hydrogels with higher shrinkage showed much lower recoveries than almost all chitin hydrogels. PG-chitin hydrogels recovered less than the IL-chitin ones likely due to the presence of impurities and the lower MW. Dye loading and release. To test the concept of pharmaceutical loading and release, all cellulose and chitin aerogels were loaded with indigo carmine, by placing the aerogels in 1 mg/mL dye aqueous solutions. Dye molecules successfully diffused into the swollen hydrogels as demonstrated by the dark blue color of the loaded gels (Fig. 6b). Dye loading in the hydrogel was calculated by determining the dye concentration of the aqueous solution before and after loading. While we expected total dye loading in the hydrogels to be related to the rehydration ratio, this was not the case: IL-2-5 showed higher dye loading (45 mg/g) than CRM-175 (29 mg/g) and IL-1-10 (28 mg/g), opposite to the rehydration order CRM-175 > IL-1-10 > IL-2-5. This may result in the conclusion that during dye loading, smaller pores (higher concentration and lower water absorbency) of the hydrogels might lead to a stronger interaction of the biopolymer with the dye, thus increasing the loading capacity of the hydrogels.


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Fig. 6. (a) Mass recoveries of hydrogels as a function of time during rehydration; (b) cumulative dye release from loaded hydrogels in phosphate buffer saline (PBS) over time (the insets are the loaded hydrogels). Chitin gel code XX-m-n, where XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. The dye release was studied by immersing the loaded gels into phosphate buffer saline (PBS) solution. In all cases, a typical diffusion-controlled release profile 59 was observed from the cumulative release plot (Fig. 6b), which includes a rapid burst release of the dye near the surface, followed by a slower release of the dye located within the hydrogel network. The burst release led to dye losses of over 80% in 1 wt% hydrogels, but as low as 30‒40% in hydrogels of higher concentrations, indicating that the release rate was primarily controlled by the biopolymer concentration, where lower concentrations led to higher rehydration and thus bigger pores exhibiting faster release. The 1 wt% hydrogels exhibited steep dye release within 3 h, while 2 or 4 wt% hydrogels released the dye much more slowly within 24 h. These release times roughly matched with the reported duration time of drugs permeated in hydrogel systems. 60 The dye loading also


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influenced the release rate, PG-2-0 (with a smaller dye loading of 25 mg/g) released slightly faster than other 2 wt% chitin gels. In addition, though at the same concentration and with similar dye loading, CRM-POM gel had higher release rate than MCC gel, likely due to the different interactions between various biopolymers (cellulose, hemicellulose, and lignin) and the dye molecules. Additionally, the aerogels were immersed in dye/ethanol solution followed by one more ScCO2-drying step to determine the dye molecule structure in the gels. Crystal form of the dye molecules was not detected in the gel (PXRD spectra shown in Fig. S6), and thus precipitation of the dye into the pores of the gels was ruled-out. CONCLUSIONS By taking advantage of the ability of ionic liquids to extract chitin or CRM of higher MWs directly from the corresponding biomass, we were able to prepare “physical” (no cross-linker) and/or “chemical” (using a cross-linker) hydrogels. The IL-extracted biopolymers of apparent higher MWs (CRM-175, IL-chitin) and PG-chitin could form both stable physical and chemical hydrogels, whereas lower MW polymers (Rec. PG, CRM-POM, and MCC) required a covalent cross-linker for hydrogel formation. Low MW pure chitin was not suitable for the preparation of hydrogels of either type. The hydrogels were supercritically dried producing amorphous, lightweight, and porous aerogels. Both chitin and cellulose aerogels showed high water absorbency, especially those prepared from CRM-175 and IL-1-10 and chemically cross-linked, again most likely due to the apparent higher MWs and lower chain entanglement (due to the incorporation of the cross-linker and the low biopolymer concentration). The results obtained from dye loading and release suggest the potential use of both chitin and cellulose hydrogels to effectively load and release


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active species. The gels with higher biopolymer concentration resulting in smaller pores exhibited increased dye loading capacity. As with water uptake, the release rate was mostly affected by the biopolymer concentration of the hydrogel, while the amount of dye and type of biopolymer had less effect. We believe that the use of chitin and cellulose hydrogels for drug delivery applications has great promise, but further research is needed. For example, the biocompatibility of CRM gels (the effect of lignin content), their mechanical properties, and lowering the burst release (through surface extracting or coating) should be investigated. Considering the unique high strength of cellulosic aerogels, we think that these materials may also find use in other fields, such as thermal insulation and tissue engineering. ASSOCIATED CONTENT Supporting Information Isolation of biopolymers with [C2mim][OAc], determination of purity of the biopolymers, measurement of the viscosity of biopolymer/NaOH/urea solutions, supercritical CO2 drying, calculation of ECH consumption, characterization methods (PXRD and SEM), and volumes and masses of all gels. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author *(R.D.R.) E-mail: [email protected] Notes


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Dr. Robin D. Rogers has partial ownership of 525 Solutions. Drs Gabriela Gurau and Julia Shamshina are part-time employees of 525 Solutions. The University of Alabama and McGill University maintain approved Conflict of Interest Management Plans. Dr. Wang’s current affiliation: Institute of Process Engineering, Chinese Academy of Sciences, No. 1 Beierjie Zhongguancun Haidian District, Beijing 100190, China. ACKNOWLEDGMENT The authors would like to thank 525 Solutions, Inc., the DOE SBIR Office of Science (DESC0010152) and the China Scholarship Council (No. 201306600007) for financial support. ABBREVIATIONS [C2mim][OAc], 1-ethyl-3-methylimidazolium acetate; CRM-175, cellulose-rich material isolated using IL at 175 °C; CRM-POM, cellulose-rich material isolated using IL with polyoxometalate (POM); ECH, epichlorohydrin; F/T, freeze/thaw; IL, ionic liquid; IL-chitin, chitin isolated from shrimp shells (SS) with IL; MCC, microcrystalline cellulose; MW, molecular weight; PG-chitin, practical-grade chitin; Rec. PG, reconstituted PG-chitin with IL; ScCO2-drying, supercritical CO2 drying.


