Deuterated Polymers for Probing Phase Separation Using Infrared

Article pubs.acs.org/Biomac

Deuterated Polymers for Probing Phase Separation Using Infrared Microspectroscopy Robert A. Russell,*,†,§ Tamim A. Darwish,† Ljiljana Puskar,‡ Danielle E. Martin,‡ Peter J. Holden,†,§ and L. John R. Foster§ †

National Deuteration Facility, Australian Nuclear Science & Technology Organisation, Lucas Heights, NSW Australia Infrared Microspectroscopy Beamline, Australian Synchrotron, Clayton, Victoria, Australia § Bio/Polymer Research Group, School of Biotechnology & Biomolecular Science, University of New South Wales, Sydney, NSW, Australia ‡

ABSTRACT: Infrared (IR) microspectroscopy has the capacity to determine the extent of phase separation in polymer blends. However, a major limitation in the use of this technique has been its reliance on overlapping peaks in the IR spectra to differentiate between polymers of similar chemical compositions in blends. The objective of this study was to evaluate the suitability of deuteration of one mixture component to separate infrared (IR) absorption bands and provide image contrast in phase separated materials. Deuteration of poly(3-hydroxyoctanoate) (PHO) was achieved via microbial biosynthesis using deuterated substrates, and the characteristic C−D stretching vibrations provided distinct signals completely separated from the C−H signals of protonated poly(3-hydroxybutyrate) (PHB). Phase separation was observed in 50:50 (% w/w) blends as domains up to 100 μm through the film cross sections, consistent with earlier reports of phase separation observed by scanning electron microscopy (SEM) of freeze-fractured protonated polymer blends. The presence of deuterated phases throughout the film suggests there is some miscibility at smaller length scales, which increased with increasing PHB content. These investigations indicate that biodeuteration combined with IR microspectroscopy represents a useful tool for mapping the phase behavior of polymer blends.

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INTRODUCTION The blending of polymers and the production of composite materials to tailor physical characteristics, biocompatibility, and biodegradability to suit specific applications necessarily requires the ability to determine the mixing or distribution of the component materials. A cogent case in point is the desirability of modifying materials produced from polyhydroxyalkanoates (PHAs) to optimize their use as biocompatible materials in biomedical applications. PHAs are a family of natural biopolyesters produced by a wide variety of microorganisms as intracellular storage compounds of carbon and energy under imbalanced growth conditions.1 Poly(3-hydroxybutyrate) (PHB) is the most commonly reported member of the PHA family and can be produced in significant quantities through bioprocessing using bacteria such as Ralstonia eutropha and transgenic Escherichia coli with an excess of glucose (Figure 1a).2,3 Modification of the bioprocessing parameters, in particular the use of more complex organic substrates and different species, has produced over 150 different monomer units resulting in a broad range of polymer properties.4 The PHA synthase enzymes of different microbial species have substrate specificity for either short chain (R)-3-hydroxy fatty acids comprising 3 to 5 carbon atoms or 6 to 14 carbon atoms, which are polymerized into short chain length (scl-) or medium chain length (mcl-) PHAs, respectively. PHB is classified as a scl-PHA, while microbial metabolism of octanoic (C8) fatty © 2013 American Chemical Society

Figure 1. Monomer structures of (a) PHB and (b) PHO.

acid in Pseudomonas oleovorans grown under oxygen limitation produces a fairly amorphous, flexible polymer that is predominantly poly(3-hydroxyoctanoate) (PHO), a mcl-PHA (Figure 1b).5 PHAs are an attractive alternative to synthetic polymers due to their biocompatibility and biodegradability. They are particularly suitable for short-lived medical implants without harmful breakdown products and have been investigated for use in wound repair as surgical sutures and nerve scaffolds.6,7 The monomeric component of PHB is the stable ketone body 3Received: November 19, 2013 Revised: December 12, 2013 Published: December 23, 2013 644

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respectively, using IR microspectroscopy to differentiate between hydrogenated and deuterated phases.

