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Bacterial Cellulose Ionogels as Chemosensory Supports Chip J. Smith, Durgesh Vinod Wagle, Hugh M. O'Neill, Barbara R Evans, Sheila N. Baker, and Gary A. Baker ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b12543 • Publication Date (Web): 10 Oct 2017 Downloaded from http://pubs.acs.org on October 11, 2017

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ACS Applied Materials & Interfaces

Bacterial Cellulose Ionogels as Chemosensory Supports Chip J. Smith II,† Durgesh V. Wagle,† Hugh M. O’Neill,‡ Barbara R. Evans,§ Sheila N. Baker,† and Gary A. Baker*,† †

Department of Chemistry, University of Missouri-Columbia, Columbia MO 65201 Biology and Soft Matter Division, Oak Ridge National Laboratory, Oak Ridge TN 37831 § Chemical Sciences Division, Oak Ridge National Laboratory, Oak Ridge TN 37831 ‡

ABSTRACT: To fully leverage the advantages of ionic liquids for many applications, it is necessary to immobilize or encapsulate the fluids within an inert, robust, quasi-solid-state format that does not disrupt their many desirable, inherent features. The formation of ionogels represents a promising approach, however, many earlier approaches suffer from solvent/matrix incompatibility, optical opacity, embrittlement, matrix-limited thermal stability, and/or inadequate ionic liquid loading. We offer a solution to these limitations by demonstrating a straightforward and effective strategy toward flexible and durable ionogels comprising bacterial cellulose supports hosting as much as 99% ionic liquid by total weight. Termed bacterial cellulose ionogels (BCIGs), these gels are prepared using a facile solvent-exchange process equally amenable to water-miscible and water-immiscible ionic liquids. A suite of characterization tools were used to study the preliminary (thermo)physical and structural properties of BCIGs, including no-deuterium nuclear magnetic resonance (no-D 1H NMR), differential scanning calorimetry (DSC), thermogravimetric analysis (TGA), scanning electron microscopy (SEM), and X-ray diffraction (XRD). Our analyses reveal that the web-like structure and high crystallinity of the host bacterial cellulose microfibrils are retained within the BCIG. Notably, BCIGs can not only be tailored in terms of shape, thickness, and choice of ionic liquid, they can also be designed to host virtually any desired active, functional species, including fluorescent probes, nanoparticles (e.g., quantum dots, carbon nanotubes), and gas-capture reagents. In this communication, we also present results for fluorescent designer BCIG chemosensor films responsive to ammonia or hydrogen sulfide vapors based on incorporating selective fluorogenic probes within the ionogels. Additionally, a thermometric BCIG hosting the excimer-forming fluorophore 1,3-bis(1-pyrenyl)propane was devised which exhibited a ratiometric (two-color) fluorescence output that responded precisely to changes in local temperature. The ionogel approach introduced here is simple and has broad generality, offering 1 ACS Paragon Plus Environment


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intriguing potential in (bio)analytical sensing, catalysis, membrane separations, electrochemistry, energy storage devices, and flexible electronics and displays. KEYWORDS Ionic liquids, ionogel, bacterial cellulose, fluorogenic probe, hydrogen sulfide, excimer

INTRODUCTION Ionic liquids (ILs) have captured the collective imagination of researchers across all branches of the physical sciences. Indeed, the unique features common to many conventional ILs (such as wide stable liquid range, extremely low vapor pressure, large electrochemical window, and excellent thermal stability, but most particularly their molecular tailorability) are responsible for their exploration for a wealth of purposes including those involving separations, liquid fuel desulfurization, biodiesel synthesis, CO2 capture, crystal engineering, battery technology, and pharmaceutics, among countless others.1-6 In spite of their unique attributes as a designer solvent or fluid, there are also a host of prospective uses wherein the liquid state is impractical or completely intolerable. For instance, for ease of handling or to avoid the issues of failure due to leakage, utilization in sensory platforms, gas separation membranes, or electrochemical/solar devices generally requires immobilization of the fluid phase.7-9 In order to expand the domain of real-world IL utility, it is imperative that supports be developed that confine ILs to allow for facile handling without sacrificing (and, ideally, actually enhancing) their original, positive attributes.10, 11 It is already known that ionogels (variously referred to an ion gels, ion-gels, or ion jellies) impart solid, self-standing, gel-like characteristics to ILs when they are immobilized or encapsulated within porous inorganic, organic, or hybrid support media.12 Alternative approaches 2 ACS Paragon Plus Environment

