Cellulose Microfibril Formation by SurfaceTethered Cellulose Synthase Enzymes Snehasish Basu,† Okako Omadjela,‡ David Gaddes,§ Srinivas Tadigadapa,§ Jochen Zimmer,‡ and Jeffrey M. Catchmark*,† †

Department of Agricultural and Biological Engineering, Pennsylvania State University, University Park, Pennsylvania 16802, United States ‡ Center for Membrane Biology, Department of Molecular Physiology and Biological Physics, University of Virginia, Charlottesville, Virginia 22908, United States § Department of Electrical Engineering, Pennsylvania State University, University Park, Pennsylvania 16802, United States S Supporting Information *

ABSTRACT: Cellulose microfibrils are pseudocrystalline arrays of cellulose chains that are synthesized by cellulose synthases. The enzymes are organized into large membraneembedded complexes in which each enzyme likely synthesizes and secretes a β-(1→4) glucan. The relationship between the organization of the enzymes in these complexes and cellulose crystallization has not been explored. To better understand this relationship, we used atomic force microscopy to visualize cellulose microfibril formation from nickel-film-immobilized bacterial cellulose synthase enzymes (BcsA-Bs), which in standard solution only form amorphous cellulose from monomeric BcsA-B complexes. Fourier transform infrared spectroscopy and X-ray diffraction techniques show that surface-tethered BcsA-Bs synthesize highly crystalline cellulose II in the presence of UDPGlc, the allosteric activator cyclic-di-GMP, as well as magnesium. The cellulose II cross section/diameter and the crystal size and crystallinity depend on the surface density of tethered enzymes as well as the overall concentration of substrates. Our results provide the correlation between cellulose microfibril formation and the spatial organization of cellulose synthases. KEYWORDS: cellulose biosynthesis, BcsA-B cellulose synthase, surface immobilization, cellulose microfibril synthesis and assembly, crystal size, crystallinity, allomorph

C

Gluconacetobacter, Agrobacterium, Pseudomonas, Rhodobacter, and Salmonella) and Gram-positive (such as Sarcina ventriculi) species can synthesize cellulose.7−9 Among the different bacteria, Gluconacetobacter hansenii (formerly Acetobacter xylinum) is a very well known bacterial strain for cellulose production. It has been used as a model micro-organism for basic and applied studies on cellulose due to its ability to produce highly crystalline cellulose fibrils.10,11 Other celluloseproducing bacteria can produce amorphous cellulose as a biofilm component.12,13 For many years, in vitro cellulose biosynthesis from purified components had not been possible due to difficulties in purifying catalytically active CSs. However, the recent expression, purification, and functional characterization of the bacterial cellulose synthase BcsA-B from Rhodobacter sphaer-

ellulose is the most abundant biopolymer on Earth and also commercially the most widely used biological material. Cellulose is used in the production of fuel, food, paper, packaging, numerous fiber composites, biomedical materials, cosmetics, and plastics, in addition to having many other industrial uses.1−3 Although cellulose has been studied for over 150 years, the mechanism of cellulose biosynthesis is not fully understood. Specifically, many fundamental questions remain regarding the structure and composition of the cellulose synthase complex (CSC), which contains multiple cellulose synthase enzymes, and its relationship to the structure of cellulose fibrils, including the formation of different crystalline allomorphs.4,5 Cellulose is a linear chain of several hundreds to many thousands of β-(1→4)-linked D-glucose units that, in most species, are linearly arranged into fiber-like structures, called cellulose micro- and macrofibrils. In addition to plants, bacteria, algae, and fungi, even animal species can synthesize cellulose.6 In the bacterial kingdom, both Gram-negative (including © XXXX American Chemical Society

Received: September 8, 2015 Accepted: January 22, 2016

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Figure 1. BcsA-B cellulose synthase requires UDP-Glc, c-di-GMP, and Mg2+ ions to synthesize cellulose in in vitro conditions. (a) Blank Ni film. (b) Only BcsA-B enzymes. (c) BcsA-B enzymes + UDP-Glc. (d) BcsA-B enzymes + UDP-Glc + Mg2+. (e) BcsA-B enzymes + UDP-Glc + c-di-GMP. (f) BcsA-B enzymes + UDP-Glc + c-di-GMP + Mg2+ + EDTA. AFM image scan size = 2 μm × 2 μm.

oides provide access to a catalytically active, heterodimeric CS. The crystal structure of BcsA-B has also been obtained and consists of a single BcsA-B complex (≈164 kDa), where BcsA is the catalytic subunit containing a cytosolic glycosyltransferase (GT) and a membrane-embedded, channel-forming transmembrane domain.14 The GT domain is expected to bind a single UDP (uridine diphosphate)-activated glucose molecule (donor) as well as the nonreducing end of the growing cellulose chain (acceptor).14,15 Bacterial cellulose biosynthesis is also allosterically activated by c-di-GMP (cyclic diguanylate), a bacterial-signaling molecule implicated in biofilm formation.15−18 Catalytic activity of BcsA requires the presence of the BcsB subunit as well as a divalent cation, either magnesium or manganese.15 Next to BcsA-B, bacterial cellulose synthase complexes likely contain a third BcsC subunit, which is predicted to form an outer membrane porin preceded by a large periplasmic domain. BcsC has been shown to be essential for cellulose biosynthesis in vivo but not in vitro.8,15,19 In addition, bacteria that form cellulose microfibrils usually contain a fourth subunit (BcsD), and bcsD knockout strains produce cellulose of reduced crystallinity.19,20 The biosynthesis of crystalline cellulose I requires an intact CSC in plants, bacteria, and algae.7,21−24 In vitro, any alteration/disruption of the native arrangement of CSC results in noncrystalline/amorphous cellulose or cellulose II.19,25,26 Therefore, crystallinity of the cellulose microfibril is dependent on the spatial arrangement of the CSs within the CSC. Cellulose microfibril structure, including characteristics such as size, shape, allomorph, and crystallinity, is believed to be determined by a complex interaction between its biosynthesis by cellulose synthase enzymes and any associated proteins and the properties of cellulose itself, including how cellulose selfassociates through noncovalent interactions such as hydrogen bonding, van der Waals forces, and hydrophobic interactions.5 In vivo studies including the impact of genetic manipulation on cellulose structure are limited by the overall complexity of such

