Immobilization of proteolytic enzymes on replica-molded thiol-ene micropillar reactors via thiol-gold interaction
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We introduce rapid replica molding of ordered, high-aspect-ratio, thiol-ene micropillar arrays for implementation of microfluidic immobilized enzyme reactors (IMERs). By exploiting the abundance of free surface thiols of off-stoichiometric thiol-ene compositions, we were able to functionalize the native thiol-ene micropillars with gold nanoparticles (GNPs) and these with proteolytic α-chymotrypsin (CHT) via thiol-gold interaction. The micropillar arrays were replicated via PDMS soft lithography, which facilitated thiol-ene curing without the photoinitiators, and thus straightforward bonding and good control over the surface chemistry (number of free surface thiols). The specificity of thiol-gold interaction was demonstrated over allyl-rich thiol-ene surfaces and the robustness of the CHT-IMERs at different flow rates and reaction temperatures using bradykinin hydrolysis as the model reaction. The product conversion rate was shown to increase as a function of decreasing flow rate (increasing residence time) and upon heating of the IMER to physiological temperature. Owing to the effective enzyme immobilization onto the micropillar array by GNPs, no further purification of the reaction solution was required prior to mass spectrometric detection of the bradykinin hydrolysis products and no clogging problems, commonly associated with conventional capillary packings, were observed. The activity of the IMER remained stable for at least 1.5 h (continuous use), suggesting that the developed protocol may provide a robust, new approach to implementation of IMER technology for proteomics research.
KeywordsThiol-enes Microreactors Microfluidics Enzyme immobilization Gold nanoparticles Mass spectrometry
Immobilization of proteolytic enzymes (e.g., trypsin, pepsin, and chymotrypsin) on solid support structures packed in capillary channels has gained considerable interest owing to its many benefits over soluble enzyme reactions [1, 2]. Enzyme immobilization on solid support structures omits the need for separation of the enzymes from the reaction solution, which not only simplifies the purification of the reaction products but also allows, for instance, online mass spectrometric (MS) analysis of the proteolytic digest and the reuse of the enzymes, which both result in significant savings in time and costs [3, 4]. Most importantly, enzyme immobilization effectively suppresses autoproteolysis even at high enzyme-to-substrate ratios. In some cases, immobilization also enhances the enzyme stability and activity, which leads to increased conversion rates [5, 6]. However, the enzyme immobilization strategy plays a key role in achieving the high conversion rates with a view to maximizing the amount of bound enzymes and maintaining them active on solid supports . The most common strategies make use of porous polymer monoliths [3, 4] or magnetic beads , which are functionalized with covalently bound enzymes and packed in a capillary channel to increase the surface area. Apart from classical esterification, covalent binding may however require harsh conditions that increase the risk of denaturation and loss of enzyme activity. The ester bond, on the other hand, is relatively unstable in aqueous conditions, which may result in leaching of the immobilized enzymes. Alternatively to covalent binding, enzymes have also been physically entrapped inside porous matrixes, but depending on the pore size, these are also prone to enzyme leaching (too large pores) or restricted diffusion of the substrate to the enzyme (small pores) . Any kinds of capillary packing (whether porous matrices or magnetic beads) may also suffer from clogging and reproducibility issues.
Modern microfabrication techniques provide appealing opportunities for the implementation of immobilized enzyme reactors (IMERs). Wafer-scale fabrication processes enable not only parallelism and high degree of system integration, but also customization of the solid support structures with respect to both architecture and the surface chemistry. For instance, dense micropillar arrays have been implemented on silicon in order to increase the surface-to-volume ratio for on-chip chromatographic separations [8, 9, 10]. Owing to the possibility to pattern well-ordered micropillar arrays simultaneous to the microchannel network in a reproducible manner, no post-processing (channel packing) is needed and the problems related to clogging can be avoided. Although oxidized silicon readily provides a variety of surface functionalization reactions via silanol groups, which are mostly suitable for covalent coupling reactions, silicon microfabrication as such requires special facilities, including cleanroom instrumentation.
