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Immobilization of Enzymes in Mesoporous Silica Particles. Protein Concentration and Rotational Mobility in the Pores Pegah Sadat Nabavi Zadeh, and Björn Åkerman J. Phys. Chem. B, Just Accepted Manuscript • DOI: 10.1021/acs.jpcb.7b00562 • Publication Date (Web): 03 Mar 2017 Downloaded from http://pubs.acs.org on March 12, 2017

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The Journal of Physical Chemistry

Immobilization of Enzymes in Mesoporous Silica Particles. Protein Concentration and Rotational Mobility in the Pores.

Pegah S Nabavi Zadeh, Björn Åkerman* Department of Chemistry and Chemical Engineering, Chalmers University of Technology, Kemivägen 10, SE-41296 Gothenburg, Sweden *Email: [email protected] Tel: +46317723052

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ABSTRACT Enzyme immobilization in porous silica particles is used to improve enzyme function in biocatalytic applications. Here we study the effective protein concentration and rotational mobility of lipase and bovine serum albumin in the pores, when confined in five types of mesoporous silica particles with different pore and particle size, exploiting the intrinsic UV-vis absorption and fluorescence anisotropy of the tryptophan residues. For all investigated combinations of proteins and particles the steady state anisotropy is higher than for the same protein in free solution, indicating a slower protein rotation inside the pores. The retardation is stronger in more narrow pores but the proteins can still move, and there is no dependence on particle size. The average number of proteins per particle N prot varies with particles diameter D as N prot ~D2.95±0.02 for both proteins, which is close to the scaling D3.0±0.1 for the available pore volume. This observation indicates that both proteins are distributed evenly throughout the particles and rules out that the proteins are only externally bound to the particle surface. Secondly, the concentration of protein in the pores depends on pore and protein size but not particle size, and corresponds to volume fractions in the range of 20-60%.


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INTRODUCTION In applied biocatalysis, solid supports are commonly used to immobilize the enzyme. One important application is when the enzyme is encapsulated in the pores of mesoporous silica MPS particles,1-2 in which case the particles and thus the enzymes can be separated from the product in the surrounding solution by simple centrifugation, so they can be reused.3 The chemical and mechanical stability of the silica material facilitate such mechanical handling, the solid particles are rigid compared to encapsulation systems based on lipids or proteins.4 In addition, the pores of the MPS afford a protective environment where the enzymes sometimes can tolerate elevated temperature5 and high salt concentration.3 However, protocols for MPS immobilization are commonly developed by trial and error, so a better molecular understanding of the enzyme immobilization process would be useful in applications.6 UV-vis spectroscopy has been used recently to study some fundamental aspects of the immobilization process.7-12 A protein-attached fluorescent dye has been used to monitor the immobilization process in real time.7 This approach has higher time resolution than previously used indirect methods,8 and revealed a short-lived intermediate which was suggested to be that the protein binds to the external particle surface before entering the pore.7 The pH which is experienced by the protein once inside the pores has been measured by using a pH-sensing fluorescent dye attached to the enzyme9 or the pore wall.10 The results indicate that the pores in mesoporous silica provide a slightly buffering environment.9 Circular dichroism spectroscopy has been used to monitor if the immobilization affects the protein conformation.11-12 While there is progress in the understanding how the proteins navigate the pores and which environment the enzymes encounter once inside, less is known on a molecular level regarding how the enzyme localization in the pores affects the subsequent catalytic step. The loaded amount of protein is commonly reported as gram of immobilized protein per gram of (dry) particle mass6, and the molar concentration of proteins in the (hydrated) pores is seldom 3 ACS Paragon Plus Environment


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measured even though it is well-known that the catalytic rate ultimately depends on the actual ratio between substrate and enzyme concentration. Another molecular aspect is how the proteins are distributed inside the particles. The high specific pore volume3 of MPS means the particles can accommodate a large amount of protein per gram of support material. However, this is an advantage only to the extent that the enzymes are available to the substrate when it enters the pores from the external solution, which means that enzymes far from the pore opening in large particles may be less efficient. Fluorescence microscopy has been used to image the distribution of proteins immobilized inside MPS,13 but only for particles hundreds of micrometers in size which is much larger than submicron-sized MPS particles used in biocatalytic applications. Better knowledge of how the immobilized enzymes are distributed in submicron-sized silica particles is thus needed. Finally, if the immobilizing forces are strong the association to the silica material may lock the enzyme in an unfavourable orientation with the active site facing the wall. A beneficial immobilization strategy may then be to allow the enzyme to rotate inside the pores. Rotation of immobilized proteins may seem like a contradiction in terms, but restricting the translational leakage into the surrounding solution does not necessarily prevent the proteins from rotating inside the pores. An illustrative example is when water soluble enzymes such as lipases9 or feruloyl esterases14 are encapsulated in the water-filled pores of MPS which are suspended in an organic solvent. While the hydrophobic substrate remains dissolved in the external nonpolar solution, these enzymes remain inside the particles in part due to their low solubility in the external nonaqueous phase. Therefore, the confining forces will not per se restrict protein rotation inside the waterfilled pores. Non-translational motion of proteins can be monitored by fluorescence anisotropy spectroscopy,15-16 making this method a powerful tool in biochemical research and medical analysis.17-19 Chen et al12 have reported an increase in the fluorescence anisotropy for organophosphorus hydrolase when it is immobilized in MPS with a pore