hydroxybutyric acid (3HBA), a normal constituent of human blood and is chemically identical in microbial and mammalian systems.8,9 However, applications of PHB are limited by its brittleness and relatively high melting temperature close to decomposition temperature.10 Consequently, the material properties of PHB have been modulated by blending with other biopolymers including mcl-PHAs.11−16 Blending of PHB with PHO has received relatively little attention because the polymers have been reported as completely immiscible, and the partially crystalline nature of the two components has made determination of miscibility difficult.17 Phase separation between PHB and PHO, reported by Dufresne and Vincendon, was characterized using SEM and thermal/mechanical analysis; however, the glass transition (Tg) data thus obtained was influenced by the crystalline phases. The freeze-fractured surface of a 50:50 (% w/w) PHO/PHB blend displayed a cocontinuous network, indicating phase separation of domains ca. 10 μm but with many voids in between. FT-IR microscopy has been used to characterize phase separation in blends of immiscible polymers, e.g., PHB with poly(L-lactic acid) and poly(ε-caprolactone).18 This study by Vogel et al. relied on the IR shift of the carbonyl peak associated with changes in crystallinity. More recently, PHB and cellulose acetate butyrate (CAB) blends were studied by IR and NIR imaging spectroscopy,19 where changes in the spectral regions associated with the first and second overtones of the CO stretching vibrations were used to characterize crystallization. Differences in FT-IR absorbance wavenumbers of the ester band were also used by Randriamahefa et al. in an attempt to distinguish between PHB and PHO.20 However, the difference in crystallinity exhibited by PHB and PHO gave similar CO stretching peaks at 1724 and 1742 cm−1, respectively. These studies highlight the difficulty in distinguishing between the spectra of these chemically similar polymers. Crystallinity affects the CO peak wavenumber and peaks attributed to stretching vibrations of amorphous and crystalline carbonyl groups occur in both polymers. Consequently, current techniques are not reliable for distinguishing between these polymers in a blend. Similarly, reliance on peak differences in the fingerprint region (ca. 800−1400 cm−1) is also problematic due to the interferences and the crowdedness of the signals. Furthermore, the position of IR signals of functional groups with polarized bonds, such as CO or C−O, significantly depends on matrix and intermolecular interactions. The bond’s force constants of such groups change unpredictably due to their susceptibility to hydrogen bonding, which results in shifts in their frequency position.21 Thus, assignment of such signals, in an attempt to map the phase separation of a polymer blend, is complicated and prone to inaccuracy. Here we report an accurate alternative to previously reported techniques for mapping phase separation, through the application of deuteration. A deuterated polymer phase can be clearly differentiated from a nondeuterated polymer based on nonoverlapping IR stretching vibrations of their carbon− hydrogen and carbon−deuterium alkyl chains (C−D stretch 2000−2250 cm−1; C−H stretch, 2800−3050 cm−1). This technique avoids the problems of overlapping peaks as well as the unpredictable shifts in IR signals due to intermolecular interactions and is independent of crystalline properties of polymers. In this study we demonstrate this technique by characterizing the phase separation between immiscible mixtures of scl- and mcl-PHAs and PHB and PHO,