In this work, we introduce rapid replica molding of ordered, high-aspect-ratio micropillar arrays from off-stoichiometric thiol-ene polymers. By mixing the thiol and allyl (“ene”) monomers in off-stoichiometric ratios, the surface chemistry and the mechanical properties of thiol-ene microdevices can be tailored toward a variety of applications [11, 12, 13]. In the past, thiol-enes have been used in several applications, including microchip electrophoresis [13, 14] and biosensing [15, 16] devices, protein and DNA arrays , and on-chip electrospray ion sources for mass spectrometric detection [18, 19]. The off-stoichiometric thiol-enes (OSTEs) have also been exploited to fabricate porous monoliths inside thiol-ene microchannels followed by covalent coupling of proteolytic enzymes onto both thiol- and allyl-rich surfaces [4, 20]. Here, we make use of the native thiol-ene surface chemistry by preparing thiol-rich micropillar arrays and functionalizing them with gold nanoparticles (GNPs) via the well-characterized and strong thiol-gold interaction [21, 22]. Finally, we demonstrate the immobilization of α-chymotrypsin (CHT) onto GNP-coated thiol-ene micropillars via the enzyme’s free thiol groups (reduced disulfide bridges). The specificity of the thiol-gold interaction between both thiol-rich surface and GNP as well GNP and CHT is also examined, and the robustness of the developed CHT-IMER setup (with respect to flow rate and reaction temperature) is demonstrated by MS analysis of bradykinin hydrolysis products.
Materials and reagents
Methanol, dimethyl sulfoxide, acetone, tetrahydrofuran, toluene, acetic acid, ammonium acetate, ammonium hydroxide, 5,5′-dithiobis(2-nitrobenzoic acid) (DTNB), phosphate-buffered saline (PBS), α-chymotrypsin from bovine pancreas (CHT, type II, 40 units/mg protein), bradykinin acetate (fragment 1–9), and DL-dithiothreitol were purchased from Sigma-Aldrich (Steinheim, Germany). All reagents and solvents used were HPLC or MS grade (≥ 98.0%). Suspensions of bare GNPs (10 nm, stabilized in 0.1 mM PBS) and dodecanethiol functionalized gold nanoparticles (d-GNPs, 3–6 nm, 0.6–0.9% solid material, 0.01% HAuCl4 in toluene) were also from Sigma-Aldrich. Irgacure® TPO-L photoinitiator (2,4,6-trimethylbenzoylphenyl phosphinate) was donated by BASF (Ludwigshafen, Germany). Water was purified with a Milli-Q water purification system (Merck Millipore, Molsheim, France). SU-8 100 negative photoresist (Microchem Corporation, Newton, MA) used for master fabrication was purchased from Micro Resist Technologies GmbH (Darmstadt, Germany). Poly(dimethyl siloxane) (PDMS) used for fabrication of the replication mold was prepared from Sylgard 184 base elastomer and curing agent (Down Corning Corporation, Midland, MI). Trimethylolpropane tris(3-mercaptopropionate) (“trithiol”) (≥ 95.0%), pentaerythritol tetrakis(3-mercaptopropionate) (“tetrathiol”) (≥ 95.0%), and 1,3,5-triallyl-1,3,5-triazine-2,4,6(1H,3H,5H)-trione (“triallyl”) (≥ 98.0%) were used for microchip fabrication and purchased from Sigma-Aldrich (Saint Louis, MO).
The activity of immobilized CHT was determined by bradykinin hydrolysis. The stock solution of bradykinin (1 mM in Milli-Q water) was diluted to appropriate concentrations with 20 mM ammonium acetate (pH 8.2) before measurements. The 20 mM ammonium acetate solution used was prepared in deionized Milli-Q water and filtered (0.2 μm) before use in MS analyses. The pH of the ammonium acetate solution was adjusted using 10% (v/v) ammonium hydroxide. For determination of the specificity of the GNP interactions, phosphate-buffered saline (pH 7.4) was used as the buffer.
Microchip design and fabrication
The final IMER was fabricated by mixing the “tetrathiol” and “triallyl” monomers in a ratio that yielded 50 mol-% excess of thiol functional groups. This composition was chosen in order to maximize the number of free surface thiols, which is known to increase as a function of increasing excess of the thiol monomer . No photoinitiators were added to the composition to facilitate straightforward bonding of two alike surfaces as described in . However, in the absence of photoinitiators, the curing of thiol-ene compositions with very large excess of the thiol monomer becomes slow and thus the 50 mol-% excess of thiols was the practical upper limit for in this study.
