Article pubs.acs.org/Langmuir
Honeycomb Films of Cellulose Azide: Molecular Structure and Formation of Porous Films William Z. Xu and John F. Kadla* Advanced Biomaterials Chemistry Laboratory, University of British Columbia, Vancouver, British Columbia V6T 1Z4, Canada S Supporting Information *
ABSTRACT: Development of value-added micropatterned porous materials from naturally abundant polymers, such as cellulose, are of growing interest. In this paper, regioselectively modified amphiphilic cellulose azide, 3-O-azidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose, with different degrees of substitution (DS) and degrees of polymerization (DP) of the poly(ethylene glycol) (PEG) side chain, was synthesized and employed in the formation of honeycomb-patterned films. With the variation of the DP and/ or DS, the amphiphilicity of the polymer and the pore size of the formed films changed accordingly. It was found that amphiphilicity of the cellulose azide played a significant role in the formation of honeycomb films. Balanced amphiphilicity was of particular importance in the formation of uniform honeycomb films. Via the CuI-catalyzed alkyne−azide [2 + 3] cycloaddition reaction, fluorescent avidin and quantum dots were attached to the films. By means of confocal microscopy, it was confirmed that the functional azido group was preferentially allocated inside the pores. This provides a platform for the development of advanced honeycomb materials with site-specific functionalities, such as biosensors.
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films.17 The number of arms in star polymers and the number of repeating units in diblock co-polymers significantly affected the honeycomb film features.15,18 Amphiphilic polymers are excellent candidates because of their ability to encapsulate the water droplets, prevent the water droplets from coalescence, and thus, stabilize the arrays of water droplets.16,19 However, to the best of our knowledge, there has not been any report on the effect of the amphiphilic polymer brush structure, specifically brush length and frequency, on the formation of honeycomb films via the “breath figures” method. To develop advanced honeycomb materials for a variety of applications, functionalization of the honeycomb films is an extremely important step. Thermo-sensitive20 and pHsensitive21 honeycomb films as well as honeycomb films with nanofillers, such as ZnO nanoparticles,22 have been reported. Because of the heterogeneous structure of honeycomb films, allocation of the functional groups on the honeycomb films is a key to success in some applications.23 For instance, the interaction of biomolecules, such as proteins and antibodies, in localized microdomains is of particular importance in the development of biosensors and biomedical devices and the research on cell growth and adhesion, tissue engineering, etc.24,25 Because of the formation mechanism of honeycomb films, the polar moieties of an amphiphilic glucopolymer were mainly oriented inside the pores.26,27 By employing an amphiphilic temperature-responsive block co-polymer, poly-
INTRODUCTION Development of highly ordered meso- and macroporous materials has become a hot research topic because of their potential applications in separation and dialysis,1 drug delivery,2 sensors,3,4 photonics,5,6 optoelectronics,7 catalysis and electrochemistry,8 biosensors,9 tissue engineering,10 etc. The porous materials can be fabricated by many methods, including photolithography,11 plasma-enhanced chemical vapor deposition (PECVD),12 and inverse microemulsion.13 Among the approaches based on bottom-up self-assembly, the “breath figures” method14 has proven to be an effective dynamic template method because of the simple, inexpensive, and robust mechanism of pattern formation. There are three major steps involved in the “breath figures” process:15,16 (1) when moist air is blown over a solution of a polymer in a highly volatile, water-immiscible, organic solvent, evaporative cooling leads to the formation of water droplets (nucleation and growth) on the liquid surface; (2) the water droplets