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to generate ionogels include use of a low-molecular-weight organic gelator,13 in situ free radical polymerization of compatible vinyl monomers (e.g., methyl methacrylate) in the presence of a small amount of a cross-linker,14 an IL solidification strategy that relies on gelation by selfassembly of suitable ABA architecture triblock copolymers,15 and nanofluid-inspired ionogels created by blending silica nanoparticles densely grafted with an IL functionality with a conventional IL phase.16 An IL confined or held inside these porous matrices offers the advantageous properties of a bulk IL, while presenting a monolithic character, opening up applications such as electrochemical platforms, actuators, separation and catalytic membranes, and drug delivery vehicles, which are otherwise impractical using a juxtaposed IL layer in a biphasic system, for example.10, 17, 18 However, earlier ionogel routes have certain limitations in terms of the sheer amount and types of IL that can be incorporated as well as imposing limitations on parameters like gel thickness, flexibility, durability, and thermal stability.19-22 For example, natural biopolymers (e.g., gelatin, agarose) generally require dissolution and reconstitution for ionogel preparation, resulting in loss of inherent structural integrity, yielding fragile, thermosensitive gels which lack the sturdiness required for many applications. Although cellulose provides a chemically and mechanically robust supporting scaffold for ionogel formation, there are limitations to prior approaches that suggest the need for significant advancements in cellulose-supported ionogel chemistry. All prior work on cellulose-supported ionogels has focused on plant-derived cellulose or modified cellulose (e.g., methyl cellulose). For example, Kadokawa et al. reported on a gel-like material formed from a 15 wt% solution of microcrystalline cellulose in 1-butyl-3-methylimidazolium chloride ([bmim]Cl) left standing for a week.23 Apart from the lengthy preparation, this approach is limited only to the use of a cellulose-dissolving IL such as [bmim]Cl (in this case, dissolution was performed at 100 °C), 3 ACS Paragon Plus Environment


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results in an almost complete disruption of cellulose crystallinity, and incorporates a large amount of water within the gels (~20 wt% based on thermogravimetric analysis). Buchtová et al. recently reported siloxane cross-linked (hydroxypropyl)methyl cellulose-based ionicallyconductive

ionogels

containing

1-propyl-1-methylpyrrolidinium

bis(trifluoromethylsulfonyl)imide.24 Although a water-immiscible IL was successfully entrapped as a biopolymer-based gel using this strategy, this preparation required a tedious graded-solvent exchange

sequence,

including

sacrificial

exchanges

with

the

hydrophilic

IL 1,3-

dimethylimidazolium methylphosphonate and acetonitrile. It should further be noted that the maximal IL loading was only 69 wt% and ionogel formation was accompanied by extreme volumetric shrinkage (~95% loss) and the development of conspicuous opacity. In a recent advance, Wunder and co-workers formed strong ionogels (MPa-range storage and loss moduli) encapsulating up to 97 wt% 1-butyl-1-methylpyrrolidinium bis(trifluoromethylsulfonyl)imide in nanofibrillar methyl cellulose networks by casting from N,N-dimethylformamide as a mutual solvent.25 In spite of these important developments, there remain limitations in the properties and chemistry of ionogels currently available. In an effort to overcome mechanical limitations and expand the scope of ionogels, we have begun to explore the utility of microbial cellulose networks for ionogel formation. Biosynthesis by some microbes (certain bacteria, fungi, and algae) is known to generate pristine cellulose absent characteristic vegetable components (e.g., lignin, hemicelluloses) which typically encase the cellulose microfibrils of plant cell walls. Bacterial cellulose (BC) is produced by strictly aerobic and non-photosynthetic gram-negative bacteria, principally of the genera Acetobacter, Sarcina ventriculi, and Agrobacterium, and is associated with different properties from plant cellulose, including a more crystalline structure with characteristic ribbon-like fibrils, higher 4 ACS Paragon Plus Environment

