Two-Photon Fluorescence Anisotropy Imaging to Elucidate the

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Two-photon Fluorescence Anisotropy Imaging to Elucidate the Dynamics and the Stability of Immobilized Proteins Alejandro H Orrego, Carolina García Rodríguez, Jose Miguel Mancheno, José Manuel Guisán, Maria del Pilar Lillo, and Fernando López-Gallego J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.5b12385 • Publication Date (Web): 30 Dec 2015 Downloaded from http://pubs.acs.org on January 2, 2016

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The Journal of Physical Chemistry

Two-photon Fluorescence Anisotropy Imaging to Elucidate the Dynamics and the Stability of Immobilized Proteins Alejandro H. Orrego#, Carolina García⊥, José M. Mancheño§, Jose M. Guisán#, M. Pilar Lillo⊥* and Fernando López-Gallego†ξ* # Enzymatic Engineering group. Instituto de Catálisis y Petroleoquímica. CSIC, c/ Marie Curie 2, 28049, Madrid, Spain §

Crystallography and Structural Biology Group. Instituto Química Física “Rocasolano”, CSIC, Serrano 119, 28006 Madrid, Spain ⊥

Fluorescence Molecular Biophysics Group. Instituto Química Física “Rocasolano”, CSIC, Serrano 119, 28006 Madrid (Spain) †Heterogeneus biocatalysis group. CIC BiomaGUNE, Pase Miramon 182, 20009, San Sebasitan-Donostia (Spain) ξIKERBASQUE, Basque Foundation for Science, Bilbao (Spain)

KEYWORDS: Green fluorescent protein. Time-resolved fluorescence. Protein stability Enzymes. Heterogeneous biocatalysis.

ABSTRACT

Time/spatial-resolved fluorescence determines anisotropy values of supported-fluorescent proteins through different immobilization chemistries, evidencing some of the molecular mechanisms that drive the stabilization of proteins at the interfaces with solid surfaces.

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Fluorescence anisotropy imaging provides a normalized protein mobility parameter that serves as a guide to study the effect of different immobilization parameters (length and flexibility of the spacer arm and multivalency of the protein-support interaction) on the final stability of the supported proteins. Proteins in a more constrained environment correspond to the most thermostable ones as was showed by thermal inactivation studies. This work contributes to explain the experimental evidences found with conventional methods based on observable measurements; thus this advanced characterization technique provides reliable molecular information about the immobilized proteins with sub-µm spatial resolution. Such information has been very useful for fabricating highly stable heterogeneous biocatalysts with high interest in industrial developments.


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INTRODUCTION Deep understanding of the molecular mechanisms that drive the stabilization of proteins at the interfaces with solid materials is fundamental to develop highly active and stable immobilized proteins1. In particular, supported enzymes are extensively used as heterogeneous biocatalysts to carry out modern chemical processes since they are highly active and selective, enable the separation and recycling of the catalyst, facilitate “in-flow” reactions and thus increase both efficiency and sustainability of the process2. Thus far, techniques for molecular characterization of solid-supported enzymes are scarce unlike chemical catalysis where molecular characterization drives the design and optimization of heterogeneous catalysts3. Recently, a handful of techniques allow the advanced characterization of heterogeneous biocatalysts, shedding light on the structure and the dynamics of the immobilized proteins

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. In addition,

recent advances in fluorescence microscopy have enabled to study single molecules in heterogeneous environments informing about spatial, dynamical and structural organization of proteins across 3D-surfaces4–6. A myriad of enzymes have been stabilized by attaching them to solid supports through different immobilization chemistries, but conclusions about the stabilization mechanisms have relied on observable measurements rather than in single molecule studies that may elicit information about the molecular basis of such stabilization. Time-resolved fluorescence is extremely useful to understand the dynamics of many biological processes8,9 and the effect of the molecular crowding on the protein dynamics and orientation. Rotational and segmental motions of proteins can be characterized by measuring the fluorescence anisotropy of chromophores attached to the biomolecules10. The fluorescent molecule, excited by a short (femtosecondpicosecond) linearly polarized light pulse at time 0 (t0) and undergoing global and/or local