arrange into a hexagonal array and sink into the polymer solution; and (3) removal of the solvent and water leaves an imprint of the water droplets as a hollow air-filled, hexagonally ordered, polymeric bubble array. There are many external factors affecting this process, including polymer concentration, relative humidity, solvent properties, and substrate.15 It should not be surprising that the only internal factor, i.e., polymer itself, plays a key role in the formation of honeycomb films. A variety of polymers and different macromolecular architectures, such as linear, star, graft, dendritic, hyperbranched polymers and coil−coil or rod−coil diblock co-polymers, were employed to form honeycomb © 2012 American Chemical Society
Received: September 24, 2012 Revised: December 18, 2012 Published: December 20, 2012 727
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Bovine serum albumin (30% aqueous solution) was purchased from EMD Millipore and used as received. Tetrahydrofuran (THF, Fisher Scientific), acetone (ACS certified, Fisher Scientific), cellulose (Fluka, Avicel PH-101, ∼50 μm particle size), and anhydrous lithium chloride (Sigma-Aldrich) were dried prior to use. Characterization. 1H and 13C nuclear magnetic resonance (NMR) spectra were measured using a Bruker AVANCE-300 spectrometer at 25 °C (small molecules) or 40 °C (polymers). Samples were dissolved in CDCl3, and chemical shifts were referenced to tetramethylsilane (TMS; 0.0 ppm). The DS of the PEG side chain in the synthesized 3O-azidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose was determined with 1H NMR, as described in the previous published paper.32 Attenuated total reflection−Fourier transform infrared (ATR−FTIR) spectra were obtained with a Perkin-Elmer Spectrum One FTIR spectrometer. Spectra were recorded at a resolution of 4 cm−1 and a total of 32 scans. The surface morphology of the cast films was observed with a Hitachi S-2600N scanning electron microscope (SEM). The pore size was measured on SEM images using ImageJ software (NIH, Bethesda, MD), and the statistical analysis of the pore size distribution was performed on the basis of more than 3000 pores for each sample by means of MATLAB software. Confocal images were collected on an Olympus FV-1000 confocal microscope with a 60×, 1.42NA objective. For the fluorescent avidin sample, the 559 nm laser was used for fluorescence excitation and emission was detected between 570 and 670 nm. For the QD sample, the 405 nm laser was used for fluorescence excitation and emission was detected between 480 and 630 nm. The confocal pinhole was set at 180 μm, equivalent to 1 optical unit (OU). Synthesis of Cellulose Azide. The synthesis of cellulose azide with three different PEG side-chain lengths (DP) is described in a previously published paper.32 PEG MW = 200 (EG4), PEG MW = 600 (EG13), and PEG MW = 1000 (EG22) were first protected by attaching an allyl group to one end of PEG to form allyloxypoly(ethylene glycol). The hydroxyl group at the other end of the PEG was then converted to the corresponding tosylate, followed by further conversion of the tosylate to the corresponding iodide. Cellulose was regiospecifically functionalized with TDMSCl to form 2,6-di-Othexyldimethylsilyl cellulose. From the reaction of the synthesized allyloxypoly(ethylene glycol) iodide with 2,6-di-O-thexyldimethylsilyl cellulose, amphiphilic 3-O-allyloxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose was synthesized. The allyloxy group at the free end of the PEG side chain was further oxidized to the hydroxypropoxy group, followed by conversion to the chloropropoxy group, and finally, the azidopropoxy group. To synthesize 3-O-azidopropoxypoly(ethylene glycol)-2,6-di-Othexyldimethylsilyl cellulose with different DS of PEG, a smaller amount of allyloxypoly(ethylene glycol) iodide was employed in the synthesis of 3-O-allyloxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose. It should be noted that, for the low DS samples, the concentration of the functional groups at the free end of the PEG side chain in a solution was relatively low. Employing