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purity, greater tensile strength with a temperature-invariant Young’s modulus, and improved water absorption capacity (BC hydrogels can hold up to 99 wt% water).26-28 BC is synthesized as an extracellular, swollen, gelatinous hydrogel mat, referred to as a pellicle, which forms a protective envelope serving to protect the bacterial cells from foreign materials whilst still allowing nutrients to be easily supplied by diffusion, and to aid in flocculation, maintenance of an aerobic environment, and the attachment to plants.29 The ultrafine network architecture of BC comprises thin microfibrils which are significantly smaller than those from plant cellulose (i.e., two orders of magnitude thinner than cellulose fibers produced by pulping wood), making BC more porous and imparting superior mechanical strength (Young’s modulus ~16.9 GPa).26, 29-31 Use of BC avoids the mechanical and thermochemical treatments such as pulping, fibrillation, and acid hydrolysis required to extract cellulose from plant sources. Given its extraordinary properties, BC has found widespread use in areas of research related to guided tissue regeneration, drug delivery, wound dressings, cosmetics, textiles, electrically-conductive films, flexible displays, acoustic membranes, and separations.28, 31-34 In this article, we present the first examples of bacterial cellulose-supported flexible ionogels (which we term bacterial cellulose ionogels) with an extremely high ionic liquid loading (up to 99 wt%) that avoid the many limitations seen with earlier ionogel approaches. In this new type of ionogel,

the

volume

spanning

network

comprises

BC

microfibrils

grown

from

Gluconacetobacter xylinus, a model microorganism selected for its ability to produce relatively high quantities of cellulose from a wide range of sources. Implementing a simple two-step solvent-exchange procedure, we prepared and studied bacterial cellulose ionogels (BCIGs) in which the IL phase percolating throughout the BC host was 1-butyl-1-methylpyrrolidinium bis(trifluoromethylsulfonyl)imide

([bmpy][Tf2N]),

1-ethyl-3-methylimidazolium 5

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bis(trifluoromethylsulfonyl)imide

([emim][Tf2N]),

or

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trihexyltetradecylphosphonium

bis(trifluoromethylsulfonyl)imide ([P14,6,6,6][Tf2N]) (Figure 1a). Notably, this approach is a general one and is applicable to virtually any IL of interest. By appropriate selection of the incorporated signaling chemistry, fluorescent sensor ionogels that respond to temperature and gas-phase ammonia or hydrogen sulfide were demonstrated. Coupled with the use of chemicallymodified BC (e.g., sulfation or phosphorylation), a plethora of soft advanced materials can be rationally built and a vast array of material properties aimed at using this strategy.

EXPERIMENTAL SECTION Materials and Reagents. Freeze-dried Gluconacetobacter xylinus bacteria (strain ATCC 700178) were obtained from ATCC (Manassas, VA). Yeast extract, peptone, and dextrose were purchased from Fisher Scientific (Waltham, MA). Sodium phosphate dibasic and citric acid were acquired from Sigma-Aldrich (St. Louis, MO). Mannitol was purchased from Calbiochem Merck (Billerica, MA) and 200 proof ethanol was obtained from Decon Laboratories (King of Prussia, PA). Ionic liquids (ILs) were synthesized in-house using previously reported methods.35 Bacterial Cellulose Hydrogel Growth. Gluconacetobacter xylinus bacteria were cultivated in a modified Hestrin-Schramm (HS) medium for BC production.36 The HS medium consisted of 2% w/v mannitol, 0.5% w/v yeast extract, 0.5% w/v peptone, 0.25% w/v sodium phosphate dibasic, and 0.15% w/v citric acid. The pH of the medium was adjusted to 6.0 by addition of 6.0 M HCl. The medium was then autoclaved at 125 °C followed by addition of 1% v/v ethanol using a sterile 0.2 µm nylon syringe filter (VWR; Radnor, PA). Starter bacterial cultures were prepared by placing 35 mL of HS medium in sterile 50 mL falcon tubes, followed by inoculation with freeze-dried G. xylinus from ATCC. The starter culture was allowed to stand static at 30 °C for 6 ACS Paragon Plus Environment