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rotations before emitting light at time (t1), may change the angle between its absorption and emission transition moments, resulting in fluorescence depolarization that decreases the steadystate fluorescence anisotropy regarding to its fundamental anisotropy value, (r0)11. This depolarization effect depends, among other factors, on the excited state lifetime (τ), the type of molecular bond of the fluorophore, the size and the shape of the rotating molecule, and the viscosity of its micro-environment11. Hence, combining time-resolved fluorescence anisotropy with confocal or multiphoton steady-state polarized microscopy would provide priceless information to understand protein dynamics with sub-µm spatial resolution. Such technical synergy may furnish experimental evidences for enzyme flexibility to address long-standing questions about operational performance of the heterogeneous biocatalyst. Unfortunately, time and spatial-resolved fluorescence polarization has been rarely used to characterize heterogeneous biocatalysts, and the handful of existing examples involve enzymes entrapped into sol-gel materials4,5 -proteins floating around the microstructure- rather than directly attached to the solid surface of porous microbeads. We know from stability data that a short spacer arm and an intensive attachment between the protein and the support surface increase the protein stability, but we poorly understand why those two parameters control the stability of immobilized proteins1,12. Based on such experimental evidence, we hypothesize that the protein fluorescence anisotropy informs about the degree of constraints to the global rotation of the immobilized proteins onto a solid surface, and consequently the fluorescence anisotropy values depend on the immobilization chemistry and specifically on the length and flexibility of the spacer arm and the number of bonds between the protein and the support surfaces. To proof this hypothesis, we have used fluorescence lifetime imaging (FLIM) and spatial-resolved fluorescence polarization to determine the fluorescence


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anisotropy of supported-proteins through different immobilization chemistries. Moreover, the experimental setup herein described allows discriminating between depolarization effects from protein mobility and possible optical artefacts of the agarose-based support material. EXPERIMENTAL SECTION Materials, bacteria strains, plasmids, protocols for the activation of solid supports and theoretical details are described in supporting information. Immobilization of fluorescent proteins on different agarose-type carriers 1g of each support was incubated in batch with 10 mL of 0,1 mg/mL solution of different proteins (LSL-EGFP, LSL-BTL2, LSL-BGL, EGFP, BTL2 and BGL) under different conditions. The residual protein in the supernatant was calculated by measuring the absorbance of EGFP at 487 nm or by colorimetric activity assay for BTL213 and BGL14. The immobilization conditions varied according to the immobilization protocol. While the immobilization on Ag10-Ni2+ support was carried out with 25mM sodium phosphate at pH 7 and 25 ºC, the immobilization on Ag10-G support was carried out with 100 mM sodium carbonate at pH 10 and 25 ºC. On the other hand, 1mL of 0.1 mg/mL His-EGFP and 1mg/mL His-LpPDC were co-immobilized on 0.1 g of AgNi2+ support with 25mM sodium phosphate at pH 7 and 25 ºC. Two-photon microscopy of immobilized derivatives of fluorescent proteins Each solid sample after immobilization was resuspended in 20 mM Tris-HCl buffer, NaCl 100 mM, pH 8.0. 30 µL of such suspension was analyzed by two-photon fluorescence polarization microscope at T=22 ºC (see Supporting Information). Thermal denaturation and inactivation of immobilized proteins For thermal denaturation of EGFP, 10 mg of immobilized EGFP were incubated with 100 µL of 25 mM sodium phosphate buffer at pH 7 in a PCR tube. The samples were denatured with a


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dynamic temperature ramp (20 ºC-100 ºC). For thermal inactivation, different preparation of immobilized enzymes and proteins were incubated in 1 mL of 25 mM sodium phosphate buffer at pH 7 at different temperatures, and samples were withdrawn after 1h and measured their residual activities. EGFP was inactivated at 55 ºC, BTL2 was inactivated at 60 ºC and BGL was inactivated at 45 ºC.