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(52) Higgins, A. M.; Jones, R. A. L. Anisotropic spinodal dewetting as a route to selfassembly of patterned surfaces. Nature 2000, 404, 476-478. (53) Tsioptsias, C.; Michailof, C.; Stauropoulos, G.; Panayiotou, C. Chitin and carbon aerogels from chitin alcogels. Carbohydr. Polym. 2009, 76, 535-540. (54) Lu, Y.; Sun, Q.; She, X.; Xia, Y.; Liu, Y.; Li, J.; Yang, D. Fabrication and characterization of α-chitin nanofibers and highly transparent chitin films by pulsed ultrasonication. Carbohydr. Polym. 2013, 98, 1497-1504. (55) Tsutsumi, Y.; Koga, H.; Qi, Z. D.; Saito, T.; Isogai, A. Nanofibrillar chitin aerogels as renewable base catalysts. Biomacromolecules 2014, 15, 4314-4319. (56) Zhou, Y.; Fu, S.; Pu, Y.; Pan, S.; Ragauskas, A. J. Preparation of aligned porous chitin nanowhisker foams by directional freeze-casting technique. Carbohydr. Polym. 2014, 112, 277283. (57) Gavillon, R.; Budtova, T. Aerocellulose: New highly porous cellulose prepared from cellulose-NaOH aqueous solutions. Biomacromolecules 2008, 9, 269-277. (58) Cai, J.; Kimura, S.; Wada, M.; Kuga, S.; Zhang, L. Cellulose aerogels from aqueous alkali hydroxide-urea solution. ChemSusChem 2008, 1, 149-154. (59) Huang, X.; Brazel, C. S. On the importance and mechanisms of burst release in matrixcontrolled drug delivery systems. J. Control Release 2001, 73, 121-136. (60) Bhattarai, N.; Gunn, J.; Zhang, M. Q. Chitosan-based hydrogels for controlled, localized drug delivery. Adv. Drug Deliver. Rev. 2010, 62, 83-99.


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Comparison of Hydrogels Prepared with Ionic Liquid-Isolated vs. Commercial Chitin and Cellulose Xiaoping Shen,†,‡ Julia L. Shamshina,§ Paula Berton,†,║ Jenny Bandomir,† Hui Wang,† Gabriela Gurau§,║ Robin D. Rogers*,†,║

Synopsis Chitin and cellulose extracted from biomass using ionic liquid require lower concentrations than commercially available biopolymers to prepare physical and chemical hydrogels. TOC


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Comparison of Hydrogels Prepared with Ionic Liquid-Isolated vs. Commercial Chitin and Cellulose Xiaoping Shen,†,‡ Julia L. Shamshina,§ Paula Berton,†,║ Jenny Bandomir,† Hui Wang,† Gabriela Gurau§,║ Robin D. Rogers*,†,║ †

Department of Chemistry, The University of Alabama, Tuscaloosa, AL 35487, USA

‡

Key Laboratory of Bio-based Material Science and Technology (Ministry of Education),

Northeast Forestry University, 26 Hexing Road, Harbin 150040, China §

525 Solutions, Inc., 720 2nd Street, Tuscaloosa, AL 35401, USA

║

Department of Chemistry, McGill University, 801 Sherbrooke St. West, Montreal, QC H3A

0B8, Canada



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ABSTRACT

Physical and/or covalently-linked (chemical) hydrogels were prepared from chitin and cellulose extracted with ionic liquid from shrimp shells and wood biomass, respectively, and compared with hydrogels prepared from commercially available biopolymers, practical grade chitin, and microcrystalline cellulose. The highly porous aerogels were formed by initial dissolution of the biopolymers in NaOH/urea aqueous systems using freeze/thaw cycles, followed by thermal treatment (with or without epichlorohydrin as a cross-linker) and supercritical CO2 drying. The ionic liquid-extracted cellulose pulp and chitin, as well as practical grade chitin could form both stable physical and chemical hydrogels, whereas biopolymers of lower apparent molecular weight such as microcrystalline cellulose required a covalent cross-linker for hydrogel formation and commercially available pure chitin was not suitable for the preparation of hydrogels of either type. Hydrogels prepared from the ionic liquid-extracted biopolymers exhibited properties substantially different from those made from the commercially available biopolymers. Loading of an active ingredient into the hydrogel and its subsequent release was demonstrated using indigo carmine and revealed that the release rate was controlled mainly by the biopolymer concentration of the gel network.

KEYWORDS: Hydrogel, aerogel, chitin, cellulose, ionic liquid, release


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INTRODUCTION Hydrogels are highly swollen, hydrophilic, three-dimensional (3D) polymeric networks capable of absorbing large amounts of water or biological fluids and swelling.1 Currently, the majority of the hydrogels are manufactured using (meth)acrylic acid or acrylamide, compounds which raise public health concerns. 2 , 3 Considering not only safety, but also biocompatibility, tissuemimicking consistency, and biodegradability, biopolymer-based hydrogels have been drawing increasing interest and attention. 4 Cellulose and chitin, the two most abundant renewable biopolymers on earth, consist of straight chain of β-(1→4)-linked D-glucose, and β-(1→4)linked N-acetyl-D-glucosamine, respectively. Because of the hydroxyl and/or acetamido groups present in their structures, fused or amorphous cellulose and chitin chains are highly hydrophilic and potentially suitable to form superabsorbent hydrogels.5,6 In addition to their biodegradability,7,8 swollen hydrogels from cellulose and chitin are elastic but soft and possess diffusive properties identical to those of a liquid, and thus can absorb and release water or biological fluids in a reversible non-irritating manner, making them compatible with living tissues. 9, 10 Consequently, these hydrogels have attracted tremendous attention as excellent candidates for biomedical applications, such as drug delivery,11 wound dressings,12 and tissue engineering matrices.13 Chitin and cellulose hydrogels are usually prepared through a two-step process involving biopolymer dissolution and chain cross-linking. To date, there are only a few specific solvent systems reported to dissolve chitin or cellulose and form hydrogels, such as ionic liquids (ILs), 14 , 15 alkali or alkali/urea (thiourea) aqueous systems, 16 - 18 and polar solvent systems including LiCl/N-methyl-2-pyrolidone (NMP),8, LiCl/dimethyl sulfoxide (DMSO),