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three days and then used to initiate all other cultures. Specifically, 10% v/v of the starter culture was combined with fresh medium and new cultures allowed to grow for three days to result in the formation of the first cellulose layer (pellicle) on the top surface of the culture medium. Initial pellicles, due to inconsistency in growth, were discarded by either removal with sterile forceps or manual agitation of the culture medium to promote pellicle collapse to the bottom of the vessel. Cultures were then allowed to incubate statically at 30 °C for a certain period of time to allow for the formation of BC pellicles of desired thickness. BC pellicles were harvested with sterile forceps and placed in a 1.0 wt% NaOH solution and heated to 95 °C for 1 h to remove cellular debris. The pellicle was then rinsed in a water bath several times, changing the water every 2 h until the pH of the water tested neutral. At this point, a clean BC pellicle swollen with water (i.e., a hydrogel) was attained. BC Alcogel Preparation. Ethanol-containing BC gels (hereafter, referred to as alcogels) were prepared from BC hydrogels by soaking them in a large excess volume (>250 mL) of absolute ethanol for 12 h to result in the exchange of water by ethanol, changing the ethanol out after the first 6 h. At this point, the alcogels can be stored in ethanol indefinitely. Alcogels were then cut into 2 × 2 cm pieces approximately 3–4 mm in thickness by first cutting with a scalpel followed by completing cuts using Toennis Dissecting Scissors (MDS0861018, Medline, Northfield, IL). Gel thickness was measured using a Pittsburgh digital micrometer (Harbor Freight, item no. 68305) at the center of the alcogel. Because the alcogels were flexible and deformable, their thickness was determined at the point where the micrometer just started to displace a tiny amount of alcohol from the gel. The initial mass of an individual alcogel was measured after gently dabbing the external surface with a KimWipe to remove excess alcohol. The average wt% of BC

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within an alcogel was 1.2 % and was determined by weighing the residual BC following complete removal of alcohol by freeze-drying at –47 °C and 175 mTorr. Bacterial Cellulose Ionogel (BCIG) Preparation. BCIGs of varying thicknesses were prepared by adding appropriate amounts of IL (i.e., 10–200% of the mass of the BC alcogel) plus 0.5 mL of ethanol to pre-weighed alcogels in pre-cleaned glass vials. To illustrate the generality of this approach,

we

initially

selected

bis(trifluoromethylsulfonyl)imide bis(trifluoromethylsulfonyl)imide

three

ILs:

([bmpy][Tf2N]), ([emim][Tf2N]),

and

1-butyl-1-methylpyrrolidinium 1-ethyl-3-methylimidazolium trihexyltetradecylphosphonium

bis(trifluoromethylsulfonyl)imide ([P14,6,6,6][Tf2N]). The vials containing the alcogel, IL, and alcohol were capped, vortexed briefly, and allowed to sit overnight. The following day, the vials were uncapped and left open to allow for ethanol evaporation for 2–5, days depending on the laboratory humidity. Following ambient ethanol evaporation, samples were weighed, dimensions carefully measured, and then samples placed under vacuum for 12 h, followed by reweighing and remeasuring. Samples were dried under vacuum until a stable mass was achieved, indicating the complete evaporation of ethanol. The wt% of IL in the BCIG was determined using the following equation, assuming that the initial cross-sectional area and thickness are matched between the BC alcogel used to produce a BCIG and the BC alcogel dried to obtain the BC dry weight in eq 1.

wt% IL = (wt. of BCIG − dry wt. of BC from alcogel)/(wt. of BCIG) × 100

(1)


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We note that, occasionally, some gels showed signs of opacity under high ambient humidity. In such circumstances, additional ethanol was added to the vial containing the gel. The vial was capped to allow for equilibration (until the gel regained clarity) and then uncapped to allow for ethanol evaporation to resume. This approach generally proved effective for recovery of ionogels showing signs of cloudiness. Nuclear Magnetic Resonance (NMR). No-deuterium (no-D) 1H-NMR experiments were performed on an Oxford AS600 NMR magnet with a Bruker AVIII HD 600 MHz console using a 5 mm CPTCI cryo-probe. Samples were measured without spinning and in the absence of deuterated solvent. X-Ray Diffraction (XRD) Analysis. XRD measurements were performed on a Bruker Prospector instrument with an Apex II CCD detector and an IMuS micro-focus Cu tube. XRD measurements were carried out on a BC aerogel, neat [bmpy][Tf2N], and BCIGs containing 14– 90 wt% [bmpy][Tf2N]. XRD was measured from 2θ = 5–45° with a 0.1° step size using a polyimide capillary sample holder. XRD diffractograms were fit to Voigt functions using PeakFit® version 4.12. BC XRD data were first fit in order to determine parameters for subsequent BCIG fitting. Data were fit to five crystalline peaks and an amorphous peak (Figure S1) following a previous report using a Voigt function for all peaks to calculate the crystallinity index (CrI) instead of the peak height method, which has been shown to overestimate the crystallinity.37 The parameters were set with a 3% tolerance in the linear baseline fit, using a deconvolution width of 0.4, standard deviation (SD) setting, a filter setting of 71.2, with varying widths, and a 7.00 Amp%. CrI was calculated by dividing the integrated area of the BC crystalline peaks by the total area of the BC XRD peaks (Figure S2). The BC diffractogram