For the specificity tests, also stoichiometric and allyl-rich (50 mol-% excess of allyl functional groups) compositions of “trithiol” and “triallyl” were used. No photoinitiators or other additives were added to any of the compositions. After mixing, the thiol-ene solution was poured onto the PDMS mold, featuring the negative replica of the micropillar array as microwells, and the mold was placed in vacuum for 2–5 min to effectively remove residual air bubbles trapped in the deep microwells (see ESM Fig. S1). Next, the thiol-ene monomer mixture was cured (without cover) under UV light for 5 min by using a Dymax 5000-EC Series UV flood exposure lamp (Dymax Corporation, Torrington, CT, USA, nominal intensity 225 mW/cm2). The planar cover layer incorporating only the inlet and outlet holes was prepared in a similar manner. The fully cured cover and bottom (micropillar) layers were then preheated to allow uniform sealing (here, ca. 70 °C was used) and laminated against each other. The bonding was finalized with additional UV exposure through the cover layer for 2 min. The bonding strength was determined by air-pressure tests using an in-house built gas delivery system consisting of an electronic regulator and solenoid valves (SMC Pneumatics Finland Oy, Espoo, Finland). The cured and bonded IMERs were stored at room temperature (RT) in the dark and under atmospheric pressure until use. Characterization of the microstructures was performed by a scanning electron microscope (SEM, FEI Quanta™ FEG, Hillsboro, OR) by attaching the samples onto the sample stage with a carbon-coated double-sided tape and sputtering (Quorum Q150TS, turbomolecular-pumped high-resolution coater, Quorum Technologies, UK) with platinum for 25 s (30 mA) to yield a ca. 5-nm-thick coating.
Surface functionalization and enzyme immobilization
The immobilization of CHT on the thiol-rich (50 mol-% excess) micropillar array included two steps, both exploiting the thiol-gold interaction. First, the micropillar array was filled with the gold nanoparticle suspension and incubated at 4 °C overnight. Next, the micropillar array was thoroughly rinsed with fresh buffer (20 mM ammonium acetate, pH 8.2) solution followed by another overnight incubation with CHT (1 mg/mL) at 4 °C. Before enzyme incubation, the disulfide bridges of CHT were reduced using 5 mM dithiothreitol (in buffer). Finally, the micropillar-based CHT-IMER was rinsed with 20 mM ammonium acetate (pH 8.2) before determination of the enzyme activity by MS.
Titration of free surface thiols
The amount of free thiol groups on micropillar surfaces were quantitated by titration using Ellman’s reagent . Briefly, a concentrated solution of 5,5′-dithiobis(2-nitrobenzoic acid) (DTNB) in PBS buffer was pumped through the micropillar array at a flow rate of 5 μL/min followed by quantitation of the reaction product, 2-nitro-5-thiobenzoate (TNB), by UV absorbance (412 nm) using Varioskan LUX Multimode Microplate Reader (ThermoScientific, Vantaa, Finland). Alternatively, the number of free surface thiols on planar surfaces was determined by submerging thiol-ene slabs (approx. 10 mm × 10 mm × 0.5 mm) into 1 mL of 200 μM DTNB in PBS for 30 min with stirring every 10 min. After 30 min, the thiol-ene slabs were removed and the reaction product TNB was quantitated by UV absorbance as described above. The number of free (reacted) thiols was determined using a molar extinction coefficient for the reaction product of ε = 14,150 M−1 cm−1 .
Water contact angle
The wetting properties of native thiol-ene and GNP-functionalized surfaces were characterized using a contact angle goniometer (Theta, Biolin Scientific, Espoo, Finland). Advancing and receding water contact angles were determined by the sessile droplet needle method using Young-Laplace fitting. The advancing contact angle was measured by increasing the droplet volume from 2 to 8 μL at a rate of 0.1 μL/s, and the receding angle by decreasing the volume from 8 to 0 μL at a rate of 0.1 μL/s.