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RESULTS AND DISCUSSION Immobilization of different EGFP variants on different agarose-based carriers We selected enhanced green fluorescent protein (EGFP) as the model protein for this work, because its fluorescence properties and thermal stability have been broadly studied15,16. EGFP is the ideal fluorescence probe for these studies since it is a rigid and symmetrical protein barrel where the fluorophore is part of protein structure, and its fluorescence and hydrodynamic properties are directly related to the protein structural integrity. The mobility of the fluorophore strictly depends on the protein mobility when the protein is attached to the support surface. Besides, we also selected agarose microbeads (Ag10) as model support because their transparency lets two-photon polarized excitation deep-material imaging, their 3D-microporous structure allows proteins freely diffusing through, and their thick fibers enable planar and direct attachment of diffused proteins17. Different EGFP variants bearing different spacer arms at their N-terminus were immobilized on Ag10 presenting the same pore size (100 nm) and fiber thickness range (25-40 nm) but activated with different reactive groups (Figure 1a-c). Firstly, we utilized the fusion protein LSL-EGFP formed by the lectin domain of the haemolytic toxin LSLa from the mushroom Laetiporus sulphureus (LSL150) genetically fused to the N-terminus of EGFP13. This fusion protein was immobilized on plain Ag10. Here, the lectin domain binds the galactose-based carbohydrates on the agarose surface which locates the EGFP core at 8.6 nm far away from the agarose surface, and avoids direct interactions between EGFP and support surfaces. Secondly, the EGFP with a hexa-His tag at its N-terminus (His-EGFP)18 was immobilized on Ag10 activated with nickel-chelates (Ag10-Ni2+). His-EGFP is attached to the surface through a shorter spacer arm than LSL-EGFP, because of the single coordination bond


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between the His-tag and the metal chelates. In this case, the EGFP core is positioned 3.4 nm away from the agarose surface.

Figure 1. Different immobilization chemistries on porous agarose beads. Panels represent the different EGFP variants immobilized on the activated nanofibers that form the porous agarose beads. LSL-EGFP/Ag10 (a), His-EGFP/Ag10-Ni2+ (b) and EGFP/Ag10-G (c). The dimension of protein and pores do not correspond to their real scale (EGFP = 2.4 x4.2 nm and pore size = 100 nm), we magnify the protein size for figure clarity. Experimental time-resolved anisotropy decays of soluble and representative immobilized EGFP variants (d). Finally, we immobilized EGFP on Ag10 activated with glyoxyl groups (Ag10-G) through multiple amine-aldehyde interactions between surface lysine residues on EGFP and the support


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surface. The EGFP is directly and multivalently interacting to the support surface, here approximately 2 nm separate the EGFP core from the agarose surface; such distance is similar to the width dimensions of the EGFP barrel18. Two-photon time-resolved fluorescence anisotropy decays of different soluble and immobilized EGFP variants The two-photon time-resolved polarization microscope set-up (see Supporting Information), unlike cuvette measurements and conventional confocal microscope set-ups, may provide us unique information about protein dynamics with sub-µm spatial resolution across the porous structure of microbeads, up to 100 µm deep-material imaging. Using such set-up, we firstly measured the experimental steady-state and time-resolved anisotropy decays of soluble EGFP variants; EGFP, His-EGFP and LSL-EGFP. The experimental parallel and perpendicular polarized fluorescence intensity decays, Ipar (t) and Iper (t), of the three variants were globally analyzed (see Supporting Information), by linking the fluorescence lifetime and the time zero anisotropy parameters, since their values depend only on both the fluorophore (EGFP) and the instrument optical setup, which are common for the three samples: τEGFP =2.7±0.1 ns; r(0)EGFP = 0.458 ± 0.005. Expectedly, EGFP and His-EGFP presented similar steady-state anisotropy (rEGFP = rHis-EGFP = 0.387±0.004), and rotational correlation time (φR