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diameter of 30 nm. This observation suggest a retardation of protein rotation in this particular case, but to our knowledge the rotational mobility of proteins in the pores of mesopourous silica particles has not been investigated systematically. Here we study two different proteins, lipase MML and bovine serum albumin BSA, immobilized in a set of five types of silica particles with different pore and particle size typically used in biocatalytic applications. We use UV-vis spectroscopy to measure the actual enzyme concentration in the MPS pores as obtained in representative immobilization protocols, and also probe how the immobilized enzymes are distributed inside the particles. Furthermore, under the same set of conditions we use fluorescence anisotropy spectroscopy to investigate to what extent the confined proteins are able to rotate inside the pores. In previous spectroscopy studies of protein immobilization7, 9 we have labelled the proteins with fluorescent dyes, but here we exploit the absorbance and fluorescence of intrinsic aromatic amino acid residues, in order to study native proteins and for the sake of comparison with the previous anisotropy study.12 The lable-free approach also avoids potential depolarization effects due to rotation of a dye attached to the protein by a flexible linker.

EXPERIMENTAL SECTION Chemicals and Particles Lipase (MML from Mucor Miehei) and albumin (BSA from bovine serum) were purchased from Sigma-Aldrich. The properties of the proteins in this study can be found in Table 1. No dye labeling was performed because the tryptophan residues in MML and BSA were used as intrinsic fluorescent reporters. If not otherwise stated, the experiments were performed in 0.1M phosphate citrate buffer at pH = 6 and 25ºC.



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Table 1. Properties of the Proteins

protein a

MML BSA

Mw b (kD)

RH c (nm)

pI d

32 66

2.25 3.5

3.8 4.7

ε 280 e (M cm )

(ns)

θ prot free g

42800 43824

3.5 6.3

23.6 88.8

-1

τ (Trp) f

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-1

(ns)

a

MML-Mucor Miehei Lipase, BSA-Bovine Serum Albumin

b

Molecular weight20-21

c

Hydrodynamic radius8-9

dIsoelectric point20-21 eExtinction coefficient8-9 f

Average excited state lifetime of tryptophan residues22-23

gGlobal rotation correlation time for free protein estimated by Eq.2 using the hydrodynamic radius

Mesoporous silica (MPS) particles of types SBA-15 (Santa Barbara Amorphous-15) and HMM (Hiroshima Mesoporous Materials) were a gift of Hanna Gustafsson (Applied chemistry, Chalmers) and were synthesized as described previously.8, 24 Nitrogen adsorption was used for characterization of the MPS to obtain average pore diameter, specific pore volume and BET surface area. Four types of mesoporous silica particles with the same pore radius and different particle diameter were used, as well as two types of particles with the same particle diameter but different pore size. The properties of the used MPS are summarized in Table 2. Table 2. Properties of the MPS Particles MPS-D a type b BJH pore sizec (nm)

BET surface aread (m2/g)

Vpore e (cm3/g)

θ part f (s)

MPS-40

HMM

9.1

463

0.91

8.5⋅10-6

MPS-300

SBA-15

9.4

606

1.03

3.5⋅10-3

MPS-1000

SBA-15

9.3

502

1.18

1.2⋅10-1

MPS-2000

SBA-15

8.9

554

1.17

1.1

MPS-2000

SBA-15

6.0

986

1.08

1.1

a

MPS-D: Mesoporous silica particles, where D refers to the average particle diameter in nm8, 24

b

Type of MPS preparation. HMM: Hiroshima Mesoporous Materials, SBA-15: Santa Barbara Amorphous8, 24

cAverage pore diameter obtained by the Barret-Joyner-Halenda method8, 24 d

Specific pore surface area obtained by the Brunauter-Emmett-Teller method8, 24

eSpecific pore volume obtained by the nitrogen adsorption-desorption isotherms8, 24 fEstimated rotational correlation time for particles calculated by Eq.2 for a spherical particle, using the average

particle radius R part = D/2.


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The morphology of the particles were investigated by SEM (See Figure 1). It is seen that the MPS-40 particles (Fig.1a) are essentially spherical, MPS-300 (Fig.1b) and MPS-2000 (Fig.1d) rod-like, whereas the MPS-1000 particles (Fig.1c) are disc-like in shape.

Figure 1. Morphology of the MPS particles by SEM imaging. (a) MPS-40, spherical particles with slit-shaped pores without an ordered structure; (b) MPS-300, rod-shaped with hexagonally ordered uniform pores; (c) MPS1000, disc-like with hexagonally ordered pores; (d) MPS-2000, rod-shaped with hexagonally ordered pores.8, 24

Protein Immobilization Aqueous solutions of the MPS were prepared by dispersing 5 mg of dry mesoporous silica particles in 1ml phosphate-citrate buffer, using vortexing for 10 min at 10 rpm followed by sonication (Ultrasonic cleaner model CD-4800 at a power of 70 W) for 20 min in order to dissolve any particle aggregates, and a final step of vortexing for 5min. Protein-particle samples were prepared by mixing 20µl of MML or BSA stock solution (20mg/ml in phosphate-citrate buffer) with 200 µl of MPS solutions (5mg/ml) diluted to a final volume of 500 µl with phosphate-citrate buffer. Each sample contained a total amount of 400 µg enzyme/mg MPS. Reference samples of free protein were prepared in the same way, but 7 ACS Paragon Plus Environment