Atomic force microscopy
An NTEGRA Prima (NT-MDT, Russia) atomic force microscope (AFM) was used for topographical analysis of the native and GNP-functionalized surfaces. The images (1024 × 1024 pixels) were captured in intermittent-contact mode at ambient conditions using gold-coated silicon cantilevers with a nominal tip radius of 10 nm (model: NSG 10, NT-MDT). The scanning rate and damping ratio were 0.2–0.3 Hz and 0.6–0.7, respectively. Image analysis was performed using the SPIPTM image analysis software (Image Metrology).
X-ray photoelectron spectroscopy
X-ray photoelectron spectroscopy (XPS) spectra were captured with a PHI Quantum 2000 scanning spectrometer, using monochromatic Al Kα x-ray source (1486.6 eV) excitation and charge neutralization by using electron filament and an electron gun. The photoelectrons were collected at an angle of 45° in relation to the sample surface with a hemispherical analyzer. The average depth of the XPS analysis was in the range of 5–10 nm. The pass energies of 187.85 eV and 29.35 eV were used for collecting survey and high-resolution spectra, respectively. The measurements were done on two different spots for each sample. The atomic concentration (%) of the different elements was derived by calculating the area of the peaks and correcting for the sensitivity factors using a MultiPak v6.1A software (Physical Electronics). The binding energies acquired in the XPS spectra were corrected using the C1s photoelectron peak at 284.8 eV as a reference.
Determination of enzyme activity
The hydrolysis of bradykinin (Arg-Pro-Pro-Gly-Phe-Ser-Pro-Phe-Arg, MW 1060) on the C-terminal side(s) of phenylalanine(s) was used as the model reaction to monitor the activity of immobilized CHT by electrospray ionization (ESI)-MS. The sample solutions (20 μM bradykinin in 20 mM ammonium acetate, pH 8.2) were infused with a syringe pump at a constant flow rate of 2.5, 5, 10, 15, or 20 μL/min and the reactants were collected for MS analysis in appropriate volumes (typically 100–150 μL per fraction). The effect of reaction temperature (RT vs. physiological temperature 37 °C) was examined by heating the CHT-IMER with a 0.5-Ω resistive heater block (Digi-Key, Thief River Falls, MN) attached to the bottom of the IMER with double-sided tape. The heating temperature was controlled with a DC power supply (Iso-Tech IPS-603, RS Components Ltd., Northants, UK) equipped with a PID temperature controller (type CN743, OMEGA Engineering, Manchester, UK). ESI-MS detection was performed on an Agilent 6330 iontrap mass spectrometer (Agilent Technologies, Santa Clara, CA) using direct infusion (5 μL/min). Before ESI-MS analysis, the collected sample fractions were diluted 1:1 with methanol-water 90:10 containing 0.2% (v/v) acetic acid. The ion trap was operated in positive ion mode with a capillary voltage set at − 3500 V and end plate offset at − 500 V. Nitrogen produced from compressed air by a Parker nitrogen generator (Cleveland, OH) was used as the drying gas (4.0 L/min, 325 °C). The MS data was acquired over a mass range of m/z 100–2000 with a maximum accumulation time of 300 ms using Data Analysis 3.4.