21

19

LiCl/N,N-dimethylacetamide (DMAc),

20

and saturated CaCl2•2H2O/methanol. 22 Among these


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solvents, alkali/urea aqueous systems are attractive due to their low cost, energy consumption, and toxicity, and the relatively low viscosity of the resulting solutions. After dissolution, physical or chemical cross-links are needed to provide structural integrity and thus form hydrogels. Physical cross-links involve various interactions such as chain entanglements, strong van der Waals forces, hydrogen bonds, crystallite associations,23 and/or ionic interactions. 24 Chemical cross-links are covalent bonds between polymeric chains themselves or between the polymer(s) and a cross-linker added to gelate the solution.25 Physical hydrogels can be obtained from solution either by curing at a certain temperature,16,18 or by coagulating with an anti-solvent in a specific fashion (e.g., molding, casting, or dropping).19,26 Chemical hydrogels can be obtained via cross-linking using irradiation (e.g., electron beam,27 γray, 28 or UV 29 ) or using a chemical cross-linker (e.g., epichlorohydrin,9, 30 , 31 1,2,3,4butanetetracarboxylic dianhydride,8,32 or succinic anhydride7,33). In our previous work, we have developed a series of methods for the isolation of cellulose-rich materials (CRMs)34-36 and chitin (IL-chitin)37,38 by direct dissolution and reconstitution of the biopolymers from their corresponding biomass using ILs. It was found that these IL-isolated biopolymers had particularly higher molecular weights (MWs) than most commercial sources. We were curious whether the unique forms of the biopolymers we extracted from biomass using ILs would provide hydrogels with properties that might differ from the currently available materials. To test this, we utilized these extracted biopolymers for the formation of both physical and chemical hydrogels in NaOH/urea as the solvent system and compared their properties to analogous gels prepared from commercial cellulose and chitin and to hydrogels published in the literature. The formation, structure and morphology, water absorbency (rehydration), and active ingredient loading and release of these hydrogels are described here.


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RESULTS AND DISCUSSION Biopolymer Extraction and Dissolution in NaOH/urea. Chitin and cellulosic biopolymers used for this study were respectively extracted from shrimp shells (SS) and poplar, using [C2mim][OAc] and procedures we have previously described (See ESI for complete experimental details and composition of the biopolymers, Table S1).35,36,38 The materials studied here include IL-chitin (isolated from dried/processed SS using microwave-assisted dissolution38), CRM-175 (extracted from poplar at 175 °C for 30 min followed by regeneration with 50% v/v acetone/H2O35), and CRM-POM (extracted from poplar with polyoxometalate (POM) at 110 °C for 2 h followed by regeneration using 50% v/v acetone/H2O36). Commercial microcrystalline cellulose (MCC), pure chitin, and practical grade (PG)-chitin, were purchased from commercial suppliers and used for comparison. The literature has reported that commercial cellulose and chitin biopolymers can be dissolved in aqueous NaOH/urea solvent systems utilizing freeze/thaw (F/T) cycles.16,17,39,40 In NaOH/urea systems, the concentration of NaOH is reported to greatly affect biopolymer dissolution due to the ability of NaOH to break hydrogen bonds, while urea (generally above 4 wt%) is suggested to act as hydrogen bond donor, creating cellulose inclusion complex and preventing reaggregation of the biopolymer chains. Here, we verified that F/T treatment is also a suitable approach for the dissolution of IL-extracted biopolymers in NaOH/urea. To accelerate dissolution, we adopted an ethanol/dry ice bath for freezing instead of a refrigerator as noted in the literature. We first attempted to replicate this dissolution procedure for the IL-extracted biopolymers using 8 wt% NaOH/4 wt% urea and 7 wt% NaOH/12 wt% urea aqueous solutions; systems reported to dissolve chitin and cellulose, respectively. 41,42 We found (Table 1) that different


5


NaOH concentrations and numbers of F/T cycles were required to dissolve the biopolymers. For chitin, a minimum of 7 wt% NaOH in NaOH/urea was required for its dissolution, while for cellulose, only the 4‒8 wt% NaOH concentrations were able to dissolve MCC and CRM-POM, and 4‒6 wt% to dissolve CRM-175. The optimized conditions were chosen as those in which the biopolymer was dissolved using the least number of F/T cycles: 8 wt% NaOH/4 wt% urea for chitin, 6 wt% NaOH/4 wt% urea for MCC and CRM-POM; and 4 wt% NaOH/4 wt% urea for CRM-175. Table 1. Alkali Solvent Selection, Viscosity Values of the Resulting Solutions, and the Maximum Concentration of the Biopolymer in the Optimum Solvent System. NaOH/urea x wt%/y wt% (F/Ta cycles)

Chitin

Biopolymer

Cellulose

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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2/2 4/0

4/4

6/4

7/12

8/4 +(2); √d +(3); √ +(3); √ +(4); √

Purechitin PGchitin Rec. PGe ILchitin

‒c

‒

‒

Swc

+(4)c

‒

‒

‒

Sw

+(8)

‒

‒

‒

Sw

+(6)

‒

‒

‒

Sw

+(8)

MCC

‒

‒

+(6)

+(2); √ +(3); √

+(2)

+(4)

Max. conc.i (wt%)

12/6b

20/0b

Viscosityf (cP)