systems and unknowns such as the composition and structure of cellulose synthase complexes. In vitro studies on membrane fractions share these difficulties and introduce additional complexities as the extent to which the cellulose synthesis biomachinery has been preserved or altered during purification. The recent availability of purified functional cellulose synthase enzymes has enabled an entirely new approach for studying the relationship between the architecture of cellulose synthase organization and the structure of cellulose produced. In this study, active cellulose synthase enzymes are immobilized and clustered onto surfaces, and the microfibrils which form under different substrate and immobilization conditions are comprehensively examined. The crystal structure of the heterologously expressed and purified BcsA-B enzyme shows that a monomeric BcsA-B complex is sufficient to synthesize a single glucan chain and also translocates it across the plasma membrane through a pore formed by BcsA.14 Thus, to test the hypothesis that cellulose microfibrils arise due to the close proximity of CSs in CSCs, we investigated the effects of surface immobilization of BcsA-B on cellulose allomorph, crystal size, crystallinity, and microfibril formation. To this end, we compared in vitro synthesized celluloses produced from BcsA-B enzymes bound to a nickel surface or in solution. Cellulose was synthesized under different conditions, including enzyme surface concentration (packing density), substrate (UDP-Glc), and cofactors (c-di-GMP, Mg2+), as a function of temperature and time. This work is a first step toward the foundation for the design and construction of artificial systems for quantitatively understanding the relationship between the assembly of cellulose biomachinery and microfibril synthesis. It also has broader impacts on possibilities for producing new biopolymer nanomaterials using engineered systems.

RESULTS Role of Substrate and Cofactors in Cellulose Microfibril Formation. It has previously been shown that the B

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Figure 2. Time-scale synthesis of cellulose microfibrils: (a) 1 min, (b) 2 min, (c) 3 min, (d) 4 min, (e) 5 min, (f) 10 min, (g) 30 min, and (h) 60 min. AFM image scan size = 2 μm × 2 μm.

first cellulose microfibrils were observed after 3 min (Figure 2c), which suggests that the polymerization was started after 2 min of incubation, or perhaps, cellulose was being synthesized but was not observed due to lack of aggregation. The density, cross section, and length of the cellulose microfibrils increased until 60 min (Figure 2c−h). The height of the microfibril measured from the Ni surface was taken to be equivalent to the cross section/diameter to avoid any AFM tip broadening effect. The maximum cross section ranged from 7 to 10 nm at 60 min, and the length of the microfibrils varied from 300 to 1000 nm after a 60 min polymerization reaction (Figure 2h). The branching of microfibrils observed may be a pattern formed simply by the aggregation of glucan chains and microfibrils from immobilized enzymes located at different positions across the surface. The AFM scanning of cellulose microfibrils has also been done under wet conditions after 4 min of synthesis. Throughout scanning, the sample was immersed in sterile filtered DI water (Figure S1). The image contains plenty of shadows due to the movement of the cellulose microfibrils, and the average cross section is 7 ± 2 nm, which is 1.5 nm (5.5 ± 1.5 nm) higher than that in the dry state (Figure S1). Due to the movement of the cellulose microfibril in wet conditions, AFM scanning was unable to image proper morphology of the cellulose fiber, thus all other images were taken under dry conditions. Effect of Temperature on Cellulose Polymerization and Microfibril Morphology. The effect of the temperature on cellulose microfibril synthesis/polymerization was studied at different temperatures (25, 37, and 50 °C) over a total of 60 min reaction time. At 25 °C, the cellulose production was slow, and only a small number of cellulose microfibrils were formed (Figure S2a). The length and cross section were 30−500 and 4−9 nm, respectively (Figure S2a). However, at 37 °C, a higher apparent polymerization rate was observed with higher cross section (7−10 nm) and longer length (300−1000 nm) of

formation of cellulose by BcsA-B requires the presence of UDPGlc, c-di-GMP, and a divalent cation such as Mg2+ or Mn2+.14−18,27−30 However, the impact of BcsA-B surface immobilization on the nature and morphology of the synthesized cellulose has not been explored to date. Therefore, we immobilized polyhistidine-tagged BcsA-B on Ni surfaces at different surface densities and initiated cellulose biosynthesis upon addition of UDP-Glc, c-di-GMP, and Mg2+ and imaged product formation by atomic force microscopy (AFM). In the absence of UDP-Glc, c-di-GMP, and Mg2+, no cellulose microfibrils were observed, resembling a smooth surface of blank Ni film (Figure 1a,b). No product was also formed in the presence of UDP-Glc only (Figure 1c). However, in the presence of UDP-Glc and Mg2+, small discrete celluloses (cross section = 4−8 nm; length = 15−230 nm) were observed (Figure 1d), suggesting that in a detergent-solubilized state, BcsA-B retains some catalytic activity in the absence of c-diGMP, as previously described.15 In the presence of UDP-Glc and the allosteric activator c-di-GMP (but no Mg2+), the immobilized BcsA-B synthesizes very short (length = 30−350 nm) and thin (cross section = 4−7 nm) cellulose microfibrils (Figure 1e), perhaps due to the presence of residual Mg2+ ions in the buffer. Upon addition of EDTA, however, no cellulose is formed in the presence of UDP-Glc and c-di-GMP only (Figure 1f), suggesting the indispensability of the Mg2+ ions for cellulose formation. Therefore, both cofactors (c-di-GMP and Mg 2+ ) are required for catalytic activity of BcsA-B. Furthermore, and as described below, in the presence of required substrate (UDP-Glc) and all cofactors (c-di-GMP and Mg2+), BcsA-B efficiently synthesizes cellulose microfibrils. Time Dependence of Microfibril Formation. To determine the microfibril polymerization rate, cellulose microfibril synthesis was studied as a function of time (1−60 min). No cellulose microfibrils were observed until 2 min after synthesis was initiated in the presence of 25 μg of immobilized BcsA-B, UDP-Glc, c-di-GMP, and Mg2+ ions (Figure 2a,b). The C