the same molar ratios of reagents would result in incomplete conversion. Hence, to achieve complete conversion of the low DS samples, the concentration of the reagents should be maintained in a similar level as used in the conversion of high DS samples. Preparation of Honeycomb Films of 3-O-Azidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl Cellulose. Honeycomb films were prepared by applying 10 μL of neat solution (1% in toluene or otherwise described) onto a glass slide in a humid environment (flow rate, 700 mL/min; relative humidity, 70−80%; and room temperature).35 Attachment of Biotin/Avidin to the Honeycomb Film of 3-OAzidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl Cellulose. Alkynated biotin was synthesized according to the procedure by Lin et al.36 with a simplified workup procedure. Linking biotin to the surface of the honeycomb film is described in detail in ref 32. For the conjugation of avidin and the biotinylated film, avidin solution (500 μg of avidin−sulforhodamine 101 in 2.5 mL of pH 7 buffer) was dropwise introduced onto the porous films (the biotinylated film and a control non-biotinylated film). Each
styrene-block-poly(N-isopropyl acrylamide) (PS-b-PNIPAAm), Stenzel et al.15 confirmed that the surface of the pores was enriched with the hydrophilic PNIPAAm groups. By means of the conjugation of the β-galactose moieties and fluorescent peanut agglutinin (PNA), Ting et al.26 developed patterned protein onto the honeycomb films of an amphiphilic diblock co-polymer, polystyrene-block-poly(2-(β-D-galactosyloxy)ethyl methacrylate-co-styrene). Min et al.28 immobilized a protein, fluorescent streptavidin, inside the hydrophilic pores of honeycomb films of an amphiphilic block co-polymer, polystyrene-block-poly(acrylic acid). Sunami et al.29 reported that fibronectin, a typical protein as a cell adhesion molecule, was site-specifically absorbed on the inside of the pores of poly(ε-caprolactone) (PCL) honeycomb film. Site-specific modification of honeycomb films is an important and challenging area of research. When amphiphilic polymers are employed in the formation of honeycomb films, it was expected that the hydrophilic groups could preferentially be allocated inside the pores. Linking functional groups to the hydrophilic moieties can effectively position the functional groups inside the pores. As the most abundant natural polymer, cellulose has been widely researched in the development of advanced materials.30,31 In our previous work,32 honeycomb films were prepared with an amphiphilic regioselective cellulose derivative 3-O-azidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose. In the present research, we aim to study the effect of amphiphilicity of the brush-shaped cellulose derivative on the formation of honeycomb films. Specifically, the cellulose azide with different degrees of substitution (DS) and different degrees of polymerization (DP) of the pendent poly(ethylene glycol) (PEG) side chain was synthesized and employed in the formation of honeycomb films, followed by examination of the effect of DP and DS on the pore structure of the formed films. The location of the functional azido group on the films was also examined with confocal microscopy by means of attaching a fluorescent protein and quantum dots (QDs) to the films. This provides a platform for site-specific functionalization of the cellulose azide films with many functional molecules containing an alkynyl group via “click chemistry”.33,34
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EXPERIMENTAL SECTION