+(2)

+(2)

5.0g

12.5(3)

+(3)

+(3)

14.1(8)h

4.3(2)

+(3)

+(3)

8.4(2)

6.5(2)

+(4)

+(4)

18.8(2)

2.4(2)

‒

‒

8.9(1)

5.2(2)

CRM‒ ‒ +(8) +(6) +(6) ‒ ‒ 9.4(1) 5.4(2) POM CRM+(4); ‒ ‒ +(10) Sw Sw ‒ ‒ 31.0(4) 1.4(2) 175 √ a F/T: Freeze/thaw; b These conditions for chitin might result in chitosan formation; c ‒: insoluble; +: soluble; Sw: insoluble but can swell; d The optimized solvent for each biopolymer; e Rec. PG: Reconstituted PG-chitin from the [C2mim][OAc] solution to remove impurities (100 °C, 1 h; ESI); f Viscosity of 1 wt% solution in NaOH/urea at 20 °C (Fig. S2); g The error was lower than 0.1; h The large error was caused by insoluble impurities in the PG-chitin solution; i Max. conc.: maximum concentration in the optimized NaOH/urea system.


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It is noteworthy that using any given NaOH/urea system, the number of F/T cycles required depended on the biopolymer, e.g., IL-chitin needed more F/T cycles than pure chitin in the same NaOH/urea solution. In addition, the viscosities of the 1 wt% chitin solutions in NaOH/urea at 20 °C increased from 5 cP for pure chitin to 18.8 cP for IL-chitin, while for cellulose solutions the viscosities of the 1 wt% solutions increased from 8.9 cP for MCC to 31 cP for CRM-175. Both of these observations suggest that the biopolymers obtained by IL extraction from raw biomass have higher MWs than the commercial biopolymers. A reconstitution process of PG-chitin with IL (Rec. PG) was performed to remove insoluble impurities. It was found that although the purity of Rec. PG was improved and the errors in viscosity were decreased, the viscosity of the Rec. PG solution, 8.4 cP, was substantially lower than that of PG-chitin (14.1 cP), suggesting that depolymerization might occur during IL dissolution. To determine the biopolymer concentrations that can be dissolved in a corresponding NaOH/urea system, the concentration above which the solution solidified within 2 min after cooling to room temperature was determined, i.e., the maximum concentration. Solutions of each biopolymer in the optimum NaOH/urea at 1 wt% were prepared using F/T treatment and then the biopolymer concentration in the solution was increased step-wise in 0.1 wt% increments, followed by several F/T cycles. The additions were continued until immediate gelation occurred at room temperature. It should be noted here that increasing the biopolymer concentration above the maximum concentration led to gelation of the solution, however, the resulting hydrogel was too weak to transfer from the mold (vial) and we considered these to be weak “pseudohydrogels.” Among all the biopolymers tested, CRM-175 and IL-chitin demonstrated the lowest maximum concentrations of 1.4 and 2.4 wt%, respectively (Table 1), again a likely indicator of a higher


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MW. The maximum concentrations for MCC (5.2 wt%), CRM-POM (5.4 wt%), PG chitin (4.3 wt%), and Rec. PG (6.5 wt%) were all similar and low. The maximum concentration found for pure chitin was much higher (12.5 wt%), again suggesting the much lower MW of this polymer, which results from its multistep processing. Interestingly, the maximum concentrations observed were independent of the particle size (not shown in the tables). However, smaller particles did result in faster dissolution, i.e., fewer F/T cycles. For example, 1 F/T cycle was needed for IL-chitin dissolution when the particle size was ˂ 125 µm vs. 8 F/T cycles when the particle size was ˃ 250 µm. This can most likely be attributed to the higher surface area and easier access of the smaller particles, although we cannot rule out the possibility of a reduction of MW due to mechanical degradation during grinding. Formation of Hydrogels. Although coagulation in acid, salt, or organic anti-solvent is frequently employed for hydrogel formation from biopolymer/NaOH/urea solutions, 43 when attempting to form bulk hydrogels or hydrogels from solutions of low concentrations,44 thermally induced gelation is considered to be more proper. Therefore, to produce stable bulk hydrogels, two strategies were attempted here: a) curing, i.e., keeping the solution at a certain temperature for a given amount of time to afford firm physical cross-linking, and b) adding a chemical crosslinker, epichlorohydrin (ECH), to form chemical hydrogels. The flowchart for each process is shown in Fig. 1.


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Fig. 1. Processes for making physical and chemical hydrogels with IL-chitin (left) and CRM-175 (right) as examples. Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. Physical Hydrogels. The effect of curing temperature on the time required to form firm physical hydrogels from solutions at maximum concentrations was studied (Table S2). At 30 °C, longer times (usually > 12 h) were required to form firm hydrogels from either cellulose or


9


chitin. Curing time significantly decreased at higher temperatures, i.e., 4 h at 60 °C for chitin or 50 °C for cellulose. We therefore selected heating regimes of 50 °C for cellulose and 60 °C for chitin, attempting to produce strong physical hydrogels in a shorter time (4 h). The effect of biopolymer concentration on hydrogel formation was also studied to determine the minimum biopolymer concentrations required to prepare physical hydrogels (Table 2). Only those biopolymers with higher apparent MWs, i.e., IL-chitin, PG-chitin, and CRM-175, were able to produce stable physical hydrogels from solutions of concentrations at and above 2 wt% (for chitin) or 1 wt% (for CRM-175). The remaining biopolymers studied, i.e. pure chitin, Rec. PG, MCC, and CRM-POM, could not form physical gels at lower concentrations or only formed pseudo gels at higher concentrations, regardless of the higher purity. For the comparative data below, we have designated the physical hydrogels of 2 wt% from PG- and IL-chitin as PG-2-0 and IL-2-0, respectively, and the cellulosic physical gel as Ph-CRM-175.