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ACS Nano microfibrils (Figure 2h), consistent with BcsA-B showing a maximum activity at this temperature.15 Cellulose microfibril formation was also observed at 50 °C, which suggests that the BcsA-B retains some activity at higher temperatures and can synthesize cellulose. However, the microfibril morphology was completely changed at higher temperature and exhibited a wider cross section (22−33 nm) (Figure S2b), suggesting that the compact microfibril structure is compromised under these conditions, possibly due to changes in enzyme activity (resulting in a change in the rate of glucan chain polymerization), the overall energetics of the system, and the impact these factors have on hydrogen bonding or van der Waals interactions, which drive cellulose assembly. Effect of Endo- and Exoglucanase on BcsA-BProduced Cellulose Microfibril. An endo- and exoglucanase was used to analyze the degree of crystallinity of the BcsA-Bproduced cellulose microfibrils. Earlier studies showed that BcsZ, a periplasmic endo-β-1,4-glucanase encoded within the bacterial celluose synthase gene operon, efficiently degrades amorphous glucan chains, yet the enzyme has very low activity toward crystalline cellulose microfibrils.15,31−33 Performing cellulose biosynthesis reactions with immobilized BcsA-B in the presence of BcsZ greatly reduced the cellulose microfibrils formed, consistent with BcsZ degrading the nascent glucan chains as they emerge from the BcsA-B complex15 (Figure S3a). However, adding BcsZ after completion of cellulose biosynthesis did not degrade the cellulose microfibrils formed, consistent with BcsZ being less active toward aggregated celluloses (Figure S3b). These results are consistent with the BcsA-B-synthesized cellulose microfibrils being highly ordered or crystalline in nature. In addition to the endoglucanase (BcsZ) treatment, the same experiments have also been performed in the presence of 0.1 mg/mL exoglucanase/Cel7A (Trichoderma reesei). However, Cel7A efficiently degrades both nascent glucan chains as well as crystalline microfibrils (Figure S3c,d). Unlike BcsZ, Cel7A could easily degrade the crystalline microfibril, likely due to the presence of a carbohydrate binding domain specific to crystalline microfibrils.34 Impact of Enzyme Concentration, Substrate, and Cofactors on Cellulose Microfibril Morphology. AFM Analysis. Microfibril formation was studied by using increasing amounts of BcsA-B (25 → 100 → 200 → 400 μg) in solution during immobilization on the Ni film, expected to result in different surface densities of the immobilized complex. The substrate (5 mM UDP-Glc) and cofactor concentration (30 μM c-di-GMP; 20 mM MgCl2) were kept constant during cellulose production. The AFM analysis shows that the microfibril cross section decreases from 8.5 ± 1.5 to 5.75 ± 0.75 nm (32%) by increasing the amount of BcsA-B from 25 (I-25) to 400 μg (I400) during immobilization. Initially, 25 μg (3.125 μM) of BcsA-B produced microfibrils exhibiting 7−10 nm/8.5 ± 1.5 nm cross-sectional areas (Figure 2h). However, microfibrils produced by 100 μg or 12.5 μM (I-100) and 200 μg or 25 μM (I-200) of BcsA-B show 5−10 nm/7.5 ± 2.5 nm cross-sectional areas (Figure 3a,b). The number of thin (5 nm) cross-sectional microfibrils increased by increasing the BcsA-B amount from 100 to 200 μg (Figure 3a,b). Moreover, in the presence of 400 μg (50 μM) of BcsA-B (I-400), a significant decrease of the microfibril cross section was observed, resulting in 5−6.5 nm thin cross-sectional microfibrils (Figure 3c). Under these experimental conditions, the BcsA-B concentration slightly exceeds the concentration of c-di-GMP, which likely results in

Figure 3. Effect of the amount of BcsA-B enzymes, UDP-Glc, c-diGMP, and Mg2+ ions: (a) I-100, (b) I-200, (c) I-400, and (d) I-40016. AFM image scan size = 2 μm × 2 μm.

the presence of inactive enzymes and suggests that the density, that is, the number of active CS, directly influences cellulose microfibril morphology. Because 400 μg of BcsA-B is 16 times the amount of 25 μg of BcsA-B during immobilization, microfibril formation was also studied in the presence of 16 times the substrate (80 mM UDPGlc) and cofactor (480 μM c-di-GMP; 320 mM MgCl2) concentrations. The I-400-16 (400 μg + 16 times more substrate and cofactors) system showed a significant increase in microfibril cross section around 13−17 nm/15 ± 2 nm (Figure 3d). These results suggest that the increased cofactors and substrate increase the rate of cellulose production. Since the amount of enzyme on the surface has remained the same in I400 and I-400-16, the increased microfibril thickness must result from a change in the structuring of the cellulose composing the microfibril as a result of increased number of cellulose chains synthesized. In addition, more CS may be active due to the presence of additional c-di-GMP in the assay buffer. Fourier Transform Infrared (FTIR) Analysis. Bacterial cellulose was previously characterized by FT-IR spectroscopy, which can easily distinguish the different allomorphs (Iα, β) and polymorphs (cellulose I, II, IIII, IIIII) of cellulose.35−38 IR studies of immobilized BcsA-B-originated (I-25, I-400, and I400-16) cellulose show sharp peaks near 3445 and 3490 cm−1 (Figure 4a), which are specific to cellulose II dipole moments aligned along the cellulose chain direction and are assigned to OH stretching vibrations arising from intrachain hydrogen bonding between O3H···O5 groups.38−40 The O2H···O6 interchain hydrogen bonding is also present in cellulose II since the exocyclic −CH2OH group at the C-6 position is in the gt conformation, where intrachain hydrogen bonding for O2H and O6H is not possible.41 The related Iα (750 and 3240 cm−1) and Iβ (710 and 3270 cm−1) IR peaks were completely absent. Therefore, immobilized BcsA-B-produced cellulose is cellulose II in nature and not cellulose I according to IR spectra. The OH stretching vibrations at 3445 and 3490 cm−1 increased D