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fitting was also used to supply the parameters for the Scherrer Equation (eq 2) in order to calculate crystallite size (τ).38 = /(! cos #)

(2)

In eq 2, τ is the size perpendicular to the lattice plane represented by the peak in question, K is a constant that depends on the crystal shape (K is often close to 1), λ is the wavelength of the incident beam in the diffraction experiment, β is the full width at half-maxima (FWHM) in radians, and θ is the location of the peak.39,

40

The d-spacing was calculated using the Bragg

equation (eq 3) assuming a first order of diffraction.40, 41 $%&' = /2 sin #

(3)

Neat [bmpy][Tf2N] diffractograms were fit by using Voigt functions as well, which allowed for an r2 value of 0.9993 while using as few peaks as possible to adequately fit the subsequent data. Fits to IL XRD data consisted of three peaks (Figure S3). The first peak at the lowest 2θ was present in all BC and BCIG samples as well, suggesting that it could be a background peak that was not completely removed in background subtraction. The two higher angle peaks, however, are commonly observed for pyrrolidinium cation containing ILs and arise from intermolecular interactions within the IL.42 BCIG samples were fit by first importing the fitting parameters and peaks from the BC fit and then adding the appropriate additional peaks to account for the presence of IL (Figures S4– 7). Each BC peak was locked at the 2θ position determined from the BC fit and IL peak positions were allowed to float. Fitting of the BCIG diffractograms was iterated until an r2 value of 0.9980 was obtained. All locked peaks were then unlocked and fit iteration resumed until r2 values of >0.9990 and F values of >60,000 were achieved.


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Differential Scanning Calorimetry (DSC). Differential scanning calorimetry (DSC) was measured under a nitrogen atmosphere using a TA Instruments model DSC Q100 fitted with a liquid nitrogen cooling system. Samples weighing 8–12 mg were hermetically sealed in aluminum pans and heated to 353 K followed by cooling to 123 K and finally heated back to 353 K, all at a rate of 10 K min–1. In order to eliminate effects from thermal history, DSC thermograms from the second heating cycle are presented and used for analysis. The crystallization temperature, Tcr, and the melting temperature, Tm, were taken at the onset of the respective transitions.43 Thermogravimetric Analysis (TGA). TGA analysis was performed on a Q50 analyzer (TA Instruments, Inc.) ramping from room temperature up to 873 K at a constant heating rate of 10 K min–1 under nitrogen flow. The decomposition temperature, Tdcp, was taken at the onset of decomposition which is defined as the temperature at which 10% mass loss had occurred.43 Scanning Electron Microscopy (SEM) Imaging. SEM imaging was performed on a FEI Quanta 600 FEG environmental scanning electron microscope equipped with a Schottky field emitter (thermal FEG). BC aerogel samples prepared by supercritical CO2 drying on a Tousimis Autosamdri-815 CPD, were imaged in low-vacuum mode at a pressure of 23 Pa and voltage of 10 kV. BCIGs were imaged under high vacuum and 5 kV while the other SEM settings remained the same as the BC aerogel samples. We were unable to image BCIG samples at high resolution due to instability of the gel in the electron beam. This could be largely avoided by incorporating lesser amounts of IL, however, this defeats our purpose which is to image a representative BCIG showing the swollen morphology maintained by the IL. Fluorescence Spectroscopy. Fluorescence measurements were performed on a Varian Cary Eclipse fluorescence spectrophotometer (Agilent Technologies, Inc.) equipped with a Nes Lab 11 ACS Paragon Plus Environment