Specificity of the functionalization protocol
After incubation with the GNPs (immersion) or d-GNPs (droplet deposition), the surfaces were carefully rinsed with PBS prior to analysis with AFM, XPS, and contact angle goniometry. In all cases, 50 mol-% excess of either the thiol or allyl functional groups was used in the bulk to achieve thiol- or allyl-rich surfaces, respectively. While thiol-rich surfaces were considered ideal for maximizing the amount of thiol-gold interactions, the allyl-rich surfaces provided a good point of comparison with negligible amount of free surface thiols but otherwise very similar surface properties (e.g., in terms of wetting/hydrophobicity ). While the immersion approach better resembles the adhesion mechanism in a microchannel and necessitates the thiol-gold interaction, the toluene drop deposition approach facilitates the deposition of d-GNPs on any surface chemistry. Based on topographical analysis by AFM, the deposition of GNPs using the immersion approach (1-h incubation) provided a significant increase in the surface roughness (Sq) from 0.77 to 5.8 nm on thiol-rich surfaces (Fig. 2a vs. b), while their impact on the surface roughness of allyl-rich surfaces was negligible (Sq = 1.6–1.7 nm, Fig. 2c vs. d). In addition to roughness, the differently treated surfaces were examined in terms of the surface area ratio (Sdr) which expresses in percent how much larger the interfacial (real) surface area is compared with the area of the projected (flat) x,y plane. As illustrated in Fig. 2a–d, the effective surface area (Sdr) was also substantially greater for GNP incubated thiol-rich surfaces (Sdr = 6.1%, Fig. 2a) than for any of the controls (Sdr = 0.9…1.8%, Fig. 2b–d). These results suggest that the surface thiols were in a key role in facilitating GNP adhesion. Instead, the drop deposition resulted in much greater overall surface roughness independent of the surface chemistry, although the change was somewhat larger in case of thiol-rich surfaces. Compared with the aqueous PBS treatment (Fig. 2b and d), toluene treatment alone was shown to increase the surface roughness (Sq) and the effective surface area (Sdr) of both thiol- and allyl-rich surfaces (Fig. 2f and h). However, when d-GNPs were included in the toluene incubation, the surface roughnesses of both thiol- and allyl-rich surfaces were further increased from Sq = 5.4 nm to Sq = 11.5 nm and from Sq = 3.6 nm to 6.4 nm, respectively, compared to those of mere toluene-treated surfaces (Fig. 2e and g). Thus, the AFM data clearly evidenced deposition of d-GNPs and formation of the dodecanethiol layer on both thiol- and allyl-rich surfaces when drop deposition method was used, as expected. Generally, the drop deposition method (Fig. 2e vs. g) resulted in somewhat nonuniformly distributed summits and larger, aggregated objects on the surface, which was likely due to nonspecific adsorption of dodecanethiol onto the polymer surfaces preventing proper self-assembly of d-GNPs. Instead, the GNP deposition by the immersion method clearly favored thiol-rich surface (Fig. 2a vs. c) and resulted in a distinct granular morphology, which laid solid grounds for GNP adhesion onto microchannel surfaces.
Fabrication and characterization of the thiol-ene micropillar arrays
Previous studies have shown that the crosslinking degree, and thus, the rigidity and the bonding strength, of thiol-ene polymer networks can be greatly altered by varying both the monomer type (e.g., trithiol vs. tetrathiol) and the (off-)stoichiometric ratio of the thiol and allyl monomers [11, 12, 18]. Upon addition of a photoinitiator to the monomer mixture, the crosslinking occurs faster and typically in a quantitative yield compared to thiol-ene curing without the photoinitiators . As a result, only surfaces with opposite excess of thiols and allyls can be effectively bond to each other . However, by omitting the photoinitiator, also two alike thiol-ene surfaces can be bond together with fairly high bonding strength , which indicates that the structure rigidity, for its part, also plays a role. These previous findings further supported the use of the tetrathiol monomer with the triallyl monomer for the fabrication of the micropillar arrays to ensure sufficient rigidity of the replicated, high-aspect ratio (here h/w ~ 4) micropillar arrays. To maximize the number of free surface thiols toward efficient binding of GNPs, a 50 mol-% excess of thiol functional groups was used in the bulk, which resulted in 162 ± 16 nmol of free thiols per device (n = 3 titrations), corresponding to ca. 131 free thiols per nm2. The theoretical (calculated) increase in the total surface area and the surface-to-volume ratio, provided by the dense micropillar array incorporating total of 14,400 pillars (each ∅ 50.4 ± 0.6 μm, height 200 μm), was ca. 3-fold and 4-fold, respectively, over a hollow microchannel with identical dimensions (width 3 mm, length 40 mm, height 200 μm).
Consequently, straightforward and good quality sealing of the micropillar arrays with another identical surface was facilitated by omitting the photoinitiator. In this manner, the reverse sides (in contact with PDMS during UV curing) of both layers could be bond together with bonding strengths exceeding 2 bar (the upper limit of our pressure controlled test system). For comparison, micropillar arrays fabricated out of “trithiol” and “triallyl” monomers with equal excess (50 mol-%) of free thiols constantly broke already at 1.5 ± 0.4 bar (n = 4 IMERs). The bonding strengths remained unchanged even after delamination (upon heating) and re-bonding of the micropillar arrays, further suggesting that the rigidity of the chosen composition greatly affects the achievable bonding strength.