Materials. PEG MW = 200 (EG4), PEG MW = 600 (EG13), PEG MW = 1000 (EG22), dimethylthexylsilyl chloride (TDMSCl, 95%), imidazole, anhydrous N,N-dimethylacetamide (DMA), anhydrous N,N-dimethylformamide (DMF), anhydrous dimethyl sulfoxide (DMSO), p-toluenesulfonyl chloride (tosyl chloride), sodium azide (NaN3), sodium hydride (60% dispersion in mineral oil), sodium thiosulfate (pentahydrate), triphenylphosphine (PPh 3 ), 9borabicyclo(3.3.1)nonane (9-BBN, 0.5 M in THF), hydrogen peroxide (35%), carbon tetrachloride, (+)-biotin N-hydroxysuccinimide (biotinNHS, ≥98%), propargylamine (98%), triethylamine (≥99%), avidinsulforhodamine 101 (fluorescent avidin), 6-heptynoic acid (90%), Nhydroxysuccinimide (NHS) (98%), N-(3-dimethylaminopropyl)-N′ethylcarbodiimide hydrochloride (EDC hydrochloride) (commercial grade, powder), and tetra-n-butylammonium iodide (TBAI) were purchased from Sigma-Aldrich and used as received. Potassium hydroxide, sodium hydroxide, anhydrous magnesium sulfate, potassium iodide, toluene [high-performance liquid chromatography (HPLC) grade], chloroform (ACS certified), methylene chloride (ACS certified), ethyl ether (ACS certified, 99.9%), sodium chloride (ACS certified), sodium bicarbonate (ACS certified), acetonitrile, L(+)-ascorbic acid, and cupric sulfate pentahydrate (ACS certified) were purchased from Fisher Scientific and used as received. QDs (545 ITK aminoPEG, 8 μM solution) were purchased from Invitrogen. 728
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introduction (drop of 50 μL) was left to infuse into the porous film before subsequent drops. The step was repeated 3 times. Unbound avidin was removed by washing 3 times with pH 8.5 buffer and 3 times with albumin solutions. Attachment of QDs to the Honeycomb Film. 6-Heptynoic acid succinimidyl ester was synthesized by following the procedure by Luo et al.37 6-Heptynoic acid (90%, 1 g) was dissolved in 100 mL of dried methylene chloride, followed by the addition of NHS (2.3 g) and EDC hydrochloride (2.3 g). The resulting reaction mixture was stirred at room temperature for 18 h, at which point 500 mL of saturated sodium bicarbonate aqueous solution was then added. The methylene chloride phase was collected before extracting the aqueous phase with 5 × 100 mL of ethyl ether. The ether extract was combined with the methylene chloride phase, washed with 3 × 165 mL of water and 3 × 165 mL of brine, and then dried with 20 g of anhydrous magnesium sulfate. A white solid product was obtained after filtering off magnesium sulfate and removing the solvents by rotary evaporation. Yield = 81%. FTIR (cm−1; ν, stretching mode; δ, bending mode): 3255 (νCH), 2945 (νCH2(as)), 2865 (νCH2(s)), 2259 (νCC), 1817 (amide, νCO), 1779 (amide, νCO), 1723 (ester, νCO), 1431 (δCH2), 1382, 1218 (νC−O), 1079, 996, 887, 812, 760, 650. 1H NMR (CDCl3, 300 MHz) δ (ppm): 1.65 (ddd, J = 14.8, 7.34, and 7.23 Hz, 2H), 1.89 (ddd, J = 15.13, 7.45, and 7.23 Hz, 2H), 1.98 (t, J = 2.85 Hz, 1H), 2.25 (td, J = 7.02 × 2 and 2.63 Hz, 2H), 2.65(t, J = 7.23 Hz, 2H), 2.84 (s, 4H). 13C NMR (CDCl3, 75.4 MHz) δ (ppm): 18.0 (HCC−CH2), 23.5 (HCC−CH2−CH2−CH2), 25.6 (N−C(O)−CH2), 27.3 (HCC−CH2−CH2), 30.4 (CH2−COO), 68.9 (HCC), 83.4 (HCC), 168.3 (O−CO), 169.1 (N−CO). The synthesized 6-heptynoic acid succinimidyl ester was reacted with the film of 3-O-azidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose. To a solution of 75 mg (0.34 mmol) 6-heptynoic acid succinimidyl ester in 2 mL of DMSO was added 4.3 mg (0.017 mmol) of cupric sulfate pentahydrate and 6.0 mg (0.034 mmol) of L(+)ascorbic acid. The formed solution was dropwise-introduced onto the porous film. The reaction mixture-coated film was then sealed in a glass container and placed in an oven at 50 °C for 4.5 days. After the film was washed with water and methanol, it was dried in air and then characterized with FTIR spectroscopy. FTIR (cm−1): 3514 (νOH), 2957 (νCH3(as)), 2870 (νCH3(s)), 2099 (νN3(as)), 1738 (νCO), 1638, 1465 (δ C H 3 , C H 2 ( a s ) ), 1378 (δ C H 3 ( s ) ), 1251 (δ S i − C ), 1115 (ν Si−O−C, C−O−C(EG) ), 1079, 1036 (ν C−O−C(AGU) ), 832, 779 (νSi−C/Si−O−C). Prior to the reaction with QDs, 75 μL of 8 μM solution of ITK 545 aminoPEG QDs was washed with pH 7 phosphate-buffered saline (PBS) buffer solution using Amicon Ultra (0.5 mL, 100 K) centrifugal filters (Millipore) and resuspended in 50 μL of pH 7 PBS buffer solution. The QDs in buffer solution were further diluted with 50 μL of acetonitrile before being introduced onto the dried 6-heptynoic acid succinimidyl ester-clicked film. After the film was sealed in a container at room temperature for 22 h, it was washed with water and then dried in air.