EGFP

= φR

His-EGFP

= 14 ± 1 ns)

values in buffer solution. These results perfectly agree with previous ones from one-photon excitation studies, using a cuvette time-resolved fluorescence system13, revealing that the EGFP fluorophore is rigidly bound to the protein scaffold and rotates together with it16,19. Soluble LSLEGFP presented a steady-state anisotropy (rLSL-EGFP = 0.406±0.004) and a global rotational correlation time (φR LSL-EGFP = 21 ± 1 ns) values higher than non-tagged EGFP in solution. These values are due to the higher molecular weight of LSL-EGFP (45 kDa) compared to the EGFP (27


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KDa). Nevertheless the fluorescence lifetime of the fluorophore was not affected by the lectin domain. This fact is possible since LSL-EGFP is a fusion protein of two domains connected by a flexible linker that enables their free motion but with some interference13. Hence, the fluorescence and rotational data of LSL-EGFP demonstrate that the spectroscopic behavior of EGFP is negligibly affected by the lectin domain although it did reduce the EGFP mobility. Next, we measured the experimental time-resolved anisotropy decays for representative regions of interest (ROIs) selected from XY anisotropy images of 3 EGFP variants immobilized on different agarose beads (Figure 1a-c) at different XY and depths locations. Figure 1d shows representative anisotropy curves for the different preparations. The time-resolved fluorescence depolarization varies with the length and flexibility of the spacer arm and the multivalency of the enzyme-support interaction. The maximum and minimum depolarization correspond to free EGFP in buffer (grey) and bound to Ag10-G (dark pink), respectively (for theoretical details see supporting information). Noteworthy, aldehyde chemistry drives to an EGFP preparation whose anisotropy remains invariable along the time (ns) (Figure 1d), suggesting that the EGFP immobilized on Ag10-G is substantially less mobile in the nanosecond time range than the same protein in the other preparations. Moreover, we have observed that experimental time-resolved polarized intensity functions (Ipar(t) and Iper(t)) show significant relative time-shift, shape and size changes for certain regions in the bead border (1-2 µm deep into the outer layer), for those samples immobilized on Ag10Ni2+ and Ag10-G beads (Figure S1). These distortions are more important for His-EGFP/Ag10Ni2+ samples, and they indicate that other depolarization processes occur in addition to protein dynamics, giving rise to an important decrease in the experimental r(0) and r values in these regions. One possible explanation for these distortions may be the existence of some


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microstructure defects at the bead border, which either alter the refraction of the light or generate an internal birrefringency of the agarose, deforming the polarized intensity functions and invalidating, only for such small area, the interpretation of experimental anisotropy values in terms of simple protein mobility. Such anomaly was not observed in plain Ag10, suggesting that those material defects might occur during the agarose activation with different reactive groups (metal chelates and glyoxyl groups). Despite such technical issue, anisotropy values at particle depths > 2 µm still serve to characterize the degree of constraints to the rotational motions of the immobilized proteins since anisotropy is independent on spatial protein localization (Figure 2ab). Two-photon steady-state fluorescence anisotropy imaging of different soluble and immobilized EGFP variants We determined both rotational properties and spatial localization of the 3 different EGFP variants immobilized as above described (Figure 1a-c) from steady-state anisotropy imaging (for data acquisition and analysis details see supporting information). XY sections of 10 different beads, at different Z depths, for each type of EGFP immobilization were measured (Figure 2). Fluorescence anisotropy images of EGFP samples immobilized on Ag10-Ni2+ and Ag10-G beads show lower anisotropy values at the bead border (green-yellow and yellow-orange colored regions respectively; Fig. 2b and S2). Based on such experimental data, average steady-state anisotropy values, , were determined from all the fluorescence anisotropy images for each preparation, eliminating the regions with anomalous low values of the anisotropy (see above). Figure S2 shows the steady-state anisotropy values per pixel determined along the diameter of the XY anisotropy images presented in Figure 2b and Figure S3. LSL-EGFP immobilized on Ag10 (= 0.424±0.005) and His-EGFP immobilized on Ag10-Ni2+ (= 0.430±0.005)


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presented slightly different values, whereas we observed significantly higher values for EGFP immobilized on Ag10-G (= 0.458±0.006). Accordingly, such latter immobilization chemistry drives to an EGFP preparation whose time-resolved anisotropy decay remains invariable along the time (ns) (Figure 1d). Both anisotropy decays and steady-state anisotropies support that the aldehyde chemistry drives to a much less mobile EGFP than in the other immobilized preparations.