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Figure 4. FT-IR spectra of BcsA-B synthesized cellulose II.

with an increasing amount of enzyme, substrate, and cofactors in immobilized conditions (Figure 4a), which suggests that the cellulose II crystallinity depends on the enzyme, substrate, and cofactor ratio since the peak position of the OH stretch is sensitive to the strength of the hydrogen bonding interactions and thus increasing crystallinity.42 Cellulose samples were also synthesized from BcsA-B in solution using the same enzyme, substrate, and cofactor ratios as described in immobilized conditions. These samples are referred to as T-25, T-400, and T-400-16 and were examined by FT-IR in the same manner as the immobilized samples (I25, I-400, and I-400-16) described above. The T-25 and T-400 samples showed very weak/broad peaks around 3445 and 3490 cm−1, which represents the very low crystalline cellulose II or amorphous cellulose. However, T-400-16 shows higher peak intensities at the same IR regions (3445 and 3490 cm−1) than T-25 and T-400 samples (Figure 4a), which suggests that the cellulose II can also form in solution, provided that the enzyme (and thus glucan) concentration is high enough. Crystallinity of the cellulose produced was then examined to determine the effect of the different factors (immobilization, mobilization, amount of the enzyme, substrate, and cofactors) on the BcsA-B-synthesized cellulose II structure. Previously, cellulose I crystallinity was calculated from the ratio of the absorption bands around 1280 and 1200 cm−1.43 However, the absorbance peak at 1280 cm−1 of cellulose I is shifted to 1278 cm−1 in cellulose II.44 Therefore, the ratio of 1278 and 1200 cm−1 absorption bands (Figure 4b) was used to determine the crystallinity of BcsA-B-synthesized cellulose II in this study. The I-400-16 showed the maximum crystallinity around 97%, which is 8 and 12% higher than the I-400 and I-25, respectively (Table 1). A decrease of the band at 1278 cm−1 (C−H bending) is observed with decreasing crystallinity. Therefore, enzyme, substrate, and cofactor ratios play an important role in cellulose II crystal structure formation under immobilized conditions. The absorbance peak at 1263 cm−1 (in-plane C− OH bending at C-2 or C-3) also increased with cellulose II crystallinity44 and further confirms that immobilized BcsA-B enzymes synthesize cellulose II (Figure 4b). Moreover under mobilized (T) conditions, T-25 and T-400 samples showed no significant peaks at the 1278 cm−1 region, which again suggests that the cellulose produced in T-25 and T-400 was not

Table 1. Crystal Size and Crystallinity Index of BcsA-B Synthesized Cellulose II sample I-25 I-400 I-400-16 T-25 T-400 T-400-16

XRD CI (%) [peak height]

XRD CI (%) [peak deconvolution]

B110 (nm) [peak deconvolution]

FT-IR CI (%)

84 ± 2.0 90 ± 1.0 95 ± 1.0

55 ± 2.0 61 ± 1.0 72 ± 3.0

6.0 ± 0.1 6.6 ± 0.1 7.6 ± 0.4

85 ± 2.0 89 ± 1.0 97 ± 1.0

79 ± 2.0

43 ± 3.0

4.6 ± 0.3

81 ± 2.0

crystalline cellulose II in nature. However, T-400-16 showed around 81% crystallinity comparable to the I-25 system (85%). X-ray Diffraction (XRD) Analysis. X-ray diffraction patterns associated with BcsA-B cellulose in the presence of different amounts of the enzyme, substrate, and cofactors in immobilized and mobilized conditions are shown in Figure 5. Crystalline cellulose II was formed by all three samples (I-25, I-400, I-400-

Figure 5. X-ray diffraction data of BcsA-B-synthesized cellulose II. E

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Figure 6. QCM study. (a) First mode center frequency shift over time by increasing the BcsA-B enzyme amount from 25 to 400 μg. (b) First mode center frequency shift over time in the presence of 400 μg of BcsA-B enzyme.

16) under immobilized conditions on Ni film since three characteristic peaks of BcsA-B cellulose at 12.1 (1−10), 20.0 (110), and 21.8° (200) were found, in agreement with previous studies on cellulose II.44−46 Under mobilized conditions (T), only T-400-16 shows crystalline cellulose II characteristics; however, T-25 and T-400 show no characteristic peaks of cellulose II and are therefore amorphous in nature (Figure 5). Table 1 summarizes the XRD data. The peak intensity at 20.0° (110) was considered to determine the crystal size and crystallinity index (CI) due to its higher intensity as compared to other peaks (12.1 and 21.8°). The peak height method was used to calculate the CI (%) of BcsA-B cellulose II according to previous work.44,45 Under immobilized conditions, it is apparent that the peak intensity associated with the (110) plane increases with increasing amounts of BcsA-B from 25 (I-25) to 400 μg (I400). The peak intensity was further increased in the presence of higher amounts of substrate and cofactors, as shown for the I-400-16 sample (Figure 5). The crystal size of I-25 cellulose II was 6.0 nm at the (110) plane, which is 0.6 nm lower than that of the I-400 cellulose II (6.6 nm). Moreover, in the presence of 16 times more substrate (UDP-Glc) and cofactors (c-di-GMP and Mg2+), the crystal size of I-400-16 is 7.6 nm at the (110) plane, which is 1.6 and 1.0 nm higher than that of I-25 and I400, respectively. Along with significant changes in crystal size, the CI value of I-400-16 was also 11 and 5% higher than that of the I-25 and I-400 systems, respectively, similar to FT-IR values (Table 1). Mobilized T-25 and T-400 cellulose samples showed amorphous character even in the presence of the same amount of the enzyme, substrate, and cofactors like I-25 and I-400, respectively. Under mobilized (T) conditions, only T-400-16 showed cellulose II characteristics (Figure 7); however, the crystal size and CI values were 1.4 nm and 5% lower than that of the I-25 even in the presence of 16 times more enzyme, substrate, and cofactors than the I-25 system (Table 1). On Ni films, the position of the BcsA-B subunits was restricted within a limited space and organized in a planar geometry, which does not allow the free movement of the catalytic subunits of the enzyme as well as the synthesized glucan chains, promoting aggregation and organization. The CI values have also been calculated by using the peak deconvolution method.47 By using this method, the CI values of I-25, I-400, and I-400-16 were 55, 62 and 73%, respectively (Table 1). The CI values obtained using the peak height