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replacing the 200 µl MPS solution with buffer. In the latter case the final protein concentration 0.8mg/ml corresponds to a volume fraction of about 0.1%, whereas the accumulation in the particle pores leads to much higher local volume fractions of 20-60% (See RESULTS). The samples were incubated at 25ºC for 48 h during gentle stirring, and then centrifuged for 6 min. The pelleted protein-particles complexes were re-suspended, and washed three times with 500 µl of phosphate citrate buffer by repeated centrifugation and resuspension. The purified MPS particles with the immobilized proteins were finally re-suspended by adding 100 µl of buffer and vortexing for a few minute until homogenous samples were obtained for the spectroscopic measurements. The fraction of the added protein which was associated with the particles is referred to as the degree of immobilization (DOI). This fraction was calculated from the difference between the total added amount of enzyme and the amount of enzyme remaining in the supernatant after particle washing, which was measured by UV absorption at 280 nm using a Varian Cary50 spectrophotometer. The protein loading PLD (µg protein/mg particle) was then obtained from the dry mass of added particles. The amount of immobilized protein was also expressed as the number of proteins per particle (N prot ) as well as the volume fraction of protein in the pores, pore filling (P f ). Both were calculated from the PLD value as described in section S1 in Supporting Information.

Fluorescence Spectroscopy Experimental. Steady state polarized emission spectra were recorded on a SPEX fluorolog 3 spectrofluorometer (JY Horiba) using Glan polarizers with excitation at 280 nm and emission recorded in the wavelength range from 300 to 450 nm. The proteins studied here contain both tryptophan and tyrosine, but the tryptophan residues are the dominant source of absorption and emission in proteins in the near UV-region, and when excited at 280 nm the emission maximum of tryptophan is at 350 nm while tyrosine emission is at 300 nm and more narrowly 8 ACS Paragon Plus Environment

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distributed.25 For simplicity we will therefore refer to the mixed set of involved chromophores as tryptophan. The polarized emission spectra were corrected for particle light scattering by subtracting a spectrum of a protein-free particle sample with the same setting of the polarizers. (See section S2 of Supporting Information). Scattering contributions were further minimized by using an excess of added enzyme (400 µg enzyme/mg MPS) in order to ensure high protein loading in the particles. The particles were washed three times with protein-free buffer before the spectroscopic measurements in order to remove free and/or loosely bound proteins, and leakage of proteins during the spectroscopic measurements were monitored and corrected for by measuring free enzyme in the supernatant obtained by centrifugation after the spectroscopic measurements. The presented anisotropy values were taken as the average value in the wavelength range 335-355 nm, with the stated uncertainty calculated as half the maximal variation between 3 or 4 independent experiments which reflects the reproducibility of the experiments. Theory. The fluorescence anisotropy of a fluorophore is a measure of its rotational mobility on the timescale of the fluorescence lifetime. For a protein with a single fluorophore with lifetime

τ, the rotational correlation time θ is related to the steady state anisotropy (r) through the Perrin equation15, 25 r=

𝑟𝑟0

1+

Eq.1

𝜏𝜏 𝜃𝜃

where r o is the fundamental anisotropy of the fluorophore in the absence of rotational motion, which for the tryptophan chromophore is r o = 0.3.15, 25 The lifetime value is sensitive to the environment in general, so τ differs slightly between the two proteins studied here (see Table 1). We neglect the effect of the immobilization on the lifetime (i.e. use the τ-value of free protein) because Czeslik et al report16 that the effect is small at least when proteins are permanently adsorbed to a silica surface. For a spherical particle with radius R, the rotation correlation time (θ) in Eq.1 is given by 9 ACS Paragon Plus Environment


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θ=

4 3

𝜂𝜂 𝜋𝜋𝑅𝑅 3

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Eq.2

𝑘𝑘 𝑇𝑇

where η is the solution viscosity, k is Boltzmann constant and T the temperature. In the case of a rigidly attached fluorophore, the radius R calculated from the anisotropy measured for (free) protein using Eq.1 and Eq.2 is expected to be close to the hydrodynamic radius (R H ) obtained by direct hydrodynamic methods such as dynamic light scattering. However, it is well established that in the case of tryptophan residues in proteins, the fluorophore undergoes a local segmental motion relative to the protein matrix as a whole, which contributes to the depolarization in addition to overall protein rotation.26-27 Applying the Perrin Eq.1 to the measured steady state anisotropy then gives apparent rotational correlation time (θ A ), and the corresponding apparent radius (R A ) of the protein obtained from Eq.2 may then be smaller than the hydrodynamic radius because a fraction of the total anisotropy is lost due to the segmental motion of the tryptophan fluorophore itself.26-27 The correlation time θ prot for the overall (global) protein rotation can be estimated by using the known hydrodynamic radius R H of the protein (Table 1) in a modified version of Eq.2

θ prot = f

4 3

𝜂𝜂 𝜋𝜋𝑅𝑅𝐻𝐻 3

Eq.3

𝑘𝑘 𝑇𝑇

where f is a factor depending on the shape and hydration of the protein, assuming the value f = 2 for globular proteins.27-28

RESULTS The amount of immobilized protein (protein loading) of lipase MML and albumin BSA were measured, and the corresponding protein concentrations in terms of volume fraction inside the pores of five different types of mesoporous silica particles (Table 2) were calculated. Secondly, we use steady state fluorescence anisotropy to monitor the rotational dynamics of the confined proteins and compare with the rotation rates of the same proteins in free solution. The two sets 10 ACS Paragon Plus Environment

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of measurements are connected because the actual protein concentrations in the pores may affect the rotational motion of the immobilized proteins compared to the dilute protein samples which are used as particle-free references. The two types of MPS-2000 particles in Table 2 were used to study the effect of pore size, whereas the four MPS-samples with approximately the same pore radius (4.5 – 4.7 nm) were used to investigate the effect of particle size for a given pore size. The latter set will be referred to as the 4.6 nm pore particles for simplicity.