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RTE-111 recirculation temperature control bath which allowed temperature control to within ± 1 °C. Gas-phase NH3 detection. BCIG ammonia (NH3)-sensing platforms were prepared by soaking 1.5-mm thick 2 × 2 cm alcogels in 1.17 mL of a 1:8 (v/v) [P14,6,6,6][Tf2N]/ethanol solution containing 1.7 µM [P14,6,6,6]3[HPTS] (see Figure S8a), followed by alcohol evaporation to afford ~0.9 mm-thick BCIGs containing 98 wt% [P14,6,6,6][Tf2N] and 13 µM [P14,6,6,6]3[HPTS]. As summarized in Figure S9, NH3 gas was produced by reacting 4.0 g of NH4Cl powder with one equivalent of NaOH (both of which had been individually ground previously using an agate mortar and pestle) inside a single-neck 50 mL round bottom flask sealed with a rubber septum. Initially, the gas was vented into the headspace of an inverted water-filled 250 mL volumetric flask and placed in a crystallization dish containing slightly-acidic water dyed with phenolphthalein. The gas was vented into the flask in order to maintain a pressure in the reaction flask of approximately 1 atm. After allowing the reaction to proceed for 15 min under mild manual agitation, the water in the volumetric flask had been completely displaced into the crystallization dish, initiating NH3(g) bubbling which resulted in the phenolphthalein-spiked water becoming pink. At this point, NH3(g) was collected from the reaction vessel via syringe as the reaction continued. Once gas bubbling from the crystallization dish resumed, the syringe was removed from the septum and the needle immediately stoppered. After collection of the gas, the volume was used in the ideal gas law equation to calculate moles of NH3, assuming ambient pressure and temperature for the gas. To perform NH3(g) sensing, the gas in the syringe was injected through the septum of a sealed quartz cuvette using a stainless steel 26-gauge needle connected to plastic tubing (REF# 682073, BD Medical, Franklin Lakes, NJ) as presented in Figure S10. The cuvette contained a 12 ACS Paragon Plus Environment

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45°-oriented glass slide with an NH3-sensing BCIG adhered to its surface. The sensing system was also equipped with a similar waste stream syringe. Fluorescence was measured 2 min after each 0.1 mL addition of NH3(g), although a complete response was achieved in under 30 s. The gel was excited at 407 nm and emission was scanned from 420 to 600 nm. To determine the primary peaks of interest in the difference spectrum, spectra were taken before and after saturation with NH3 and the initial fluorescence intensity (F0) was subtracted from the NH3saturated fluorescence intensity (Fsat). The ratio of the intensities measured at these sodetermined spectral positions (i.e., F511/F445) was then used to plot the NH3(g) response. Fluorescence temperature monitoring. This study was conducted using 2-mm thick BCIGs comprising 25 µM 1,3-bis(1-pyrenyl)propane (BPP) dissolved in [bmpy][Tf2N] accounting for 99 wt% of the total gel mass. The BCIG was prepared by adding 84.5 µL of a 100 µM BPP in ethanol solution to 0.3 mL of ethanol, 0.476 g of [bmpy][Tf2N], and a BC alcogel. After incubation and drying, samples were placed on a solid aluminum block set at 45° to the excitation beam direction. For comparison, a solution of [bmpy][Tf2N] containing 5 µM BPP was also prepared and measured. The fluorescence spectra were blank subtracted using a corresponding probe-free BCIG or [bmpy][Tf2N]. Samples were excited at 325 nm and emission was monitored from 350 to 600 nm for excitation/emission slits of 10/10 nm. The fluorescence spectra were collected at 10 °C increments from 30 to 120 °C. The sample temperature was allowed to equilibrate for 10 min prior to fluorescence measurement.

RESULTS AND DISCUSSION BC and BCIG Preparation. The bacterial strain Gluconacetobacter xylinus (ATCC 700178) was chosen for this study because cellulose synthesized by this particular strain is well known to display values for mechanical strength, water absorption capacity, and crystallinity, as well as 13 ACS Paragon Plus Environment