method are higher than values obtained by the peak deconvolution method.47 Peak height method always gives higher values compared to the peak deconvolution method because the peak height method tends to underestimate the amorphous components and therefore overestimates the crystallinity.11,47 However, the peak height method was used to determine the CI value of cellulose II more frequently than the peak deconvolution method.44−48 Adsorption Pattern of BcsA-B Enzyme Complex on Ni-Coated Quartz Crystal Microbalance (QCM) Resonator. In order to determine whether a packed monolayer of enzyme has formed on the Ni surface, QCM studies on enzyme adsorption using Ni-coated resonators were performed. Figure S4 shows an optical picture of the fabricated and packaged resonator array with eight resonators in the array and the Teflon fixture used in the measurement. Each resonator is individually addressed through backside (etched side) electrodes, which are extended to the rim of the sensor array chip. The top electrode is common to all the pixels of the array. The mounting and packaging of the array and the interconnection were also carefully optimized to maintain a high-quality factor of the resonators. The resonance frequency of the fabricated devices was measured to be 115 MHz with a Q-factor of 3573.535 in air and corresponds to a quartz resonator thickness of ∼14.1 μm, and QCM resonator frequency shift measurement was done in the presence of an increasing concentration of BcsA-B enzyme from 25 to 400 μg. Figure 6a shows the frequency shift of the resonators at various protein loading concentrations (25 → 100 → 200 → 400 μg). Another QCM resonator frequency shift measurement was done only in the presence of 400 μg of the BcsA-B enzyme complex by using a different resonator. The resonance frequency of the fabricated devices was measured to be 63.92 MHz with a Q-factor of 10 136 in air and corresponds to a quartz resonator thickness of ∼25.5 μm. The packaged QCM device was placed into a plastic encasing, which held the device in place and provided a well for the liquid measurements. Initially, 200 μL of buffer A solution was deposited onto the micro-QCM device. After the resonator was allowed to stabilize, 400 μg of enzyme was added to the buffer A solution. The resonance shift was observed for almost 1 h (Figure 6b). For analysis, the adsorbing BcsA-B protein layer is considered to be viscoelastic in nature, whereas the surrounding buffer A solution, containing a very low volume fraction of enzyme molecules, can be represented as a Newtonian fluid. F

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Figure 7. Schematic model of the BcsA-B-derived cellulose II formation: (a) I-25, (b) I-400, and (c) I-400-16.

frequency and Q-factor shifts was 0.5 and 0.1%, respectively, thereby indicating a good convergence with the model.

Using expressions given by eqs 3 and 4, the expected frequency and Q-factor changes can be calculated. In order to model the system, the thickness and elastic modulus of the protein layer are considered as parameters and varied to fit to the experimental data. Based on the experimental results and the fit to the model, thickness and elastic modulus were calculated for the BcsA-B protein film adsorbed on the resonator. QCM studies showed that the surface density of the BcsA-B enzymes increased when the enzyme solution concentration increased from 25 to 400 μg. After 400 μg of BcsA-B enzyme was incubated on the Ni-coated QCM resonator for 30 min, a 19.65 nm layer of enzyme was formed on the surface (Figure 6b). Since one 12-His-tagged BcsA-B enzyme measures approximately 16 nm, it is hypothesized that a monolayer of enzyme was formed. Based on QCM analysis, the number density of BcsA-B enzyme on the surface is 7.65 × 1012/cm2. This is the same number density calculated assuming an enzyme with a cylindrical morphology hexagonally packed, where the long axis of the cylinder is perpendicular to the surface and the diameter or cross section of the cylinder is approximately 4 nm. The crystal structure of the enzyme14 suggests a maximum dimension of approximately 4 nm × 6 nm × 16 nm, where the 16 nm dimension would align with the long axis of the cylinder. This is slightly larger than the estimated cross-sectional size, but the actual size and conformation of the packed enzyme are unknown. The adsorption of enzyme from a 25 μg solution of BcsA-B enzymes formed a sub-monolayer (or patchy layer) with a resultant thickness much less than the saturation thickness, which is estimated from a very low frequency change (kHz) profile (Figure 6a). The effective thickness in this case is reduced to only 1.6 nm, which represents a fractional layer thickness due to the averaging effect. Furthermore, the elastic modulus decreased significantly (486.5 kPa) compared to that measured using 400 μg of enzyme, demonstrating that the film is not continuous. The viscosity of I-25 (0.0001987 mPa·s) is also much lower than that of the I-400 system (0.0023 mPa·s). On the basis of the QCM results, the proposed model (Figure 7) describes how the sub-monolayer and monolayer were formed and how the antiparallel glucan chains of cellulose II assemble from the immobilized BcsA-B enzymes. For these fitting parameters of the viscoelastic film on the resonator surface, the error between the calculated and measured