Table 2. Effect of Biopolymer Concentration on the Formation of Physical Hydrogels at 50 °C (for Cellulose) or 60 °C (for Chitin) for 4 h.

Chitin

Biopolymer

Cellulose

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Pure chitin PG-chitin Rec. PGa IL-chitin MCC CRMPOM

NaOH/urea (x wt%/y wt%)

1.0 wt%

1.5 wt%

2.0 wt%

3.0 wt%

4.0f wt%

8/4 8/4 8/4 8/4 6/4

—b — — — —

— Pseudoc — Pseudo —

— √, PG-2-0d Pseudo √, IL-2-0d —

— √ Pseudo N/Ae Pseudo

— √ Pseudo N/A Pseudo

6/4

—

—

—

Pseudo

Pseudo

√, PhN/A N/A N/A N/A CRM-175 a Rec. PG: Reconstituted PG-chitin from [C2mim][OAc] to remove impurities; b No hydrogel was formed; c Pseudo: The resulting gel was not strong enough to maintain integrity during transfer; d Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin; e N/A: limited by the maximum concentration of the biopolymer; f The results were the same even at maximum biopolymer concentrations. CRM-175

4/4


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Freshly-prepared chitin physical hydrogels appeared yellowish and non-transparent. The opacity is primarily caused by the architecture of the macromolecules or the hydrogel formation mechanism (phase separation vs. crystallite formation), which was previously reported as the key factor for the transparency of the hydrogel; when phase separation precedes crystallite formation (molecular association), an opaque gel is obtained.20 Further, the high gelation temperature (50 or 60 °C) increased the degree of phase separation and thereby heterogeneities (i.e., the number and size of polymer aggregates),45,46 resulting in the complete opacity of chitin hydrogels. After washing with DI water, the chitin hydrogels appeared ivory-colored possibly due to the removal of NaOH and urea molecules. Ph-CRM-175 had a yellow color before and after washing, probably caused by the presence of lignin. The physical chitin hydrogels appeared swollen after washing, whereas Ph-CRM-175 hardly swelled, suggesting over-entanglement with much stronger chain intertwining and hydrogen-boding developing during physical cross-linking of the cellulosic biopolymer chains. Chemical Hydrogels. The generalized mechanism of cross-linking of the biopolymers with ECH is shown in Scheme 1.47 Lower heating time was needed to form chemical hydrogels (1 h for cellulose, and 2 h for chitin) than that needed for physical hydrogels (4 h) at the same temperature, indicating the effectiveness of ECH in cross-linking. The real mass of ECH consumed for cross-linking was estimated taking into account the MW increase due to the incorporation of ‒CH2CH(OH)CH2‒ connecting units, and under the assumption that the mass of biopolymers in the chemical gel from the biopolymer solution (1 g) was equal to that in the physical gel of the same biopolymer concentration (see ESI for calculation). The consumption of ECH for gel formation was found to be low, i.e., 0.0033 g for 2 wt% chitin gels and 0.0015 g for


11


1 wt% CRM-175 gels (Table 3), with others unable to be calculated due to the failure of physical gel formation.

Scheme 1. Chemical Cross-linking Reaction between ECH and Biopolymer (chitin is shown). Reproduced from Ref. 47 with permission from the Royal Society of Chemistry. Table 3. Preparation of Chemical Hydrogels at 50 °C/1 h for Cellulose and 60 °C/2 h for Chitin. Conc. (wt%)

Mass ratio of ECH/biopolymerf

Gel Codeg

MCC CRM-POM

1 2a 1 2 2c 1 2 4d 4

10 10 10 5 5 10 5 2.5 2.5

— — √, PG-1-10 √, PG-2-5 Too weak √, IL-1-10 √, IL-2-5 √, MCC √, CRM-POM

ECH consumptionh (g) N/A N/A N/A 0.0033(2) N/A N/A 0.0033(2) N/A N/A

CRM-175

1e

10

√, CRM-175

0.0015(2)

Biopolymer

Chitin

Pure chitin PG-chitin Rec. PGb IL-chitin

Cellulose

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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a

Even increasing to maximum concentration, no firm gels from pure chitin were formed; b Rec. PG: Reconstituted PG-chitin from [C2mim][OAc] to remove impurities; c This concentration was used attempting to produce gels for comparison with PG-2-5; d Lower concentration led to very weak hydrogels; e The concentration is limited by the maximum concentration in NaOH/urea; f Mass ratio of ECH/biopolymer: e.g., 10 represents the mass of ECH added is 10 times that of the biopolymer in the solution; g Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin; h The real mass of ECH consumed for cross-linking.


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The influence of biopolymer concentration on chemical gel formation was tested, and it was found that chemical hydrogels were formed from the three biopolymer solutions at concentrations that successfully formed physical hydrogels (2 wt% PG- and IL-chitin, and 1 wt% CRM-175). Additionally, 1 wt% chitin (PG- and IL-chitin) solutions and 4 wt% CRM-POM and MCC solutions that could not form firm physical gels did produce chemical hydrogels, whereas pure chitin of the lowest MW was not able to form either type of hydrogel. After washing with DI water, swelling was observed in both chitin and cellulose hydrogels. The chitin chemical hydrogels appeared similar (nontransparent) to the physical ones except for the much higher swollen volumes after washing (Table S3, ESI). Although the preparation conditions were similar, cellulosic hydrogels exhibited higher transparency than chitin ones, indicating a lower degree of phase separation. In particular, MCC hydrogels had the highest transmittance among all hydrogels before and after washing, while CRM hydrogels of lower purity became translucent after dramatic swelling during washing. The transparency of the CRM hydrogels increased after swelling compared with the as-prepared gels, which may suggest that transparency of cellulosic hydrogels is also related to the degree of swelling. As expected, the firmness of the 1 wt% swollen chitin gels were lower compared with analogous 2 wt% gels, while the highly absorbent cellulosic gels were so soft and sticky that extra care had to be taken when transferring these from washing bath to the ethanol container (for the following solvent exchange and supercritical drying). Structural confirmation of chemical cross-linking was assessed through Fourier transform infrared (FTIR) spectroscopy (taking chitin as a benchmark, Fig. S3, ESI). A series of narrow absorption bands, typical for chitin, including O-H stretch (3445 cm-1), different types of C-H stretches (3099, 2918, 2870 cm-1), amide I (C=O stretch at 1653 and 1620 cm-1 due to the