Figure 1. Correlation between the pore diameter and PEG length for cellulose azide samples with high and low DS.
hydrophilic PEG side chain. However, this trend was not completely maintained for low DS (0.11−0.13) samples. For these low DS samples, the pore size varied slightly with an increasing PEG length from EG4 to EG22. For samples with low DS and short PEG side chain, they are quite hydrophobic; therefore, they have poor stability for the growing water droplets. When the PEG length is increased, they became more hydrophilic, resulting in the better ability to stabilize the growing water droplets during the formation of honeycomb films. The effect of DS on the pore size is displayed in Figure 2. For samples with a relatively short PEG chain (EG4 and EG13), the
Figure 2. Correlation between the pore diameter and degree of substitution for three cellulose azide samples with different PEG lengths.
pore size increased with an increasing DS. This trend became very significant for the sample with short PEG length (EG4), because the pore diameter increased by more than 70% when DS increased from 0.11 to 0.47. This phenomenon confirmed that increasing hydrophilicity of hydrophobic samples improved their stability for growing water droplets. However, for samples with a relatively long PEG chain (EG22), the pore size decreased slightly with an increasing DS from 0.13 to 0.45. Because of their long PEG chain, more micelles were formed during the nucleation process with an increasing number (DS) of the hydrophilic PEG chain, resulting in a smaller pore size. According to the above discussion, the very hydrophobic samples with a small number of EG units, e.g., EG4 and DS = 0.11, formed small pores and the pore size increased with an increasing DS and/or PEG length. On the other hand, the very hydrophilic samples with a large number of EG units, e.g., EG22 and DS = 0.45, also formed small pores and the pore size increased with a decreasing DS and/or PEG length. Hence, the balance of amphiphilicity seems to be an important factor in the
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RESULTS AND DISCUSSION Formation of the Honeycomb Films. As described in a previously published paper,32 the synthesized amphiphilic 3-Oazidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose is ready for the formation of the honeycomb film. However, there are many factors affecting this process, including the molecular structure, solution concentration, and casting conditions. The effect of the molecular structure, such as the DP and the DS of the PEG side chain and the solution concentration, was studied. Figure 1 demonstrates the relationship between the pore diameter and the chain length of PEG. For high DS (0.37− 0.47) samples, the pore size decreased with an increasing PEG length. This can be explained because more micelles were formed during the nucleation process because of their longer 729
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formation of honeycomb films. A summary of SEM images of the formed honeycomb films is shown in Figure 3. The images
The effect of the concentration on the pore size was also studied. For the sample with a short PEG chain (EG4) and high DS (0.47), the mean pore diameter decreased from 2.61 to 2.20 μm when the concentration increased from 1 to 2% (see the Supporting Information). With the same PEG chain length (EG4), the sample with low DS (0.11) showed the same trend. The mean pore diameter decreased from 2.45 to 1.52 μm when the concentration increased from 0.3 to 1% (see the Supporting Information). Stenzel at al.15 stated that the pore size increased with a decreasing concentration and provided a correlation between the pore size (PS) and the polymer concentration (c), which is close to PS = k/c, where k is a constant that is dependent upon the material used. Our observation was in good agreement with the trend but did not strictly follow this correlation. In addition, a second porosity in the honeycomb films of their amphiphilic block co-polymers reported by Beattie et al.38 and Escale et al.39 was not observed with our PEG-functionalized amphiphilic cellulose azide films in the present study. In comparison to the honeycomb films of the amphiphilic block co-polymers, our films had a much smaller wall thickness/pore size ratio. This may be due to the high density of hydrophilic PEG brushes surrounding a relatively slim hydrophobic backbone and the well-distributed hydrophilic groups in the molecular structure of our regioselectively modified cellulose azides. The thinner wall may better inhibit the formation of the aforementioned second porosity in the honeycomb films. To study the effect of amphiphilicity by varying the DS and DP on the formation of honeycomb films, the casting conditions were remained fixed. Hence, although the mean pore diameter measured in the present study was between 1.2 and 2.6 μm, honeycomb films with a wider range of pore sizes could be obtained by varying the casting conditions, such as the concentration, solvent system, humidity, air flow rate, etc.40 Allocation of the Azido Group on the Films. When the “breath figures” method was employed, honeycomb films were formed with the synthesized amphiphilic 3-Oazidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose. The importance of amphiphilicity of the polymer lies in the stabilization of water droplets by the hydrophilic PEG segments and the formation of a surface enriched with the hydrophobic backbone. In theory, the azido functional group at the end of the hydrophilic PEG side chain should be preferentially self-directed surrounding the water droplets and finally inside the pores. To prove this hypothesis, two