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MATERIALS AND METHODS

Material and Reagents. Deuterated octanoic acid (octanoic acidd15, 98% D by 1H NMR) was produced by catalyzed 1H/2H exchange under hydrothermal conditions (modification of Hughes et al.)22 and used as the precursor for microbial biosynthesis of deuterated PHO (dPHO) polymer. Heavy water (D2O) used in the growth medium for dPHO production was obtained from Sigma-Aldrich, >99.8% D isotopic purity. CHCl3 used for polymer purification and blending was obtained from Ajax FineChem (Australia). Octanoic acid-d15 added during oxygen-limited cultivation of Pseudomonas oleovorans (ATCC 29347) was metabolized to form a terpolymer of C6, C8, and C10 monomer units within intracellular inclusion bodies as previously reported.5 Polymer Extraction and Film Blend Preparation. Cell biomass was lyophilized, and the deuterated biopolymer was extracted using CHCl3. Culture was centrifuged, and the frozen cell pellet was lyophilized prior to solvent extraction in 10× volume of CHCl3 (60 °C, stirring, 12 h). The crude polymer fraction was concentrated by rotary evaporation, and purification of dPHO from the crude extract was achieved by precipitation of the polymer following the addition of cold methanol. The polymer was redissolved in CHCl3 and the precipitation repeated. PHB (natural origin, Sigma-Aldrich) was dissolved by stirring in CHCl3 at 60 °C. Polymer solutions were poured into Teflon-lined Petri dishes (Welch Fluorocarbon Inc., USA) to allow solvent evaporation, then placed under vacuum (−100 kPa) at room temperature in a vacuum oven (Emtivac Engineering, Dandenong South, Australia) to remove residual solvent. Molecular weights of dPHO and PHB (149 and 522 kDa, respectively) were determined by gel permeation chromatography (Polymer Laboratories GPC50, Agilent, USA) against a series of narrow molecular weight polystyrene standards. Samples of dPHO and PHB films were weighed into glass ca. 20 mm diameter vials and dissolved in CHCl3, stirring at 60 °C. Solvent was then evaporated to yield polymer films of different blend compositions covering the base of the vials. When removed from vials, these films measured 50−100 μm thick using digital calipers (Starret model 727, Jedburgh, Scotland). Films were left to equilibrate in ziplock bags for one week prior to microtoming and infrared mapping. To better understand the polymer phase behavior through the bulk of the films, cross-sections of films were microtomed to 5 μm thin sections and pressed flat onto 0.5 mm thick CaF2 discs (Crystran Limited, Poole, U.K.). NMR Spectroscopy. Protonated and perdeuterated PHO dissolved in CDCl3 (Sigma-Aldrich) were analyzed by 1H and 2H NMR (Bruker 400 MHz spectrometer at 298 K in CDCl3) to identify moieties in the PHO monomer and to determine the extent of deuterium substitution in the polymer resulting from biosynthesis using deuterated substrates. FT-IR Spectroscopy. An ATR spectrometer (Nicolet iS10 with diamond ATR accessory, Thermo Scientific) was used to characterize the differences between the FT-IR spectra of protonated (PHB) and deuterated (dPHO) components. Sixteen scans were coadded, and the background was subtracted, resolution 4 cm−1. Infrared Microspectroscopy. Film thin sections were examined in transmission mode using a Bruker FT-IR spectrometer attached to a Hyperion 3000 IR microscope equipped with a 64 × 64 pixel focal plane array (FPA) imaging detector. With the microscope reflective IR Cassegrain objective and condenser (15×, 0.4 N.A.) each detector field of view corresponded to ca. 170 μm2 on the sample. Larger sample areas were investigated by compiling several of these frames. All data were recorded using Bruker OPUS version 6.5 software, and data analysis was performed using Bruker OPUS 7.0 imaging software.

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RESULTS AND DISCUSSION NMR Characterization. We have previously reported the partial deuteration of PHO, whereby the combination of NMR 645

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Figure 2. 1H NMR spectra of PHO produced from (a) protonated substrates; (b) 2H NMR spectra of dPHO showing the corresponding deuterium signals; (c) deuterated substrates showing proton residues. Peaks indicate proton moieties shown in the PHO monomer.

and GC/MS showed the substitution of all but one proton by deuterium via biosynthesis.5 Figure 2 shows (a) the 1H NMR spectra of a protonated version of PHO and (b) 2H NMR of dPHO showing the expected chemical shift of the deuterium signals, and Figure 2c shows the proton residues in the dPHO produced in this study. Deuteration was confirmed at greater than 95% D by NMR, reflecting the isotopic purity of substrates used in microbial polymer production. It is noteworthy that the deuterium signals of the CD2 and CD moieties next to the polymer backbone are broad in nature (Figure 2b) and do not show the typical splitting pattern of peaks in the protonated sample. This is unlike the terminal CD3 groups, which showed a relatively sharp deuterium signal due to free and unrestricted rotations. FT-IR Characterization. The FT-IR spectrum of dPHO shows strong peaks at 2050 to 2200 cm−1, which were attributed to C−D stretching vibrations (Figure 3). The peaks observed around 2800 cm−1 indicated the presence of some C− H stretching vibrational signal from the relatively small protonated component of substrates that were biosynthesized into the final product. The C−D component of dPHO derived from the integrated peak areas was 0.72 of the total. In contrast, the spectrum for PHB shows only the C−H stretching vibrations from 2800 to 3050 cm−1, which would indicate an absence of deuterated polymer in a particular region of a blend. Thus, we propose that the FT-IR spectrum of areas within a well-mixed blend of PHB and dPHO would show consistent signals in both C−H and C−D peaks, while areas that show predominantly C−H or C−D signatures would be an indication of phase separation of the two polymers. Mapping Deuterated/Protonated Phases in Film Sections. Cross sections of film blends were pressed flat onto CaF2 discs for IR microspectroscopy in transmission mode. This approach has the additional advantage of characterizing polymer miscibility throughout the bulk of the film rather than the phase separation at the solution/air interface as reported in previous IR microspectroscopy studies.18,23 In the study here, optical images of analyzed areas are overlain with contour maps where red and blue represent high and low absorbances for that wavenumber range, respectively. The cross section of a PHB/dPHO blend (50:50% w/w) is shown in Figure 4. Phase separation in the analyzed area cannot