Protein Loading and Pore Filling The amount of immobilized proteins in porous particles is commonly expressed as the mass of bound protein per gram of (dry) particles, referred to as the protein loading (PLD ).6 Table 3 shows the measured P LD -values obtained for all the combinations of proteins and particles in this study, as well as the degree of immobilization DOI (the fraction of the added proteins which remain bound to the particles after washing) which were used to calculate the PLD . It is seen from Table 3 that the DOI-values (and hence P LD ) are essentially constant for the four particle types with 4.6 nm pores. That the particle size tends to have little influence on protein loading has been noted before8, 24 although the amount of loaded protein obtained here is higher, probably because the added amount of protein per particle is four times higher. Next, when comparing the two types of MPS-2000 particles, the degree of immobilization and protein loading is considerably lower for the smaller pore size (3nm), especially for the larger BSA where P LD is 2.2 times lower than in the 4.6 nm pores for the same particle size. In the case of MML the lower loading is in agreement with a previous study with the MPS-2000 particles8 and consistent with that less space is available inside the MPS pores with the smaller diameter. However, the comparison is somewhat obscured due to the variation in specific pore volume (see Table 2) between the different particle types. Therefore, the amount of immobilized protein was expressed in two more direct measures. First, the actual protein concentration in the pores



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1 2 3 which is given as the volume fraction protein (pore filling P f ), and secondly the average number 4 5 of proteins per particle N prot (see section S1 of Supporting Information). 6 7 8 Table 3. Protein Loading, Pore Filling and Average Number of Proteins per Particle 9 MML BSA MML/BSA 10 11 particle R pore b DOIc P LD d Pf e N prot f DOIc P LD d Pf e N prot f N prot -ratio g 12 a type (nm) (%) (µg/mg) (%) (µg/mg) 13 4.5 72 288 0.284 133 84 336 0.604 75 1.77 14 MPS-40 15 4.7 75 300 0.261 5.4⋅104 87 348 0.553 1.78 MPS-300 3.0⋅104 16 17 MPS-1000 4.6 70 280 0.213 1.7⋅106 85 340 0.472 1.70 9.9⋅105 18 19 MPS-2000 4.5 71 284 0.218 1.4⋅107 84 336 0.470 1.74 7.9⋅106 20 3.0 63 252 0.209 1.3⋅107 38 152 0.230 3.42 21 MPS-2000 3.8⋅106 22 a MPS-D, where D indicates average particle diameter in nm (see Table 2) 23 b 24 Average pore radius8, 24 25 cDegree of immobilization, fraction of added proteins which were immobilized. 26 dProtein loading (µg protein per mg of dry particles) 27 28 ePore filling, see section S1 of Supporting Information 29 f Average number of proteins per particle, see section S1 of Supporting Information 30 g 31 Ratio of number of MML and BSA proteins per particle 32 33 34 Table 3 shows the pore filling values which are calculated from the P LD -values. It is seen that 35 36 37 for the particles with the larger pores, MML generally fills a significantly smaller fraction of 38 39 the available pore volume (P f = 20-30%) than the larger BSA (Pf = 50-60%). By contrast, in 40 41 the case of the smaller pores both proteins occupy a similar fraction of the pore volume, 2142 43 44 23%. Notably, in all cases studied here the protein concentration in the pores is much higher 45 46 than the starting volume fraction of about 0.1% in the protein solutions in which the empty 47 48 49 particles were incubated. 50 51 It should be noted that pore filling reported here only gives the average protein concentration 52 53 in the pores, it may also be important how the proteins are distributed throughout the particles. 54 55 56 Table 3 also shows the average number of bound proteins per particle (N prot ) calculated from 57 58 the PLD -values. It is seen that for the smallest MPS-40 there is only about 100 proteins per 59 60

particle on the average, and that N prot increases strongly with increasing particle size. In the 12 ACS Paragon Plus Environment

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Discussion we investigate to which extent this increase matches the increase in available pore volume per particle. It is also seen that the ratio between the numbers of MML and BSA immobilized in a given type of particle is roughly constant at about 1.75 for the large pore size, and about twice as high for the smaller pores where the ratio is 3.42.

Fluorescence Anisotropy The red curves in Figure 2 shows the anisotropy spectra (r) for BSA either in free solution (Fig. 2a) or immobilized in MPS-1000 (Fig. 2b), calculated by applying Eq.S4 (section S2 of Supporting Information) to the polarized component spectra in Figure S1. The black curves in Figure 2 show the corresponding total intensity spectra (I tot ) obtained by Eq.S5.