experiments were conducted with the honeycomb film of 3O-azidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose, with DS being 0.37. The medium DP (EG13) was selected because this should be more representative among the three, while higher DS (0.37) was selected because of its higher reactivity in the click reaction. Avidin is a tetrameric glycoprotein, which can bind up to four biotin molecules. The avidin−biotin complex has one of the strongest known non-covalent bonds, with an absolute (standard) free energy of binding ΔAo = −85.4 kJ/mol and dissociation constant Kd ∼ 10−15 M.41 Each avidin chain is arranged in an eight-stranded antiparallel β-barrel, in which biotin is bound. The tryptophan residues 70, 97, and 110 form part of the avidin binding site and are anchored through hydrogen bonds to other residues, leading to stabilization of the binding site.42 As described in the previous work,32 biotin-NHS was first reacted with propargylamine to form alkynated biotin. The
Figure 3. SEM images of the honeycomb films of cellulose azide with varying DS and PEG lengths.
on the lower left corner represent the most hydrophobic sample, while the images on the upper right corner represent the most hydrophilic sample. Not only are the pores on the two corners smaller than those on the other area, but they are also less uniform than those on the other area. Hence, balance of amphiphilicity played a key role in the formation of the honeycomb film, in the way of both pore size and regularity. An obvious question exists, and that is if similar amphiphilicity would result in similar pore structure. To answer this question, samples with a similar number of EG units but different PEG lengths were compared. Figure 4
Figure 4. Comparison of the pore size distribution for samples with different PEG lengths but similar DS × DP value (a and b, 1.69−1.88; c and d, 0.91−1.00). Note: ⟨φ⟩, mean pore diameter; σ, standard deviation.
displays the pore size distribution of two groups of samples. The first group has a DS × DP value between 1.69 and 1.88 (panels a and b of Figure 4), while the second group has a value between 0.91 and 1.00 (panels c and d of Figure 4). In both groups, samples with shorter PEG chains (panels a and c of Figure 4) formed honeycomb films with a larger pore size than those with longer PEG chains (panels b and d of Figure 4). Hence, samples with a similar amphiphilicity but longer PEG side chains tend to form smaller sized pores. 730
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honeycomb films, the unbound avidin should have been removed by washing with buffer and albumin solutions. However, a small amount of avidin still remained on the control sample of non-biotinylated film. The remaining unbound avidin was mainly absorbed on the surface in between the pores rather than inside the pores, indicating that avidin located inside the pores on the biotinylated film was specifically absorbed by the biotin moieties. The second experiment was to observe the preferential allocation of the azido groups by clicking QDs to the film. CdSe/ZnS/PEG2000−NH2 QDs (diameter ∼ 9 nm; see the Supporting Information for a TEM image) were first functionalized with propiolic acid. The QDs were slightly aggregated after the reaction according to the TEM measurement. The CdSe/ZnS/PEG2000−NH2 QDs were also functionalized with NHS propiolate, followed by click reaction on the honeycomb film of 3-O-azidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose. However, there was no fluorescence observed with confocal microscopy. This may be due to luminescence inhibition resulting from the functionalization and the CuI-catalyzed click reaction.43 Hence, an alternative way was proposed to link QDs to the honeycomb films. NHS heptynoic ester was then synthesized and clicked to the honeycomb film of 3-O-azidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose, followed by reaction with CdSe/ZnS/ PEG2000−NH2 QDs. From this procedure, exposure of the QDs to the chemical reaction environment was minimized. Particularly, the exposure of the QDs to the CuI catalytic system was avoided. The click reaction of NHS heptynoic ester to the honeycomb film of 3-O-azidopropoxyEG13-2,6-di-Othexyldimethylsilyl cellulose was confirmed with ATR−FTIR as the azido peak at 2099 cm−1 decreased significantly, while the new carbonyl peak appeared at 1738 cm−1 (see the Supporting Information). The click reaction took place heterogeneously on the surface of the honeycomb film without stirring; therefore, it was expected to obtain a low conversion of the azido group. A full conversion was confirmed with FTIR spectroscopy by complete disappearance of the azido peak when conducting a homogeneous click reaction using the solvents of THF/DMSO (1:1). The QD-treated honeycomb film was characterized with confocal microscopy. A combined confocal fluorescent and optical image is displayed in Figure 7a, while Figure 7b shows a confocal fluorescent image only. It is clearly seen that the fluorescent QDs were preferentially located inside the pores. These results clearly show successful allocation of the functional azido groups inside the pores and provide a platform for site-specific functionalization of the honeycomb films for a
alkynated biotin was then clicked to the honeycomb film of 3O-azidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose. The ATR−FTIR spectra of the film before and after the click reaction are compared in Figure 5. Successful linkage of biotin
Figure 5. Comparison of ATR−FTIR spectra of the film of 3-OazidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose (a) before and (b) after the click reaction with the alkynated biotin.