DISCUSSION In the past decades, membrane fractions containing CSCs were isolated from bacteria to study cellulose synthesis in in vitro conditions in the presence of UDP-Glc, c-di-GMP, and Mg2+ ions, which primarily produced cellulose II as well as amorphous cellulose.49−53 Therefore, native/cellulose I could not be synthesized in in vitro conditions due to some alteration of the CSC and/or functionally associated proteins present in native conditions.52,53 To examine the role of spatial positioning of CSCs on cellulose microfibril formation, functional His-tagged BcsA-B synthase derived from Rhodobacter sphaeroides was immobilized onto Ni surfaces with varying densities and examined under differing substrate, cofactor, and temperature conditions. The hypothesis was that cellulose microfibril size, crystal size, crystallinity, and allomorph may change as a function of enzyme surface density under immobilized conditions and the presence of substrate and cofactors, possibly with the emergence of cellulose I as the density of surface enzymes increased, where the enzymes would be closely packed, promoting the parallel glucan chain crystal organization found in cellulose I. The current system offers some exciting observations in this regard. By increasing the amount of BcsA-B enzyme complex from 25 to 400 μg, the surface density or positioning of immobilized BcsA-B enzyme complexes on the Ni film apparently affects the morphology as well as the crystal structure of cellulose microfibrils, as shown using AFM, FT-IR, and XRD analysis. However, surface-tethered BcsA-B enzyme produces crystalline celluose II and not cellulose I. The formation of cellulose II could arise simply from the random orientation of the enzymes on the surface, which may not align the produced glucan chains to promote the less energetically favorable parallel chain arrangement of cellulose I. However, the situation may be more intricate, as the BcsA-B enzyme studied here extrudes a glucan which is not positioned parallel to the long axis of the enzyme. Given the location of the His tag and the hypothesized orientation of the enzyme on the surface, it is quite possible that glucan chains are extruded in a direction initially more parallel to the substrate surface, as shown in G

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ACS Nano

enzymes but also position them spatially in an intricate fashion to predispose the extruded glucan chains to form cellulose I. Another possibility is that the Rhodobacter sphaeroides BcsA-B enzyme lacks some structural or functional feature, which would enable the production of cellulose I. Finally, other aspects of the model system (Figure 7) here may preclude its formation. As described above, the BcsA-B enzyme used in our studies is embedded in a detergent micelle that surrounds the transmembrane region of BcsA, which may preclude native-like interactions between neighboring BcsA-B enzymes. Under, immobilized conditions, closely positioned BcsA-B CSs produce crystalline cellulose II. In solution, the BcsA-B enzymes form generally disordered glucan chains, likely due to their random orientation and diffusion. These results strongly suggest that the spatial arrangement, and in particular the packing, of the cellulose synthase enzymes is important for synthesis of highly crystalline cellulose. The production of glucans with increased spatial density may promote improved interaction, resulting in additional hydrogen bonding and van der Waals interactions, which drives microfibril crystallization. Besides the spatial organization or orientation of the synthase subunits, a higher amount of cofactors and substrate also regulates the microfibril morphology and crystal properties, which could suggest that microfibril formation is a timedependent phenomenon where aggregation, crystallization, and organization of glucan chains depend on the rate of cellulose synthesis. For example, crystallization of cellulose after polymerization results in a precrystallization region where extruded glucans interact and order. An increased polymerization rate could lead to a larger precrystallization region allowing more glucan chains to interact at that stage, resulting in larger crystals, larger microfibrils, and larger aggregates (Figure 7a−c).

Figure 7. As the enzyme surface density increases, given the random orientation of enzymes on the surface, it may be more likely for glucan chains to interact in the antiparallel cellulose II configuration. This analysis, however, assumes that cellulose II forms through the interaction between multiple glucan chains. It has been hypothesized that an individual glucan chain with a high degree of polymerization can fold to form the antiparallel arrangement.54,55 In our studies, no cellulose II was observed under tube conditions without excess substrate, suggesting that this process may not occur spontaneously. However, once the cellulose produced begins to aggregate and form a network, the force of polymerization associated with an immobilized enzyme may provide the energy needed to fold an extruding glucan chain which has at least a portion that is no longer free but associated with a larger network of cellulose. In native or in vivo conditions, thermodynamically unfavorable parallel chain cellulose I was synthesized from unidirectional oriented glucan chains with the same polarity.54 This enigmatic process could occur through the combination of spatial arrangement within the CSC as well as in the presence of other associated factors. All microfibril-forming bacterial cellulose synthases (Bcs) also contain the BcsD subunit,8,56 which could be implicated in organizing the BcsA-B complexes in the membrane and assists the formation of crystalline cellulose.19 A previous study also showed that the acsD or bcsD gene is required for normal cellulose production in Acetobacter xylinum, and mutation leads to a very low yield of low crystalline cellulose I and II allomorphs.19 Thus, in the absence of the BcsD subunit, the native crystallization of cellulose I in in vitro conditions is disturbed. Subsequent work has shown that the COBRA protein in plant systems is required for the parallel orientation followed by deposition of cellulose I microfibrils in native conditions.57 Another recent study proposes that COBRA is a “polysaccharide chaperon” that acts transiently at the initiation of cellulose microfibril formation, causing cellulose I to form as glucans emerge from the cellulose synthase complex.58 Therefore, on the basis of these three concepts (absence of BcsD subunit and polysaccharide chaperon protein, and the orientation of BcsA-B enzyme on the surface), a conceptual model (Figure 7) for the molecular arrangement of antiparallel folded glucan chains has been developed, which could explain the formation of cellulose II by immobilized BcsA-B enzymes in this in vitro system. Such a process could also be responsible for the rough microfibrils observed in Figure 3d. In fact, observed microfibrils may not resemble the traditional cellulose I microfibril thought to exist in natural systems but rather may consist of a string of ordered domains of cellulose II formed from the association of multiple glucan chains and possibly folded individual chains. Previously, it was proposed that a larger number of CS proteins may be essential to the production of cellulose I.53 To explore this possibility, I-400 and I-400-16 systems were examined, which produced densely packed surface-immobilized CSs. However, additional enzymes, cofactors, and substrate did not result in the formation of cellulose I. Therefore, it could be concluded that the assembly of the cellulose I microfibril needs other accessory proteins that allow a unique assemblage of catalytically active subunits for the formation of the native cellulose I. It would also be important to determine the nature or type of interaction between the CS subunits in the organization of CSC required for the formation of cellulose I. The assembly of the native CSC may not only aggregate CS