Figure 2. Spectra for anisotropy r and total intensity I tot for (a) free BSA and (b) BSA immobilized in MPS-1000 with 9.3 nm pore size, calculated from the corresponding polarized components in Figure S1 using Eq.S4 and Eq.S5 in section S2 of Supporting Information. Solid horizontal black line represents the presented anisotropy values, taken as the average between 335 nm and 355 nm. Excitation at 280nm.

The overall shape of the total intensity spectrum is the same for free and immobilized protein (including an emission peak at 335nm). This observation indicates that the tryptophan reporter chromophore is not strongly perturbed by the pore environment and supports our assumption that the life time of the free protein can be used for the immobilized proteins (See Experimental section). The major spectral difference is that the total intensity in the particle-containing sample (Figure 2b) increases more strongly below 300 nm compared to the particle-free sample (Figure 2a), mainly due to enhanced light scattering of the excitation light (280 nm) by the 13 ACS Paragon Plus Environment


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particles. Turning to the anisotropy it is seen to be essentially independent of wavelength until a marked increase arises below 325 nm, especially in the particle-containing sample in part due to the light scattered by the MPS being highly polarized. The anisotropy values presented below (Table 4) were taken as the average value in the range 335-355nm, representing the limiting value for long wavelengths in the emission band of tryptophan. Finally, it is seen in Figure 2 that the fluorescence anisotropy of BSA is higher in the presence of particles, an effect which is observed for all particle/protein combinations studied here.

Effect of Pore and Particle Size In Table 4 the measured steady state anisotropy of free protein is compared with the same protein when immobilized, either in two types of MPS-2000 particles with different average pore sizes or in four types of particles with varying diameter but essentially the same pore radius (4.6±0.1 nm). The first observation is that in all five cases both MML and BSA exhibit an anisotropy in the presence of the particles which is significantly higher compared to the same enzyme in free solution, given the uncertainty ±0.01 in the r-values. This observation indicates a slower protein rotation in the pores, but possibly also slower internal protein dynamics which also may affect the anisotropy (See DISCUSSION). Secondly, for a given protein type the anisotropy in the MPS-2000 particles is higher in the smaller pores. Thirdly, there is no obvious trend in the anisotropy values with particle size. The highest values are observed in the intermediately sized MPS-300 where the anisotropy is equal or close to 0.3. Finally, we note that in our hands free MML has a lower anisotropy than free BSA, consistent with that the smaller MML rotates faster in free solution than BSA and in agreement with well-established results.25, 29


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Table 4. Fluorescence Anisotropy for Free Proteins and in Different Particle Types

MML

BSA

0.06

θA e (ns ) 0.87

0.08

θA e (ns ) 2.73

4.5

0.23

11.5

0.18

9.45

MPS-300

4.7

0.30

-----

0.27

56.70

MPS-1000

4.6

0.15

3.50

0.16

7.24

MPS-2000

4.5

0.17

4.58

0.18

9.45

MPS-2000

3.0

0.21

8.17

0.24

25.20

particle type a

R pore c (nm) ---

MPS-40

Free proteinb

rd

rd

a

MPS-D where D is the average particle diameter in nm.

b

Protein in particle-free buffer solution.

c

Average pore radius8, 24

dMeasured tryptophan steady state anisotropy. The uncertainty is ±0.01 as calculated from

the variation between 3 to 4 independent experiments. eApparent rotational correlation time calculated from anisotropy by Eq.1. The uncertainty is ±0.2ns

from the variation between 3 to 4 independent experiments.

DISCUSSION The pore filling and anisotropy results are discussed first in terms of the particle properties, including possible mechanisms for the enhanced protein anisotropy in the presence of the particles. Finally some implications for the efficiency of encapsulated enzymes are discussed.

Particle Properties The particles used here constitute a series of porous hosts which is suitable for a systematic investigation of protein immobilization. They form a nearly homologous set regarding the nanometer scale pore properties in spite of their morphological differences on the micrometer length scale (Figure 1). In a set of particles with effective diameters of 40, 300, 1000 and 2000 nm and essentially constant pore radius 4.6±0.1 nm (See Table 2) the average pore volume per particle exhibits the expected cubic dependence Vpore ~ D3.0±0.1 on the effective particle size D with good accuracy (See Figure S2 in Supporting Information). This set will for simplicity be



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referred to as 4.6 nm pore particles, in contrast to the smaller 3.0 nm pores which are also available in the largest 2000 nm particles.

Amount of Immobilized Protein The present investigation constitute a systematic study of how protein size as well as pore and particle size affect the degree of pore occupation by proteins immobilized in mesoporous silica particles. That the degree of immobilization (DOI in Table 3) is nearly constant and well below 100% for the set of particles with 4.6 nm pores indicates that the added protein is in excess, and that these particles are essentially saturated with protein for both BSA and MML under our conditions. Consistent with this interpretation the DOI-values are even smaller with the smaller 3.0 nm pores, although the comparison is somewhat obscured by the fact the specific pore volume varies between the particle types. An important conclusion from the present work is that the amount of bound protein is better understood in terms of the number of proteins per particle (N prot ) and the actual concentration of proteins in the pores (pore filling Pf ), two measures that will be discussed below. Proteins per particle. Figure 3 shows a plot of the average number of proteins (N prot ) per particle (from Table 3) plotted versus particle diameter (D), and the strong increase shows that the larger particles accommodate many more proteins as expected. Importantly, the inset in Figure 3 shows that the number of protein per particle with the large pore radius 4.6 nm varies with the linear particle size D as N prot ∼D2.94±0.01 for MML and Nprot ∼D2.95±0.01 for BSA.