to the film was confirmed by the decreasing azido peak (2096 cm−1), new carbonyl peak (1696 cm−1), and new triazole ring peaks (1655 and 1622 cm−1) after the click reaction. Because of the strong affinity between biotin and avidin, the film treated with fluorescent avidin was expected to show fluorescence at the spots where alkynated biotin was clicked, i.e., where azido group was located on the film. Figure 6 compares the combined
Figure 6. Comparison of the combined confocal fluorescent (pink) and optical (blue) images of the honeycomb films (a) with and (b) without biotin attachment.
confocal fluorescent and optical images of the honeycomb films with and without biotin attachment. The blue color represents the image from the optical microscope, showing the surface structure, while the pink color represents the image from the confocal microscope, showing the location of fluorescent avidin. With biotin attached, avidin was located inside the hexagonal pores (Figure 6a), while a small amount of avidin was absorbed only by the surface between the pores in the film without biotin treatment (Figure 6b). After avidin was applied to the
Figure 7. (a) Combined confocal fluorescent (blue) and optical (gray) image and (b) confocal fluorescent image of the QD-attached honeycomb film. 731
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variety of potential applications through simple “click chemistry”.
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CONCLUSION Amphiphilic cellulose derivative 3-O-azidopropoxypoly(ethylene glycol)-2,6-di-O-thexyldimethylsilyl cellulose of different DS from 0.07 to 0.47 and different DP from 4 to 22 of the hydrophilic side-chain PEG was synthesized and employed in the formation of honeycomb films via the “breath figures” method. The effect of DP and DS on the pore size was studied. Balanced amphiphilicity of polymers was found to be very important in the formation of uniform honeycomb films. For polymers with short PEG side chain (EG4), the pore size increased with an increasing DS. For polymers with long PEG side chain (EG22), the pore size decreased slightly with an increasing DS. For polymers with high DS (0.37−0.47), the pore size decreased with an increasing DP. For polymers with a similar number of hydrophilic EG segments, the pore size decreased with an increasing length of the pendant PEG. In addition, fluorescent avidin was attached to the biotinylated honeycomb film by means of biontin/avidin conjugation. Together with the experiment of attaching QDs to the film, preferential allocation of the functional azido group inside the pores was confirmed by confocal fluorescence microscopy.
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ASSOCIATED CONTENT
S Supporting Information *
Pore size distribution changing with the polymer concentration, ATR−FTIR spectra of the honeycomb film of 3-OazidopropoxyEG13-2,6-di-O-thexyldimethylsilyl cellulose before and after the click reaction with NHS heptynoic ester, and TEM image of the CdSe/ZnS/PEG2000−NH2 QDs. This material is available free of charge via the Internet at http:// pubs.acs.org.
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AUTHOR INFORMATION
Corresponding Author
*Telephone: (604) 827-5254. Fax: (604) 822-0661. E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS This work was financially supported by the Sentinel Bioactive Paper Network. Special thanks go to Kevin Hodgson, light microscopy technician at the University of British Columbia bioimaging facility, for assistance in the confocal microscopy measurements.
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