Figure 2. Fluorescence intensity (a) and fluorescence anisotropy (b) of XY sections (80 x 80 µm) at the equator of the representative microbeads immobilizing different EGFP variants through different chemistries: Grey scale corresponds to fluorescence intensity (It). Rainbow-like scale corresponds to fluorescence anisotropy (). Fluorescence intensity allows us to characterize the spatial location of the different EGFP variants across the porous structure of the differently activated agarose beads; we expectedly observed that different immobilization chemistries lead to different patterns for spatial distribution of EGFP, as we previously reported13,20 (Figure 2a and S3). However, figures 2a-b demonstrate that the fluorescence anisotropy was independent of the protein localization, since


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both His-EGFP and LSL-EGFP presented similar () values localizing at the outer part of bead and across the whole bead diameter respectively. Likewise, His-EGFP and EGFP immobilizates presented quite different () values although both proteins were heterogeneously distributed across the bead surfaces of both samples (Figure 2a-b). In regions with high density of EGFP proteins like the outer layer of those samples, the proximity of two vicinal EGFP molecules can provoke an energy transfer phenomenon (homo-FRET)9 that might cause fluorescence depolarization non related with protein dynamics, leading to lower (r) values depending on the relative orientation of the chromophores. To discard homo-FRET issues during the anisotropy measurements, we co-immobilized His-EGFP with 10-fold excess of a His-tagged but non fluorescent protein; p-coumaric acid decarboxylase from Lactobacillus plantarum-(HisLpPDC)21 on Ag10-Ni2+ (Figure S2-4), minimizing the possibility that two His-EGFP molecules were proximal enough to suffer homoFRET. We determined = 0.425 ± 0.006 that corresponds to the found for the His-EGFP solely immobilized on Ag10-Ni2+, demonstrating that even at high protein density, fluorescence anisotropy of immobilizates is not affected by homo-FRET issues. On the other hand, we incubated Ag10-Ni2+ microbeads immobilizing His-EGFP (( = 0.430 ± 0.005) with imidazole to in situ disrupt the proteinsupport coordination bonds, letting the protein float within the whole porous environment. We measured = 0.400 ± 0.005, a value closer to the soluble enzyme (Figure S2-3). This experiment evidences that the higher (r) value expressed by the His-EGFP immobilized on Ag10-Ni2+ without imidazole incubation was due to the bonding between both the protein and the agarose surfaces.


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Calculation and comparison of mobility factors of different immobilized EGFPs based on anisotropy values Interestingly, the experimental time-zero anisotropy value of free EGFP in a buffer solution ( r(0)EGFP ) represents the maximum experimental steady-state anisotropy value, rmax = 0.458 ± 0.005, that can be measured for a completely immobile EGFP protein with our instrumental setup, while steady-state anisotropy values of the soluble EGFP variants (rmin EGFP and rHis-EGFP = 0.387 ± 0.004; rmin LSL-EGFP = 0.406 ± 0.004) provide the minimum anisotropy value that means a fully mobile EGFP variants. Hence, rmax only depends on the fluorophore nature and the instrumental set-up, while rmin strictly depends on the fluorophore and the size and shape of the attached protein. These two parameters set the limits for comparing the anisotropy of different immobilized EGFPs in order to elucidate the effect of the immobilization chemistry on the final mobility of the heterogeneous proteins. To this aim, we have calculated a normalized mobility factor (Mf) based on the steady-state anisotropy (r) of each sample corrected by (r(0)EGFP) and (rmin) of the corresponding soluble EGFP variant determined under the same experimental conditions (eq 1): = ((0) − )/((0) − ) (1) According to eq. (1), immobilized proteins with reduced mobility (high bound rigidity) will present Mf values close to 0, while immobilized proteins highly mobile (low bound rigidity) will present Mf values closer to 1. Figure 3 shows the Mf values of soluble and immobilized EGFP variants. Mf value of His-EGFP immobilized on Ag10-Ni2+ was 1.7-fold lower than the Mf value for LSL-EGFP immobilized on Ag10 according to that His-EGFP possess a spacer arm 2.5 times shorter than the one presented by LSL-EGFP. Furthermore, the direct immobilization of EGFP core on Ag10-G throughout several covalent attachments presented Mf =0, significantly lower