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Figure 3. The average number of proteins per particle for MML (squares) and BSA (circles) versus the effective particle diameter D, for pore radius 4.6±0.1 nm (solid symbols; curves are guide to eye) and 3.0nm (open). Inset: Double logarithmic plot for data with 4.6 nm pore radius where straight lines are least square linear fits giving slopes 2.94±0.02 for MML and 2.95±0.01 for BSA.

The near cubic scaling (i.e proportionality to particle volume) supports that under our immobilization conditions both types of proteins are more or less evenly distributed inside the particles with the 4.6 nm pores, since the pore volume per particle exhibits a similar scaling (~ D3.0±0.1 Figure S2 in section S3 of Supporting Information). Notably, the particle-size dependence rules out that the proteins are adsorbed only to the external particle surface because then the expected scaling is Nprot ∼D2. Turning to Nprot in the case of the MPS-2000 particles with the smaller 3 nm pores, it is clear from Figure 3 (open symbols) that the number of proteins per particle is reduced compared to the same 2000 nm particle size with the larger pores (solid symbols), and more so for the larger BSA. Pore filling. The pore filling (Table 3) is another useful measure of how much protein is bound to the particles. Firstly, the observed Pf -values (20-60%) in general are much higher than the volume fraction in the protein solution before the particles are added (0.1%), which quantifies how pore and protein size affect the capacity of porous silica particles to accumulate a protein from a dilute aqueous solution. In fact, the highest volume fractions measured here (0.6) are



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approaching the theoretical upper limit for close packing of spheres in a cylindrical pore (0.67),6, 30 which supports our assertion that the particles are nearly saturated with protein. Secondly, the high loading of both proteins shows that a pore size of around 4.6 nm is suitable for immobilization of proteins in the present size range, probably because the radii of MML and BSA are smaller than but comparable to the pore size. It has been reported previously that the pore size is important for enzymatic activity, and Rpore = 4.4 nm was found to give both high loading and high enzymatic activity for MML.8 Thirdly, the high protein concentrations reveal a crowded environment in the pores which is taken into account when the anisotropy values of immobilized and free protein are compared below. By combining the two measures for the amount of bound protein, the Pf -values can be well understood in terms of the data on Nprot. From Table 3 it is seen that the number of MML per particle is a factor of 1.75±0.03 higher than the number of BSA in the four particle types with the 4.6 nm pores, explaining the higher pore filling with BSA. The volume of each BSA is 3.8 times larger compared to the MML molecule and therefore more efficient (per protein) at filling the pores. In a deviation from this overall pattern, BSA has a pore filling of only 23% in the MPS-2000 particles with the smaller 3.0 nm pores and essentially as low as for MML (21%). This effect is due to that the number of MML per particle is now about a factor 3.4 higher than for BSA, almost exactly compensating for the smaller volume per MML protein.

Anisotropy The increase in anisotropy for immobilized proteins compared to the same protein free in solution (Table 4) shows that the motions causing the depolarization are retarded in the pores. When anisotropy is monitored by tryptophan fluorescence, it reflects the rotational motion of the whole protein, but also the local motion of the tryptophan residues.26, 31 It is important to note that in the steady state anisotropy technique employed here these two contributions are convoluted into an apparent rotational correlation time (θA). The fact that θA is significantly 18 ACS Paragon Plus Environment

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larger in the pores than for free protein (Table 4) just reflects that one of those motions or both are retarded in the pores. Notably, we show in section S4 of Supporting Information that the potential depolarizing effect of whole-particle rotation is neglible in the present study, because all studied particles are large enough to be viewed as non-rotating on the time scale of the tryptophan fluorescence lifetime. The maximum anisotropy in our system is r = ro = 0.3,25 the fundamental anisotropy for tryptophan, which is obtained for a protein which does not rotate in the pores (for instance due to pore wall adsorption) and does not exhibit any local tryptophan depolarizing motion. All the observed r-values (Table 4) are intermediate between the limiting anisotropy (0.3) for fully adsorbed and rigid protein and the anisotropy value of respective protein when free in solution. This observation strongly indicates that both proteins do undergo some depolarizing motions inside the pores, and also that these motions are retarded compared to the free protein. This interpretation is supported by the observation that the anisotropy is higher in the MPS-2000 with more narrow pores (3.0 nm) for a given protein, as expected since stronger confinement is likely to affect the motion more strongly. There is no trend in the anisotropy values with particle size (for 4.6nm pore size) in Table 4, which is consistent with that none of the MPS particles studied here undergo any depolarizing rotation during the tryptophan fluorescence lifetime. Therefore, particle rotation is not the cause for the difference in anisotropy between different particle sizes (for a given protein), but more likely due to different pore morphology or possibly to differences in protein-particle interaction strength. The anisotropy of MML in MPS-300 is equal to limiting anisotropy value ro = 0.3 which indicates non-rotating proteins with negligible depolarizing internal protein motions (within our present experimental accuracy). Gustafsson et al24 have noted a higher activity of lipase in this particular type of particles and an interesting possibility is that an essentially static encapsulation of the enzyme affects its activity. For BSA in the MPS-300 particles the



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anisotropy value (r) is 0.27. In the other three particle types (Table 4), the measured anisotropy of both proteins is in an intermediate range (0.15-0.27) which shows that depolarizing motions occur albeit more slowly than for free protein.