13


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occurrence of hydrogen bonding C=O···HN and C=O···HOCH2, respectively48), amide II (N–H bend and C-N stretch at 1550 cm-1), amide III (complex vibrations of NHCO at 1305 cm-1), 49 and C–O–C and C–O stretches (four intensive bands at 1153, 1110, 1065 and 1022 cm-1) were detected. While physical gels exhibit the typical FTIR signature of chitin, the FTIR spectra of chemically cross-linked gels looks slightly different. If we compare, for example, the band corresponding to the stretch of the C–O bond in chemical (1028 cm-1) and physical (1022 cm-1) gels, the latter looks broader and more pronounced, an indication that there are more OH groups bonded to the carbon backbone. There is also a noticeable difference in the amide I and II bands. In native chitin, the amide I band at 1620 cm-1 is higher in intensity than the one at 1653 cm-1, and the amide II band is higher than the amide I bands. In chemically cross-linked chitin, the relative intensities of the two amide I bands are reversed and amide II becomes lower than amide I, suggesting that a) chemical cross-linking results in shortening the distances between biopolymeric molecules thus C=O and NH become involved in the hydrogen bond interactions to a larger extent; b) chemical cross-linking may occur at the CH2OH sites. Due to the very low amount of cross-linking, other changes were not detected in the FTIR spectra. Preparation of Aerogels. Physical and chemical hydrogels were oven-dried, freeze-dried, or supercritically CO2 dried (ScCO2-dried). Oven-drying totally destroyed the structure of the hydrogel network, leading to collapse of the pores due to the recrystallization of the biopolymers (Fig. S4). Additionally, powder X-ray diffraction (PXRD) analysis showed that the (020) crystalline plane of α-chitin at 9.2° or the ( 110 ) plane of cellulose II at 11.8° shifted (partially) to 7.6° after chemical cross-linking (Fig. S5). (While these observations suggest a change in crystalline form induced by the chemical cross-linking, additional characterization techniques are


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needed to completely understand these changes.) Although freeze-dried gels didn’t collapse, drying gels using this process required several days. ScCO2-drying was employed to provide porous aerogels. The hydrogels were first immersed in ethanol forming alcogels, which were much more compact and firmer than the original hydrogels. Chitin and CRM-175 alcogels appeared glassy and very regular, whereas MCC and CRM-POM alcogels were rough and lumpy (Fig. 2). After ScCO2-drying, all of the resulting aerogels were lightweight, and the PXRD analysis showed that the crystalline peak at 19.2° (for 110 plane of α-chitin) or 20.0° (for 110 plane of cellulose II) almost disappeared and an amorphous peak at about 11.8° occurred (Fig. S5). The amorphous scattering for the aerogels is different from that of amorphous cellulose or chitin (ca. 16°,50,51) which may be an important clue for understanding the molecular mechanisms of aerogel structure. Most of the aerogels appeared white or nearly white except for the CRM-POM aerogel that exhibited a yellow color. Additionally, cellulosic aerogels were much harder and stronger (by pinch testing) than the chitin aerogels, which might be associated with both the nature of the biopolymer and the more compact aerogel microstructure.

Fig. 2. Photographs of (a) IL-chitin and (b) cellulose gels (original gels ‒ left, alcogels ‒ middle,


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and aerogels ‒ right) during preparation. Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. Morphology of Aerogels. The surfaces and cross-sections of both the chitin and cellulosic aerogels were studied using scanning electron microscopy (SEM, Fig. 3). As the cellulosic aerogels were much more compact, higher magnification was required. Each aerogel had a characteristic channel-like structure of phase separation (so-called spinodal decomposition 52 ) resulting from the thermodynamic instability of the biopolymer/NaOH/urea solutions. The chitin aerogels consisted of a discontinuous hierarchical porous network, which entailed a welldeveloped macro-porous system with pore diameters up to 40 µm and a nanoscale substructure. Contrarily, the cellulosic aerogels were much more compact and homogeneous, consistent with the lower phase separation and higher transparency. Most of the pores in the cellulosic aerogels were unobservable and the maximum pore size was less than 1 µm. The CRM aerogels also included a few uneven fibrils, possibly due to the presence of lignin or hemicellulose impurities.

Fig. 3. SEM images of surfaces and cross-sections of IL-chitin (IL-2-5, top) and cellulose (CRM175, bottom) aerogels at magnifications in the range of 500‒2500, and 2000‒4000, respectively.