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than the other two immobilized EGFP samples, which means EGFP immobilized on Ag10-G is practically immobile in the nanosecond time-range. In the light of these results, we suggest that proteins immobilized through longer spacer arms or univalent interactions are more mobile than proteins closely attached to the agarose surface by several bonds.

0.48

0.44

1.0

0.5

Mf

0.40

(r)

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0.36

0.32

0.0

Figure 3. Average anisotropy values (grey bars) (r) and mobility factors (black circles) (Mf) of different EGFP preparations. For each sample, Mf values are calculated from the corresponding experimental (r) values regarding to the minimum and maximum EGFP anisotropy values. armax was r(0)EGFP in all cases.b rmin was rEGFP. c rmin was rLSL-EGFP. Experimental correlation between thermal stability and fluorescence anisotropy of immobilized EGFP The spectroscopic and optical data herein obtained demonstrate that fluorescence anisotropy, and consequently rotational mobility of the immobilized EGFP directly depends on the manner how it is attached to the support. To additionally demonstrate that the degree of constraints to protein rotational motions is related to the thermal stability of the protein, we studied both thermal denaturation and thermal stability (Figure 4 and Figure S5) of EGFP immobilized on Ag10-G (low Mf) and LSL-EGFP immobilized on plain agarose (high Mf). We observed a temperature


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range where significant EGFP fluorescence can be measured for the system EGFP/Ag10-G but not for the system LSL-EGFP/Ag10 (Figure 4), which is consistent with a scenario where a fraction of EGFP remains properly folded in the first case whereas no folded EGFP is present in the second one. Moreover, the EGFP immobilized on Ag10-G kept 100% of its initial fluorescence after 1h incubation at 55 ºC unlike LSL-EGFP immobilized on plain Ag10 that only retained 20% of its initial fluorescence under the same conditions. Taken together, these results would suggest higher protein stability for the first system; namely, the results are consistent with a higher stability for the EGFP multivalently attached to the solid surface. Both thermal denaturation and inactivation data support that the immobilization chemistries that yield high (r) and consequently low Mf promote higher thermal stabilizations of the immobilized proteins than those chemistries driving to high Mf values. Thus, according to both spectroscopic and functional studies, the higher or lower protein mobility is determined by both the length and flexibility of the spacer arm and the multivalency of the protein-support interaction. Finally, it is worth to mention that despite the qualitative shape of the registered curves of EGFP fluorescence versus temperature is similar to those of fluorescent proteins in solution22, the intrinsic complexity of our systems, both in terms of composition and processes involved, precluded any attempt to quantitatively analyse these results.


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Relative fluorescence / %

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100 80 60 40 EGFP/Ag10-G

20

LSL-EGFP/Ag10

0 20

30

40

50

60

70

80

90

100

Temperature / ºC

Figure 4. Thermal denaturation of LSL-EGFP/Ag10 (dotted line) and EGFP/Ag10-G (solid line). Anisotropy studies guide the selection of the optimal chemistry to immobilize two different enzymes Fluorescence anisotropy studies using a two-photon polarization microscope have allowed us screening different immobilization chemistries to determine the crucial parameters that drive to protein stabilization. Spatially resolved anisotropy can therefore provide a predictable outcome of those immobilization chemistries that thermally stabilize proteins by reducing their mobility. In fact, the outcome resulted from the anisotropy studies with EGFP variants immobilized through different chemistries was used to stabilize two non-fluorescent hydrolases with industrial interest; lipase 2 from Bacillus stearothermophilus (BTL2)23 and a β-galactosidase from Lactobacillus plantarum (BGL)14. The native enzymes and their corresponding LSL-fusion proteins were immobilized on Ag10-G and plain Ag10 respectively. Figure 5 shows, as the anisotropy predicts, that immobilization on Ag10-G through lysine residues on protein surfaces resulted in more thermostable biocatalysts than chimeric enzymes immobilized on Ag10 through the lectin domain. Although protein immobilization on Ag10-G


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supports has stabilized dozens of industrially relevant enzymes12, this study sheds light on why glyoxyl chemistry is so efficient improving protein stability. It had been suggested that both the length of the spacer arm and the number of protein-support interactions strongly contributed to increase the stability of immobilized protein on porous beads12, but grounded mechanistic interpretations based on molecular and dynamics studies were not available.