Figure 3. FT-IR spectra of (a) PHB and (b) dPHO.

be assessed using optical microscopy (Figure 4a), as the two polymers do not show distinct areas that are often observed in chemically diverse polymer blends.4 In contrast, the integrated areas associated with the C−D and C−H stretching vibrations show distinct regions of localized intensity. The C−D map (Figure 4c, contour plot of peak 2000 to 2250 cm−1) shows that dPHO is distributed in irregular domains ca. 50 μm throughout the film. Areas of low C−D absorbance (dark blue) indicate a lower concentration of dPHO, which corresponds to areas of predominantly PHB (high absorbance in the C−H map, Figure 4b). Comparatively less C−H absorbance was observed in the corresponding areas of dPHO, due to the relatively strong C−H signal resulting from substrate purity described above. A persistent low level of dPHO was observed across most of the section (intermediate absorbance in C−D map) and the 646

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Figure 4. PHB to dPHO 50:50 (% w/w) film cross-section showing (a) optical image with no discernible difference between polymers; IR contour map of (b) C−H (2800−3050 cm−1) and (c) C−D (2000− 2250 cm−1). Color bar shows normalized intensity. Figure 5. PHB to dPHO 60:40 (% w/w) film cross-section showing (a) optical image; IR contour map of (b) C−H (2800−3050 cm−1) and (c) C−D (2000−2250 cm−1).

boundaries between PHB-rich and dPHO-rich areas overlapped to some extent. Analysis of extracted spectra from areas of lowest intensity in the C−D contour map showed the minimum ratio of the C−D component in the polymer blend was 0.15, suggesting some association of PHB and dPHO at smaller length scales. By controlling film thickness to less than the size of most domain sizes, the likelihood of FT-IR transmission through overlapping domains of the film section was minimized. Thus, the chemical information from FT-IR microspectroscopy, coupled with the use of a deuterated polymer probe, provides a suitable technique to determine miscibility in polymer blends. The distribution of C-D rich areas was also observed in the blend comprising a ratio of 60:40 (% w/w) PHB to dPHO (Figure 5c). As the deuterated polymer component is lower, these domains become fewer and protonated matrix becomes more continuous through the film (Figure 5b). Compared to the 50:50% w/w blend, there appears to be a greater degree of overlap between C−D and C−H regions, indicating a greater degree of miscibility in this blend. As the deuterated polymer component is reduced further to 20% in the blend (80:20 (% w/w) PHB to dPHO), increased miscibility was observed (Figure 6). There was minimal perturbation of the PHB matrix through the thickness of the films (Figure 6b), and the regions of low intensity for both

polymer phases were only observed at the edges of the section in both maps, due to edge effects of IR interferograms. In order to determine whether the deuteration had any influence on the phase separation observed in these blends, a control sample of 50:50 (% w/w) dPHO/PHO was also examined. As this material was difficult to microtome, a solution of polymer in chloroform was drop cast directly onto a CaF2 slide, and the film was analyzed following solvent evaporation. Segregation of isotopic species in binary blends has been reported;24 however, under the conditions of this experiment, deuteration had no observable effect. Figure 7 exhibits no domains of protonated or deuterated phases, rather a steady drop in relative intensity of both polymer species, which reflected the decreasing thickness toward the edge of the film, indicating that the difference in bond energy between C− H and C−D did not affect the distribution of the polymer chains as observed by IR microspectroscopy. This agrees with previous work describing the comparable physicochemical properties of protonated and deuterated forms of a polymer.5 Thus, this technique is sensitive to the phase separation between polymers, even those that are chemically very similar, such as PHB and PHO, that may be difficult to distinguish using existing techniques. 647

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polymers consistent with that reported previously from a freeze-fractured surface of a PHB/PHO blend. The chemical information provided an indication of partial miscibility at smaller length scales not previously identified and showed a trend of increasing miscibility with decreasing PHO content. Our method is independent of crystallinity and thus can be applied to a variety of polymer blends where overlapping peaks in the IR spectrum would otherwise be difficult to distinguish. The separation of C−H and C−D peaks in the IR spectrum also makes possible the determination of a deuterated phase in a complex polymer mix. In addition to the biodeuterated polymer presented here, deuteration can be readily applied to chemically synthesized polymers or small molecules and is therefore suitable for probing miscibility of a range of polymer or small molecule blends.