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Shrinkage and Porosity. Air-drying led to collapse of the gels with up to 98% shrinkage compared with the original hydrogels, while ScCO2-drying maintained the pore structure and provided much lower shrinkage. During both solvent exchange and ScCO2-drying, the volumes of the physical gels hardly changed, whereas all chemical hydrogels suffered relatively severe volume reduction (Table 4). Such differences in shrinkage could be explained by the proposed mechanism shown in Fig. 4. Specifically, in chemical hydrogels, the polymer chains are likely to be less intertwined than those in physical gels due to the presence of the cross-linker and the shorter time required for preparation. When immersed in ethanol and then ScCO2-dried, the stretched biopolymer chains in chemical hydrogels can be re-entangled, resulting in significant shrinking. Less entanglement might also be the reason for the higher shrinkage of the chemical chitin hydrogels at 1 wt% concentration than those at 2 wt%. In other words, the bigger pores in the more swollen original hydrogels, even with thinner walls in the 1 wt% gels, are assumed to be easier to collapse during solvent exchange and ScCO2-drying. Table 4. Shrinkage, Density, and Porosity of Chitinous and Cellulosic Gels. Density Porosity (%) (×10-2 g/cm3) PG-2-0 -0.3c -1.7c 2.6(1) 96(1) c IL-2-0 -0.5 -20(3) 2.6(2) 95(1) PG-1-10 -34(3) -57(2) 2.4(1) 95(1) PG-2-5 -9.0(3) -38(2) 3.3(1) 94(1) IL-1-10 -37(3) -65(1) 2.7(1) 94(1) IL-2-5 -24(3) -48(2) 3.4(1) 94(1) MCC -76(2) -88(3) 18(3) 62(3) CRM-POM -71(3) -84(3) 7.7(5) 87(2) CRM-175 -58(2) -87(2) 5.8(4) 91(2) Ph-CRM-175 0 0 5.9(1) 90(2) a Chitin gel code XX-m-n: XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. b Salco and Saero: Volume shrinkage of alcogels and aerogels compared with the original gels, respectively; c The deviation is higher than the average value. Salcob (%)

Saerob (%)

Chitin

Gel Codea

Cellulose

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Fig. 4. The proposed structural change of chemical and physical gels before and after shrinking caused by solvent exchange and ScCO2-drying. The type of biopolymer also had an effect on shrinkage. Chemical hydrogels of cellulose sharply shrank after drying, forming both denser and harder aerogels than chitin hydrogels. The MW of biopolymers seemed to have little impact on the total shrinking of the gels, which can be deduced from the CRM-175 and CRM-POM chemical gels (Table 4). Additionally, the purity of the biopolymers might play a role. PG-2-5 (made from PG-chitin as received) showed less shrinkage than purer IL-2-5 (made from IL-chitin isolated from SS with [C2mim][OAc]), although we cannot completely eliminate the influence of the lower MW of PG-chitin. All of the chitin aerogels in this work had densities an order of magnitude lower (2.4‒3.4 ×10-2 g/cm3) than those reported in the literature (12‒27 ×10-2 g/cm3),45,53 which may be related to the different solvent systems, chitin concentrations, or gelation methods (heating or coagulation) used. Moreover, the densities we obtained approached those of chitin nanowhisker-based aerogels (0.5‒2.1 ×10-2 g/cm3).54-56 These low densities correlate well with the observed high porosities (94‒96%).


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The cellulosic aerogels were denser than the chitin gels, in agreement with their appearances in the SEM images. Their density and porosity were roughly in agreement with the literature data.57,58 Of all the cellulosic aerogels, CRM-175 gels (physical and chemical) had the lowest densities and thus the highest porosities, while the MCC chemical gel possessed the highest density and thus the lowest porosity. Rehydration Properties. The water rehydration ability of the hydrogels was tested by placing the aerogels in water for 24 h to evaluate both the rehydration ratio and rate. Immediately after placing the hydrogels into DI water, all aerogels would first shrink before swelling due to the gas venting from the aerogels. Rehydrated cellulosic hydrogels were quite tough, whereas the chitin gels were relatively fragile, which may affect their service life and recycling use. The presence of lignin did not inhibit the water uptake of CRM hydrogels. Out of all gels, the CRM-175 chemical hydrogel showed the highest water uptake (9300% or 93 times the weight of the aerogel), followed by the higher MW chitin gels IL-1-10 (6800%), IL-2-5 (4900%), and finally IL-2-0 (4350%; Fig. 5). These results suggest that higher biopolymer MW, lower biopolymer concentration, and chemical gel type are favorable parameters in achieving higher ultimate aerogel water uptake capacity. These three conditions favor the extension of the polymer chains and thus the formation of bigger pores in the rehydrated hydrogels, which might explain the higher rehydration ratios.


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Fig. 5. Rehydration ratio of chitin and cellulosic hydrogels after soaking the aerogels in water at room temperature for 24 h. Chitin gel code XX-m-n, where XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. The rehydration rate was evaluated by continuously monitoring hydrogel mass increase during water uptake (Fig. 6a). The rehydration rate depended on the biopolymer concentration, i.e., 1 wt% chitin hydrogels rehydrated faster than the 2 wt% gels, presumably due to faster water diffusion in the bigger pores of the former. The pore structure of the aerogels also affected the rate of water uptake, with cellulose gels of denser structure absorbing water much slower than chitin gels because their microporous network is harder to unfold. MCC and CRM-POM chemical hydrogels absorbed water particularly slowly since both were the most compact aerogel structures and contained the highest biopolymer concentrations. Although the aerogels could be well rehydrated, the volumes and masses of the fully rehydrated hydrogels were lower than observed for the original gels. Mass recoveries (calculated using Eq. 7, ESI) for the chitin hydrogels depended on chitin concentration, with higher concentrations resulting in higher recoveries (Fig. 6a), which was likely to be related to the lower


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shrinkage (collapse) of these hydrogels during drying (Table 4, Saero column). For this same reason, the chemical cellulose hydrogels with higher shrinkage showed much lower recoveries than almost all chitin hydrogels. PG-chitin hydrogels recovered less than the IL-chitin ones likely due to the presence of impurities and the lower MW. Dye loading and release. To test the concept of pharmaceutical loading and release, all cellulose and chitin aerogels were loaded with indigo carmine, by placing the aerogels in 1 mg/mL dye aqueous solutions. Dye molecules successfully diffused into the swollen hydrogels as demonstrated by the dark blue color of the loaded gels (Fig. 6b). Dye loading in the hydrogel was calculated by determining the dye concentration of the aqueous solution before and after loading. While we expected total dye loading in the hydrogels to be related to the rehydration ratio, this was not the case: IL-2-5 showed higher dye loading (45 mg/g) than CRM-175 (29 mg/g) and IL-1-10 (28 mg/g), opposite to the rehydration order CRM-175 > IL-1-10 > IL-2-5. This may result in the conclusion that during dye loading, smaller pores (higher concentration and lower water absorbency) of the hydrogels might lead to a stronger interaction of the biopolymer with the dye, thus increasing the loading capacity of the hydrogels.