CONCLUSION Functional His-tagged BcsA-B cellulose synthase enzymes were immobilized on Ni films and cellulose microfibril morphology; crystal size and crystallinity changes were examined as a function of enzyme surface density. Highly crystalline cellulose II emerged as the density of surface enzymes increased. This resulted from the spatial organization and packing density of the immobilized enzymes. Several possibilities for the absence of cellulose I were discussed. The demonstrated system allows much future work on cellulose biosynthesis to be performed. The role of the BcsD subunit and polysaccharide chaperon protein (e.g., COBRA) can be studied using this system to determine its impact on cellulose allomorph formation. In addition, other surfaceimmobilized or solution-phase scaffolds can be created to more closely mimic a CSC by better controlling the number or spatial position of CS enzymes within a complex. Such engineered CSCs or rosettes could enable a deep understanding of the relationship between CS assembly and cellulose formation, paving the way for engineered cellulose and lignocellulose for many applications ranging from improved feedstocks for biofuels to nanomaterial production. MATERIALS AND METHODS Sputter-Deposited Nickel Thin Films on Si Substrates. Nickel thin films of ∼50 nm were deposited using a Kurt J. Lesker CMS-18 DC (MRI, Penn State University, USA) sputtering system. The sputter sources were optimized to have a uniform thickness/composition over a 6 in. diameter Si wafer. The base pressure of the vacuum chamber H

DOI: 10.1021/acsnano.5b05648 ACS Nano XXXX, XXX, XXX−XXX

Article

ACS Nano was below 2.0 × 10−7 Torr. The films were deposited for 500 s at room temperature at an argon pressure of 5 mTorr to produce asdeposited films. The power was maintained at 200 W for the Ni target. The deposition rate at these conditions is around 1.0 Å/s. To maintain uniformity of the film, the substrate stage was rotated during all depositions. Before every deposition, sputter cleaning of the targets was performed. The target substrate distance was fixed at 6 in. After Ni coating, the 6 in. Ni-coated wafer was broken into small pieces (1 cm2) for experimental purposes. The pieces of Ni thin film containing Si wafers were stored under appropriate conditions (e.g., vacuum) to prevent oxidation of the Ni surface prior to experimental use. BcsA-B Enzyme Expression and Purification. The bacterial cellulose synthase bcsA and bcsB genes from Rhodobacter sphaeroides were cloned into the pETDuet (Novagen) expression vector as described previously.14 BcsA contains a C-terminal dodeca-histidine tag for metal affinity purification. BcsA and BcsB were expressed in Escherichia coli C43 in autoinduction medium and were purified by metal affinity (Ni-NTA agarose, QIAGEN) followed by size-exclusion chromatography as described previously14,15 with the exception that 1 mM LysoFos choline ether 14 (LFCE14) detergent was used instead of LDAO (lauryl-N,N-dimethylamine oxide). The purified complex was concentrated to 10.0 mg/mL final concentration and remained active for more than 1 month stored at 4 °C. Immobilization of Histidine-Tagged BcsA-B Enzyme on Ni Thin Film and Synthesis of Cellulose. The purified His-tagged BcsA-B complex (25−400 μg) was incubated with the Ni thin film for 30 min at room temperature to allow proper immobilization through binding of histidine residues to Ni. Nonspecific binding on the Ni film was removed by washing in the presence of buffer A (25 mM sodium phosphate pH 7.4, 0.1 M NaCl, 5 mM cellobiose, 10% glycerol, 1 mM LFCE14). Immobilized BcsA-B was incubated in the presence of 5.0 mM UDP-Glc, 30 μM c-di-GMP, and 20 mM MgCl2 in buffer A at 37 °C for 60 min to synthesize cellulose. The polymerization reaction was terminated with 40 mM EDTA. Sample Preparation for AFM Study. The EDTA-treated samples were washed in the presence of sterile, filtered deionized water (DI) for 10 min using a shaker at 150 rpm followed by vacuum drying using N2 gas at room temperature just before the AFM study. Sample Preparation for FT-IR and XRD Study. The EDTAtreated samples collected from the Ni film surface were centrifuged at 14 000g at room temperature for 20 min. The supernatant was discarded, and the pellet was washed twice in 500 μL of sterile DI water (filtered), freeze-dried for 12 h, and characterized using FT-IR and XRD. Atomic Force Microscopy. The dried samples were imaged with a Dimension Icon atomic force microscope (Bruker, CA, USA) operated in ScanAsyst and PeakForce tapping mode. Nanoscope (v 8.10b44) and Nanoscope Analysis (v 1.40) software were used for AFM operation and image analysis. SCANASYST-AIR+ probes (Bruker, CA, USA) with a spring constant of 0.4 N/m, deflection sensitivity factor of 60 nm/V, and a tip radius of 2 nm (nominal) were used for all experiments. All samples were scanned at 512 × 512 sampling rates. The scan size, scan rate, and peak force set point were 2 μm × 2 μm, 0.5 Hz, and 500 pN, respectively. The cross section/ diameter and length of the cellulose fibers were calculated by using a step method tool available in Nanoscope Analysis software (v 1.40). Before step analysis, a “Flatten” tool was used to process each and every image. Step analysis makes relative height measurements between two regions (steps) on sample surfaces and works with an averaging box cursor drawn on the cellulose fibril surface of interest and determines the average height. The average height measurement was performed after selection of 25 different spots in each image. The cross section and the length measurements were obtained from the height mode AFM images for each sample. At least five different samples from each set were scanned, and representative images were chosen for analysis. The AFM scanning of cellulose fiber under hydrated conditions was performed after 4 min of cellulose synthesis. SCANASYST-FLUID+ probes (Bruker, CA, USA) with a spring constant of 0.7 N/m, deflection sensitivity factor of 30 nm/V, and a tip radius of 2 nm