Mechanisms of Enhanced Protein Anisotropy in the Pores Inside the pores the protein dynamics can be retarded by two principle mechanisms, proteinprotein or protein-wall interactions. The first scenario invokes the high protein concentration in the pores that we report (Table 4). The retardation could then be caused by the same proteinprotein hydrodynamic interactions in the pores that has been demonstrated by Roosen-Runge et al 32 to retard BSA motion in non-dilute solutions of free proteins (no particles). However, a depolarized light scattering study of the rotational motion of spherical particles in concentrated suspensions33 indicates that such effects depend too weakly on the volume fraction to explain the retardation we observe. Also speaking against such a protein-protein explanation is the observation that both MML and BSA exhibit higher anisotropy in the more narrow pores (Table 4) even though the pore filling is lower (Table 3). The second possible mechanism, which indeed is expected to be stronger in more narrow pores, is that the rotation is retarded by hydrodynamic interactions between the (individual) proteins and the pore wall. Rotation of confined particles has been studied theoretically in detail by Jones.34 A quantitative comparison with the theory is precluded by the fact that Jones models the rotation of a (single) spherical particle confined in a slit between two planar walls, in contrast to our essentially cylindrical pore geometry. However, in qualitative terms, the predicted retardation is stronger the narrower the gap, in agreement with that the anisotropy for both MML and BSA is higher in the more narrow pores (Table 4). Moreover,the predicted effect is of the right order of magnitude. When the slit is twice as wide as the particle diameter the prediction by Jones34 corresponds to an increase in the rotational correlation time (θrot by a factor of about 1.7 compared to the same particle when rotating in free solution. This is 20 ACS Paragon Plus Environment

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comparable to the increase in θA by a factor of 0.17/0.06 = 2.8 that we observe in the case for MML (Table 1, RH = 2.25 nm) in the 4.6nm pores (Table 3), in which case the relative pore width is D/2RH = 2. The hydrodynamic rotational retardation of a sphere in a cylindrical pore has not been investigated theoretically to our knowledge, but is expected to be somewhat stronger than in a slit because the particle is more strongly confined than by two planar walls. This presumption that more walls will cause stronger retardation is supported by a theoretical study of the rotational retardation of a sphere in a spherical cavity.35 It is important to note that the comparison with one-particle theories regarding particle-wall interactions rests on the assumption that the proteins are evenly distributed in the pores, an issue on which we have presently no experimental data.

Implications for Potential Encapsulation-Effects on the Catalytic Step The amount of immobilized protein is commonly stated as the protein loading (weight of protein per weight of dry particles) because it is convenient when describing the sample preparation protocol, but on a molecular level our results show that the effective number concentration of protein (pore filling, Pf ) is very high under typical immobilization conditions, sometimes approaching close packing. Such crowded pores are likely to retard the transport of the substrate inside the pores. Notably, our results indicate that the enzymes are distributed essentially evenly throughout the particles. From a design point of view it is then probably better to immobilize a certain amount of protein in a higher number of particles with smaller size, because substrate diffusion can be expected to be slow to reach the enzymes at the bottom of long narrow pores. In addition to average concentration of the proteins and their distribution inside the particles, the confinement may also influence the catalytic efficiency of an enzyme by affecting the dynamics of the proteins. One suggested possibility is through altering the internal dynamics of the enzyme,27, 31, 36 but we argue that also the global rotation of the whole enzyme may be 21 ACS Paragon Plus Environment


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important. In fact, for water soluble lipase9 or feruloyl esterase14 the possibility to rotate inside the pores may be crucial, so that (at least intermittentently) the active site is oriented towards the substrate which is dissolved in the external nonaqueous solution. The present results were obtained with the particles in aqueous solution. However, our conclusions regarding the rotation of lipase in the pores most likely also apply with the MPS suspended in a organic solvent, since we have recently used fluorescence spectroscopy of dye-labelled proteins to support the notion that the water-soluble lipase remain localized in water-filled pores when a non-aqueous solvent surrounds the particles.9

CONCLUSION Our results indicate that tryptophan UV-absorption and fluorescence anisotropy of non-labelled proteins can be used to measure the protein concentration in the pores and study the rotation of the confined enzymes in a manner similar to free proteins. Our experimental approach lends support from the observation that the experimental values for pore filling are within the theoretically expected range (0-0.67) and the same holds for the anisotropy where the r-values should fall between the free protein anisotropy and the limiting anisotropy (r0 = 0.30) as long as the particles are so large their rotation can be neglected. The protein rotation is slower in the pores than outside the particle, with protein-wall rather than protein-protein hydrodynamic interactions possibly being the more likely mechanism. Notably, a slowing down of internal protein dynamics may contribute to the increase in steady state anisotropy measured here upon immobilization. Czeslik et al16 have studied lysozyme and SNase bound to a planar silica/water interface using time resolved anisotropy measurments. The results show that a slow component corresponding to global protein motion (θprot) is the dominant contribution, but also that the correlation time for the internal motion (θTrp) can both decrease or increase after adsorption. Time-resolved anisotropy would be an interesting


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extension of the present study to investigate the effects of the immobilization on the local residue motion in the MML and potential effects on its catalytic activity.