Residual Activity / %

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

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100 75 50 25 0 LSL-BTL2

BTL2

LSL-BGL

BGL

Figure 5. Thermal inactivation of different immobilized enzymes immobilized on Ag 10 (blue) and Ag10-G (purple). BTL2 and BGL were incubated for 1h at 60 ºC and 45 ºC respectively and their residual activities were measured and normalized to their corresponding initial activities. Here, we have advanced in the characterization of such heterogeneous systems by elucidating two parameters that control the anisotropy of proteins and consequently their mobility regardless the protein distribution across the 3D-porous structures. The enormous value of fluorescence anisotropy in the characterization of fluorescence proteins can be expanded to enzymes strongly tethering fluorescence cofactor such as FAD+ to perform time-resolved fluorescence studies.


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CONCLUSIONS We have designed a new methodology to characterize heterogeneous bio-molecules (i.e enzymes), connecting molecular and dynamical aspects with protein function under drastic conditions. This methodology has allowed the quantification of fluorescence anisotropy across the porous structure of microbeads, up to 100 µm deep-material imaging with sub-µm spatial resolution, providing a mobility factor that informs us about the degree of constraints to the rotational motions of the attached protein promoted by the immobilization chemistry. The mobility factor represents a general protein mobility scale regardless both instrumental setup and the size of the attached protein. This factor would facilitate the comparison between immobilized proteins from other studies and/or the use of different instrumental setups. Noteworthy, the data from the time/space-resolved fluorescence anisotropy suggest that multivalent protein immobilization through extremely short spacer arms -like glyoxyl chemistry provides- promotes a highly rigid interaction between the protein and the solid surface that turns proteins into highly thermostable. This conclusion had been previously withdrawn from indirect experimental evidences based on functionality, but here we have linked conventional functional studies with advanced molecular characterization of immobilized proteins. Moreover, this platform aids in the design of better immobilization strategies to achieve more stable heterogeneous biocatalysts with interest for the biodiesel and food industries. Therefore, molecular characterization of such biocatalysts engenders an emerging opportunity to address operative aspects that occur at different scale lengths (nm to mm) during the reaction process.


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ASSOCIATED CONTENT Materials, bacteria strains, plasmids, protocols for the activation of solid supports, theoretical details are described in supporting information and some experimental methodologies and some additional experimental data are supplied as Supporting Information. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author FLG: e-mail: [email protected] MPL: e-mail: [email protected] ACKNOWLEDGMENTS We acknowledge COST action CM103-System biocatalysis and IKERBASQUE for the funding to FLG. We also acknowledge financial support for MPL, CG [grant number CTQ 20101645] and JMG (project CTQ2009-07568) from the Ministerio de Economía y Competitividad, for JMM from the Ministerio de Educación y Ciencia (BFU2010-17929/BMC). ABBREVIATIONS Ag10: Plain and cross-linked 10% agarose beads: Ag10-Ni2+: Cross-linked 10% agarose beads activated with nickel chelates. Ag10-G: Cross-linked 10% agarose beads activated with aldehyde (glyoxyl) groups. EGFP: Enhanced green fluorescent protein. LSL: lectin domain of the haemolytic toxin LSLa from the mushroom Laetiporus sulphurous. LpPDC: p-coumaric acid decarboxylase

from

Lactobacillus

plantarum.

BTL2:

lipases

2

from

Bacillus


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Table of Contents Graphic


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Two-Photon Fluorescence Anisotropy Imaging to Elucidate the

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