(nominal) were used for all experiments. All samples were scanned at 512 × 512 sampling rates. The scan size, scan rate, and peak force set point were 2 μm × 2 μm, 0.5 Hz, and 450 pN, respectively. Fourier Transform Infrared Spectroscopy. FT-IR spectra were collected using a FT-IR spectrometer (Bruker Vertex V70) equipped with an MVP-PRO ATR unit (diamond crystal, 45° angle of incidence) and a KBr beam splitter. All spectra were collected from the freeze-dried sample pellet in the region of 400−4000 cm−1 with a 6 cm−1 resolution and averaged over 100 scans. The one-point baseline correction of FT-IR spectra at 3800 cm−1 absorbance was performed with OPUS software (Bruker Optics). The 3800 cm−1 absorbance was set to zero due to the absence of any absorbance feature. The degree of crystallinity was evaluated from the ratio of the absorption bands around 1278 [C−H bending] and 1200 cm−1 [in-plane C−OH bending at the C-6 position].43,44 X-ray Diffraction. Crystalline structures of the freeze-dried cellulose samples were analyzed by XRD using a Rigaku DMAXRapid II microdiffractometer with Cu Kα radiation with a wavelength of λ = 0.15406 nm generated at 50 kV and 40 mA in transmission geometry. The 0.3 mm collimator was used. The data were integrated (intensity vs 2θ) with a step size of 0.02° across the range of 5−40° 2θ using AreaMax (Rigaku software). By using the peak height method, CI (crystallinity index) was calculated for cellulose II as the ratio of the crystalline scatter of the 110 reflection at 2θ of 20.0° (crystalline height, Icr) with the height of the amorphous reflection at 2θ of 16° (amorphous height, Iam) by using eq 1.44,45,48,59 MDI Jade 2010 software (Materials Data, Inc., Livermore, CA) was used to process the diffraction patterns and to determine intensity values for eq 1 after linear baseline correction. CI = [(Icr − Iam)]/Icr] × 100

(1)

where Icr is the intensity of the 110 reflection/20° at 2θ, Iam is the minimum intensity between the 1−10 and 110 reflections/16° at 2θ. Alternative XRD analyses (peak deconvolution) were used, which provide more accurate CI assessment for cellulose I.47 However, eq 1 was used more often specifically to assess CI for cellulose II.44,45,48,59 Given that cellulose II was observed in our studies (see below), eq 1 was also used to assess CI. In the peak deconvolution method, the diffraction peaks were fitted with pseudo-Voigt peak functions using MDI Jade 2010 software. In this method, a pseudo-Voigt function was used to profile the peak shape and area, assuming a linear background. A broad peak around 21.5° was assigned to the amorphous contribution. Crystallinity was calculated from the ratio of the area of all crystalline peaks to the total area.47 The dimension of the crystal perpendicular to the diffracting planes with hkl Miller indices, Bhkl, was evaluated by using Scherrer’s expression (eq 2).60

Bhkl =

Kλ (Δ2θ )2 − (Δ2θins)2 cos θ

(2)

where Bhkl is the average crystalline width at the (110) plane, K is the shape factor (K = 0.9), λ is the wavelength of incident X-rays (λ = 0.15406 nm), θ is the Bragg angle, Δ2θ is the fhwm of the reflection peak, and Δ2θins is the instrumental broadening. The instrumental broadening was determined from the fwhm of four reflections of a silicon standard (NIST Si 640). Quartz Crystal Microbalance. A micromachined quartz resonator array was used to measure the changes in the frequency and the Q-factor upon adsorption of the His-tagged BcsA-B enzyme complex as a function of concentration in the solution phase. The QCM frequency shift resulting from the deposition of a viscoelastic layer, such as a protein/enzyme film, in a viscous liquid can be analyzed using a continuum mechanics approach.61,62 In order to model this situation, the QCM surface is considered to be in intimate contact with a continuous viscoelastic layer with an infinitely thick Newtonian liquid overlayer on one of its surfaces. Under the assumption that the thickness of the bulk liquid layer is much larger than the decay length of the acoustic wave in the liquid, the frequency and Q-factor changes with respect to liquid loading conditions can then be written as62 I

DOI: 10.1021/acsnano.5b05648 ACS Nano XXXX, XXX, XXX−XXX

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ACS Nano Δf ≈ −

⎞ ⎛ ⎛ ηliq ⎞2 ηviscω2 1 ⎜ ⎟ ⎟⎟ tviscρvisc ω − 2tvisc⎜⎜ 2 2 2 ⎟ 2πρq tq ⎜ δ liq ⎠ μvisc + ω η ⎝ visc ⎠ ⎝

⎛ ⎞ μvisc ω ⎜2t ⎟ 2 2 + ω2ηvisc ⎝ visc μvisc ⎠ ΔQ ≈ − 2πf0 ρq tq ⎡ ⎛ ⎛ ηliq ⎞ μ ω ⎞⎤ ⎢1 + ⎜2tvisc⎜ δ ⎟ μ2 +viscω2η 2 ⎟⎥ ⎝ liq ⎠ visc ⎝ ⎣ visc ⎠⎦

Additional results in Figures S1−S4 (PDF) (3)

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

(4)

where μvisc, ηvisc, and ρvisc are the elastic modulus, viscosity, and density of the viscoelastic layer, respectively, and tvisc is its thickness and ηliq is the viscosity of the liquid overlayer (buffer A solution); δliq is the penetration depth of the acoustic wave in the liquid; f 0 is the resonance frequency of the resonator, ρq is the density of quartz, tq is the thickness of the quartz resonator, and ω is the angular frequency = 2πf, where f is the frequency under viscoelastic loading conditions. It has been demonstrated that high-frequency micromachined quartz resonators are especially suited for monitoring small changes in the viscoelastic loading in films adsorbed on their surface. Miniaturized resonators provide a greater resolution of the density, thickness, viscosity, and the elastic modulus properties of the thin adsorbing films.63 A large variation in the Q-factor and the frequency for small changes in the viscoelastic properties of the adsorbed film is highly desirable for accurate determination of material properties and for the observation of conformational changes in these nanometer thin films. Thus, careful interpretation of the frequency and Q-factor changes can provide a sensitive tool for the analysis and investigation of the biomolecular films in terms of film assembly, packing density, and other conformational changes. The fabrication of the devices from polished 1 in. diameter AT-cut quartz disks was carried out using a recently developed inductively coupled plasma etch process operating at 1 mTorr to provide 76 μm deep × 500 μm diameter × 24 μm floor thickness circular wells with a mirror finish (