AUTHOR CONTRIBUTION PN as first author designed and performed the experiments, analyzed the data and wrote the manuscript, BÅ as senior investigator has contributed to designing the experiments, analyzing the data and writing the manuscript.

SUPPORTING INFROMATION Calculation of pore filling and number of proteins per particle, polarized fluorescence emission spectra in the absence and presence of particles, calculation of average pore volume per particle vs particle size, rotation of the whole MPS particle (θPart) and its contribution in anisotropy.

ACKNOWLEDGEMENTS

We would like to thank Hanna Gustafsson for synthesis and characterization of all the MPS particles, Nils Carlsson for all valuable inputs and Sam Safaei Moghaddam for helpful discussion and skilfull assistance. Funding from the Swedish research council to B.Å is acknowledeged as a part of the Linnaeus Centre for Bio-inspired Supramolecular Function and Design – SUPRA.



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21. Hirayama, K.; Akashi, S.; Furuya, M.; Fukuhara, K., Rapid Confirmation and Revision of the Primary Structure of Bovine Serum Albumin by Esims and Frit-Fab Lc/Ms. Biochem Biophys Res Commun 1990, 173 (2), 639-46. 22. Graupner, M.; Haalck, L.; Spener, F.; Lindner, H.; Glatter, O.; Paltauf, F.; Hermetter, A., Molecular Dynamics of Microbial Lipases as Determined from Their Intrinsic Tryptophan Fluorescence. Biophys. J . 1999, 77 (1), 493-504. 23. Joshi, N. V.; Joshi, V. O. d.; Contreras, S.; Gil, H.; Medina, H.; Siemiarczuk, A. In Fluorescence Lifetime Measurements of Native and Glycated Human Serum Albumin and Bovine Serum Albumin, Advances in Fluorescence Sensing Technology IV, 1999; pp 124-131. 24. Gustafsson, H.; Johansson, E. M.; Barrabino, A.; Oden, M.; Holmberg, K., Immobilization of Lipase from Mucor Miehei and Rhizopus Oryzae into Mesoporous Silica--the Effect of Varied Particle Size and Morphology. Colloids Surf. B 2012, 100 (0), 22-30. 25. Lakowicz, J. R., Principles of Fluorescence Spectroscopy. 3rd ed.; Springer: Baltimor, USA, 2006. 26. Lakowicz, J. R.; Freshwater, G.; Weber, G., Nanosecond Segmental Mobilities of Tryptophan Residues in Proteins Observed by Lifetime-Resolved Fluorescence Anisotropies. Biophys J 1980, 32 (1), 591-601. 27. Lakowicz, J. R.; Maliwal, B. P.; Cherek, H.; Balter, A., Rotational Freedom of Tryptophan Residues in Proteins and Peptides. ACS. Biochem 1983, 22 (8), 1741-52. 28. B. Massey, J.; Churchich, J. E., Nanosecond Spectroscopy of a Dimeric Enzyme: Plasma Amine Oxidase. Biophys. Chem. 1979, 9 (2), 157-162. 29. Puri, B. R.; Sharma, L. R.; Pathania, M. S., Principles of Physical Chemistry. Vishal Publishing Company: 2008. 30. Miyahara, M.; Vinu, A.; Ariga, K., Adsorption Myoglobin over Mesoporous Silica Molecular Sieves: Pore Size Effect and Pore-Filling Model. Mat.Sci. Eng. C-Bio S. 2007, 27 (2), 232-236. 31. Nishimoto, E.; Yamashita, S.; Szabo, A. G.; Imoto, T., Internal Motion of Lysozyme Studied by Time-Resolved Fluorescence Depolarization of Tryptophan Residues. ACS Biochem 1998, 37 (16), 5599-607. 32. Roosen-Runge, F.; Hennig, M.; Zhang, F.; Jacobs, R. M.; Sztucki, M.; Schober, H.; Seydel, T.; Schreiber, F., Protein Self-Diffusion in Crowded Solutions. Proc Natl Acad Sci U S A 2011, 108 (29), 11815-20. 33. Degiorgio, V.; Piazza, R.; Jones, R. B., Rotational Diffusion in Concentrated Colloidal Dispersions of Hard Spheres. Physical Review E 1995, 52 (3), 2707-2717. 34. Jones, R. B., Rotational Diffusion of Colloidal Particles near Confining Walls. J Chem Phys 2005, 123 (16), 164705. 35. Jones, R. B., Dynamics of a Colloid in a Spherical Cavity. In Theoretical Methods for Micro Scale Viscous Flows Feuillebois, F. S., A. , Ed. Transworld Research Network: Kerala, India, 2009; pp 61-104. 36. Careri, G.; Fasella, P.; Gratton, E., Enzyme Dynamics: The Statistical Physics Approach. Annu Rev Biophys Bioeng 1979, 8 (1), 69-97.



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TOC Graphic


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Figure1. Morphology of the MPS particles by SEM imaging 101x68mm (600 x 600 DPI)

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Figure 2a. . Spectra for anisotropy r and total intensity for free BSA 79x61mm (300 x 300 DPI)


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Figure 2b. Spectra for anisotropy r and total intensity for immobilized BSA 79x61mm (300 x 300 DPI)



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Figure 3. The average number of proteins per particle for the proteins 91x72mm (300 x 300 DPI)


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TOC Graphic 228x149mm (300 x 300 DPI)


Immobilization of Enzymes in Mesoporous Silica ... - ACS Publications

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