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Fig. 6. (a) Mass recoveries of hydrogels as a function of time during rehydration; (b) cumulative dye release from loaded hydrogels in phosphate buffer saline (PBS) over time (the insets are the loaded hydrogels). Chitin gel code XX-m-n, where XX represents the chitin type, m is the chitin concentration, and n is the mass ratio of ECH to chitin. The dye release was studied by immersing the loaded gels into phosphate buffer saline (PBS) solution. In all cases, a typical diffusion-controlled release profile 59 was observed from the cumulative release plot (Fig. 6b), which includes a rapid burst release of the dye near the surface, followed by a slower release of the dye located within the hydrogel network. The burst release led to dye losses of over 80% in 1 wt% hydrogels, but as low as 30‒40% in hydrogels of higher concentrations, indicating that the release rate was primarily controlled by the biopolymer concentration, where lower concentrations led to higher rehydration and thus bigger pores exhibiting faster release. The 1 wt% hydrogels exhibited steep dye release within 3 h, while 2 or 4 wt% hydrogels released the dye much more slowly within 24 h. These release times roughly matched with the reported duration time of drugs permeated in hydrogel systems. 60 The dye loading also


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influenced the release rate, PG-2-0 (with a smaller dye loading of 25 mg/g) released slightly faster than other 2 wt% chitin gels. In addition, though at the same concentration and with similar dye loading, CRM-POM gel had higher release rate than MCC gel, likely due to the different interactions between various biopolymers (cellulose, hemicellulose, and lignin) and the dye molecules. Additionally, the aerogels were immersed in dye/ethanol solution followed by one more ScCO2-drying step to determine the dye molecule structure in the gels. Crystal form of the dye molecules was not detected in the gel (PXRD spectra shown in Fig. S6), and thus precipitation of the dye into the pores of the gels was ruled-out. CONCLUSIONS By taking advantage of the ability of ionic liquids to extract chitin or CRM of higher MWs directly from the corresponding biomass, we were able to prepare “physical” (no cross-linker) and/or “chemical” (using a cross-linker) hydrogels. The IL-extracted biopolymers of apparent higher MWs (CRM-175, IL-chitin) and PG-chitin could form both stable physical and chemical hydrogels, whereas lower MW polymers (Rec. PG, CRM-POM, and MCC) required a covalent cross-linker for hydrogel formation. Low MW pure chitin was not suitable for the preparation of hydrogels of either type. The hydrogels were supercritically dried producing amorphous, lightweight, and porous aerogels. Both chitin and cellulose aerogels showed high water absorbency, especially those prepared from CRM-175 and IL-1-10 and chemically cross-linked, again most likely due to the apparent higher MWs and lower chain entanglement (due to the incorporation of the cross-linker and the low biopolymer concentration). The results obtained from dye loading and release suggest the potential use of both chitin and cellulose hydrogels to effectively load and release


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active species. The gels with higher biopolymer concentration resulting in smaller pores exhibited increased dye loading capacity. As with water uptake, the release rate was mostly affected by the biopolymer concentration of the hydrogel, while the amount of dye and type of biopolymer had less effect. We believe that the use of chitin and cellulose hydrogels for drug delivery applications has great promise, but further research is needed. For example, the biocompatibility of CRM gels (the effect of lignin content), their mechanical properties, and lowering the burst release (through surface extracting or coating) should be investigated. Considering the unique high strength of cellulosic aerogels, we think that these materials may also find use in other fields, such as thermal insulation and tissue engineering. ASSOCIATED CONTENT Supporting Information Isolation of biopolymers with [C2mim][OAc], determination of purity of the biopolymers, measurement of the viscosity of biopolymer/NaOH/urea solutions, supercritical CO2 drying, calculation of ECH consumption, characterization methods (PXRD and SEM), and volumes and masses of all gels. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author *(R.D.R.) E-mail: [email protected] Notes


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Dr. Robin D. Rogers has partial ownership of 525 Solutions. Drs Gabriela Gurau and Julia Shamshina are part-time employees of 525 Solutions. The University of Alabama and McGill University maintain approved Conflict of Interest Management Plans. Dr. Wang’s current affiliation: Institute of Process Engineering, Chinese Academy of Sciences, No. 1 Beierjie Zhongguancun Haidian District, Beijing 100190, China. ACKNOWLEDGMENT The authors would like to thank 525 Solutions, Inc., the DOE SBIR Office of Science (DESC0010152) and the China Scholarship Council (No. 201306600007) for financial support. ABBREVIATIONS [C2mim][OAc], 1-ethyl-3-methylimidazolium acetate; CRM-175, cellulose-rich material isolated using IL at 175 °C; CRM-POM, cellulose-rich material isolated using IL with polyoxometalate (POM); ECH, epichlorohydrin; F/T, freeze/thaw; IL, ionic liquid; IL-chitin, chitin isolated from shrimp shells (SS) with IL; MCC, microcrystalline cellulose; MW, molecular weight; PG-chitin, practical-grade chitin; Rec. PG, reconstituted PG-chitin with IL; ScCO2-drying, supercritical CO2 drying.


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Comparison of Hydrogels Prepared with Ionic Liquid-Isolated vs. Commercial Chitin and Cellulose Xiaoping Shen,†,‡ Julia L. Shamshina,§ Paula Berton,†,║ Jenny Bandomir,† Hui Wang,† Gabriela Gurau§,║ Robin D. Rogers*,†,║

Synopsis Chitin and cellulose extracted from biomass using ionic liquid require lower concentrations than commercially available biopolymers to prepare physical and chemical hydrogels. TOC


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Comparison of Hydrogels Prepared with Ionic-Liquid-Isolated vs

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