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possessing an ultra-fine, pure fibrous network structure.44 ILs were incorporated into the BC network using a simple strategy based on diffusion-displacement of a mutually-compatible cosolvent such as ethanol, followed by evaporation of said co-solvent (Figure 1b) to leave behind BC gels swollen with an IL component. BCIGs were chemically stable and found to support up to 99% of the IL by weight, without loss of structural integrity. Even for high loadings, BCIGs were extremely flexible (Figure 2a), being amenable to manual bending and contortion with negligible loss of the sequestered IL phase. The flexibility and robustness of the BCIGs derives from the continuous interconnected network of cellulosic microfibrils in BC (Figure S11) which is absent in reported ionogels, which are generally more fragile or less compliant.21, 23, 45 A series of BCIGs of varying thickness (Figure 2b, inset) were prepared by manipulating the amount of IL incorporated or by intentionally culturing the cellulose pellicles to particular thicknesses. The negligible vapor pressures of the ILs allowed for thickness manipulation of the BCIGs with no discernible change in size or shape for more than 6 months under ambient storage. The IL phase could also be easily recovered by co-solvent extraction at any point, similar to previous reports of silica-based ionogels.46 While planar cellulose pellicles can be cut into desired shapes after harvesting, more interestingly, they can actually be grown into arbitrary cross-sectional shapes and sizes, provided a suitable template is available. A similar process has been reported previously for growing tubular bacterial cellulose.47 An example of this templated growth is illustrated in panels c and d of Figure 2 showing BCIGs prepared from individual pellicles grown using stainless steel letter-shaped molds placed at the surface of the BC culture medium (in this case, the letters M-I-Z signify Mizzou, our institutional nickname since 1905). Importantly, in addition to catering the ionogel geometry to the particular needs of device applications and testing equipment (e.g., dynamic mechanical analysis), this templated growth 14 ACS Paragon Plus Environment

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strategy also avoids the necessity of casting ionogels from dissolved cellulose solutions, which can degrade the natural highly crystalline structure of BC.7,

19

The individual letter-molded

BCIGs were differentially stained with unique fluorescent dyes (Nile Red, rhodamine 6G, and pyrene, respectively) to illustrate the utility of BCIGs for display and light-emitting applications (Figure 2c,d). Notably, the chemically-inert nature of the BCIGs and the involatility of ILs similarly allows for the incorporation of other task-specific molecules (e.g., bioactive molecules, nanoparticles, polymers) within the ionogels, introduced via co-solvent assisted diffusion followed by co-solvent removal by evaporation.48, 49 1

H-NMR. No-deuterium (no-D) 1H-NMR was used in this study to directly compare the neat IL

with IL confined within a BCIG. As expected, ILs incorporated in BCIGs exhibit peak broadening (Figure 3a), a phenomenon which has been similarly observed for ILs confined in silica-based ionogels.50, 51 The characteristic peak broadening seen for [emim]+ and [bmpy]+ ILs (assigned in the Supporting Information, SI) upon incorporation in BCIGs likely stems from specific interactions between the IL components and cellulose fibers, restricting the diffusional dynamics of the IL cations (noting that the [Tf2N]– anion contains no protons, it is spectroscopically silent in 1H NMR).7,

17

This spectral broadening is noteworthy given the

mesoscopic confinement within large pores and voids within the BC and the fact that the BCIGs predominantly comprise IL (above 99 wt%) in these samples, suggesting long-range structural perturbations introduced by interfacial interactions. Although beyond the scope of the present investigation, the mesoscopic confinement of ILs in BCIGs forms the subject of current study in our group. Thermal Characterization. The differential scanning calorimetry (DSC) scans for BCIGs (taken from the second heating cycle) reveal a reduction in the melting (Tm) and crystallization 15 ACS Paragon Plus Environment


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(Tc) temperatures compared to that of the bulk liquid (Figure 3b). Neat [bmpy][Tf2N] presents complex melting behavior, showing the presence of two peaks whereas the corresponding BCIG for contents of 98.6 wt% [bmpy][Tf2N] and higher reveal a slightly reduced melting point (by ~6–10 K) with a loss in melting complexity (Figure S12).42, 52 This decreased complexity for the BCIGs is attributed to changes in molecular ordering due to surface interactions with the cellulose fibers under mesoscopic confinement. This perturbation in apparent Tm and Tc for ILs confined in BCIGs (Table S1) further confirms the strong interactions between IL components and the fibrous cellulosic network, leading to significant alteration in the bulk solvent properties, an observation made earlier for IL confinement within silica ionogels.53,

54

It is peculiar that

ionogels containing

Bacterial Cellulose Ionogels as Chemosensory ... - ACS Publications

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