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AUTHOR INFORMATION

Corresponding Author

*(R.A.R.) E-mail: [email protected]. Tel: +61(2)9717-3691. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This IR analyses were undertaken on the Infrared beamline and associated instruments at the Australian Synchrotron, Victoria, Australia. Deuteration and preparation of films was undertaken at the National Deuteration Facility, which was partly funded by the National Collaborative Research Infrastructure Strategy, an initiative of the Australian Government.

Figure 6. PHB to dPHO 80:20 (% w/w) film cross-section showing (a) optical image; IR contour map of (b) C−H (2800−3050 cm−1) and (c) C−D (2000−2250 cm−1).

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ABBREVIATIONS FT-IR, Fourier transform infrared; NMR, nuclear magnetic resonance spectroscopy; ATR, attenuated total reflection spectroscopy

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REFERENCES

(1) Dawes, E. A.; Senior, P. J. The role and regulation of energy reserve polymers in micro-organisms. Adv. Microb. Physiol. 1973, 10, 135−266. (2) Lenz, R. W.; Marchessault, R. H. Bacterial polyesters: Biosynthesis, biodegradable plastics and biotechnology. Biomacromolecules 2005, 6, 1−8. (3) Park, S. J.; Choi, J.; Lee, S. Y. Engineering of Escherichia coli fatty acid metabolism for the production of polyhydroxyalkanoates. Enzyme Microb. Technol. 2005, 36, 579−588. (4) Rehm, B. Polyester synthases: natural catalysts for plastics. Biochem. J. 2003, 376 (1), 15−33. (5) Foster, L. J. R.; Russell, R. A.; Sanguanchaipaiwong, V.; Stone, D. J. M.; Hook, J. M.; Holden, P. J. Biosynthesis and characterization of deuterated Polyhydroxyoctanoate. Biomacromolecules 2006, 7 (4), 1344−1349. (6) Shishatskaya, E. I.; Volova, T. G.; Puzyr, A. P.; Mogilnaya, O. A.; Efremov, S. N. Tissue response to the implantation of biodegradable polyhydroxyalkanoate sutures. J. Mater. Sci.: Mater. Med. 2004, 15, 719−28. (7) Mosahebi, A.; Fuller, P.; Wiberg, M.; Terenghi, G. Effect of allogeneic Schwann cell transplantation on peripheral nerve regeneration. Exp. Neurol. 2002, 173, 213−23. (8) Zinn, M.; Witholt, B.; Egli, T. Occurrence, synthesis and medical application of bacterial polyhydroxyalkanoate. Adv. Drug Delivery Rev. 2001, 53 (1), 5−21. (9) Foster, L. J. R.; Tighe, B. J. Enzymatic assay of hydroxybutyric acid monomer formation in poly(beta-hydroxybutyrate) degradation studies. Biomaterials 1995, 16, 341−343.

Figure 7. C−D contour map of 50:50 (% w/w) dPHO/PHO blend. The distribution of the deuterated polymer shows no localized intensity and a steady drop in intensity toward the thin edge of the film (lower right).

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CONCLUSIONS Reports in the literature using IR microspectroscopy to investigate phase separation in biopolymers have relied on shifts of the carbonyl peak due to chemically different species and have been confined to mapping the surface of solvent cast films. Limitations on characterizing chemically similar polymers such as PHB and PHO include subtle shifts and impact of crystallinity on the carbonyl peak and peak differentiation in the crowded fingerprint region. The proposed technique using deuteration to differentiate between polymers using IR microspectroscopy demonstrated phase separation of these 648

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(10) Kunioka, M.; Doi, Y. Thermal degradation of microbial copolyesters: poly(3-hydroxybutyrate-co-3-hydroxyvalerate) and poly(3-hydroxybutyrate-co-4-hydroxybutyrate). Macromolecules 1990, 23, 1933−1936. (11) Wang, W.; Zhu, X.; Wang, X.; Chen, G.-Q.; Xu, K. Thermal degradation of microbial copolyesters: poly(3-hydroxybutyrate-co-3hydroxyvalerate) and poly(3-hydroxybutyrate-co-4-hydroxybutyrate). J. Appl. Polym. Sci. 2008, 107, 166−173. (12) Zhao, K.; Deng, Y.; Chun, J.; Chen, G. Q. Polyhydroxyalkanoate (PHA) scaffolds with good mechanical properties and biocompatibility. Biomaterials 2003, 24, 1041−1045. (13) Furukawa, T.; Sato, H.; Murakami, R.; Zhang, J.; Duan, Y.-X.; Noda, I.; Ochiai, S.; Ozaki, Y. Structure, dispersibility, and crystallinity of poly(hydroxybutyrate)/poly(L-lactic acid) blends studied by FT-IR microspectroscopy and differential scanning calorimetry. Macromolecules 2005, 38, 6445−6454. (14) Sodian, R.; Hoerstrup, S. P.; Sperling, J. S.; Martin, D. P.; Daebritz, S.; Mayer, J. E., Jr.; Vacanti, J. P. Evaluation of biodegradable, three-dimensional matrices for tissue engineering of heart valves. ASAIO J. 2000, 46 (1), 107−110. (15) Sodian, R.; Hoerstrup, S. P.; Sperling, J. S.; Daebritz, S.; Martin, D. P.; Moran, A. M.; Kim, B. S.; Schoen, F. J.; Vacanti, J. P.; Mayer, J. E., Jr. Early in vivo experience with tissue-engineered trileaflet heart valves. Circulation 2000, 102 (19, Suppl 3), 22−29. (16) Furrer, P.; Panke, S.; Zinn, M. Quantitative analysis of bacterial medium-chain-length poly([R]-3-hydroxyalkanoates) by gas chromatography. J. Microbiol. Methods 2007, 69, 206−213. (17) Dufresne, A.; Vincendon, M. Poly(3-hydroxybutyrate) and poly(3-hydroxyoctanoate) blends: Morphology and mechanical behavior. Macromolecules 2000, 33 (8), 2998−3008. (18) Vogel, C.; Wessel, E.; Siesler, H. W. FT-IR imaging spectroscopy of phase separation in blends of poly(3-hydroxybutyrate) with poly(L-lactic acid) and poly(e-caprolactone). Biomacromolecules 2008, 9 (2), 523−527. (19) Suttiwijitpukdee, N.; Sato, H.; Unger, M.; Ozaki, Y. Effects of hydrogen bond intermolecular interactions on the crystal spherulite of poly(3-hydroxybutyrate) and cellulose acetate butyrate blends: Studied by FT-IR and FT-NIR imaging spectroscopy. Macromolecules 2012, 45 (6), 2738−2748. (20) Randriamahefa, S.; Renard, E.; Guérin, P.; Langlois, V. Fourier transform infrared spectroscopy for screening and quantifying production of PHAs by Pseudomonas grown on sodium octanoate. Biomacromolecules 2003, 4 (4), 1092−1097. (21) Li, G.; Ye, S.; Morita, S.; Nishida, T.; Osawa, M. Hydrogen bonding on the surface of poly(2-methoxyethyl acrylate). J. Am. Chem. Soc. 2004, 126 (39), 12198−12199. (22) Hughes, J. R.; Luckhurst, G. R.; Praefcke, K.; Singer, D.; Tearle, W. M. Chemically-induced discotic liquid crystals. Structural studies with NMR spectroscopy. Mol. Cryst. Liq. Cryst. 2003, 396 (1), 187− 225. (23) Ling, S.; Qi, Z.; Knight, D. P.; Shao, Z.; Chen, X. FTIR imaging, a useful method for studying the compatibility of silk fibroin-based polymer blends. Polym. Chem. 2013, 4, 5401−5406. (24) Müller, G.; Schwahn, D.; Eckerlebe, H.; Rieger, J.; Springer, T. Effect of the Onsager coefficient and internal relaxation modes on spinodal decomposition in the high molecular isotopic blend polystyrene/deutero-polystyrene studied with small angle neutron scattering. J. Chem. Phys. 1996, 104, 5326−5337.

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