Before this work, high-aspect thiol-ene micropillar arrays have been achieved via direct photolithographic patterning only [30, 31]. Although the lithography approach allows high feature resolution, it necessitates the use of photoinitiators (and inhibitors), which often complicates adhesive bonding and results in a higher crosslinking degree, and thus lower thiol density on the surface. To examine the effect of the crosslinking degree on the free thiol density, the number of free surface thiols was determined for tetrathiol-rich (50 mol-%) thiol-enes cured (5 min) in the absence and in the presence of the photoinitiator (0.1% TPO-L). As a result, the amount of free thiols dropped dramatically from the initial 190 ± 43 thiols to only 12 ± 1 thiols per nm2 (ESM Fig. S3a). Similar decrease in the amount of free surface thiols (15 ± 1 thiols per nm2, ESM Fig. S3a) was also achieved by re-exposing the reverse “PDMS side” of the thiol-ene layer for another 5 min (in the absence of the photoinitiator). The impact of curing time was also confirmed by determining the amount of free thiols as a function of UV exposure dose (ESM Fig. S3b). On the average, these thiol densities correspond to ca. 14–18 nmol thiols per device, which is an order of magnitude lower amount than that on a tetrathiol-rich device cured in the absence of the photoinitiator and about an order of magnitude higher amount compared with stoichiometric (2.2 ± 0.3 nmol/device, n = 3) and allyl-rich (2.1 ± 0.05 nmol/device, n = 2) devices. Since the high density of free surface thiols was found crucial for the efficient coupling of GNPs (as shown in the previous chapter), the replication approach developed herein is likely the only way to achieve sufficiently high coverage of free surface thiols, and thus of GNPs, as it allows fabrication of thiol-ene micropillars without the photoinitiators. The sole critical step in the micropillar replication protocol developed in this study was to remove the air trapped in the deep wells of the PDMS negative mold, as illustrated in ESM Fig. S1, suggesting that the method is robust and feasible for low-cost fabrication of highly ordered micropillar arrays.
Enzyme immobilization and performance characterization
In this work, we introduce rapid replica molding of ordered, high-aspect-ratio, thiol-ene micropillar arrays for implementation of microfluidic immobilized enzyme reactors (IMERs) by exploiting thiol-gold interaction. The replica-molding method developed herein provides a straightforward approach for the fabrication of ordered micropillar arrays in non-cleanroom conditions. The possibilities to avoid the use of photoinitiators and to tune the thiol-ene surface chemistry via off-stoichiometry enable not only straightforward bonding but also good control over the number of free surface thiols available for GNP binding. Owing to the vast excess of thiol functional groups, we were able to bind GNPs on the native thiol-rich micropillars in an efficient manner so that these could be further exploited to immobilizing CHT also based on thiol-gold interaction between the GNPs and the thiol residues of the enzyme. Compared with microchannel packing with porous polymer monoliths or magnetic beads, the well-ordered, microfabricated pillar arrays allowed us to avoid the common pitfalls, such as clogging commonly associated with post-processed microchannel packings. The method qualification evidenced that the developed CHT-IMERs performed proteolytic hydrolysis (of bradykinin) in a robust and stable manner at RT and physiological temperature. The product conversion rate was most dependent on the flow rate (residence time), and almost complete (product/substrate ratio > 10) hydrolysis was achieved at a residence time of as short as 10 min (2.5 μL/min). Furthermore, the activity of the IMER remained stable for at least 1.5 h (continuous use), suggesting negligible leakage of CHT out of the IMER. As the enzymes were firmly immobilized, no further purification of the reaction solution was required prior to mass spectrometric detection. In all, the developed protocol is significantly straightforward, yet robust, while being also somewhat universal, since it can be applied for the immobilization of any proteolytic enzymes by their thiol residues. These are the main advantages of the developed IMER technology, which are likely to provide new opportunities for modern proteomics research.
We thank the Electron Microscopy Unit of the Institute of Biotechnology, University of Helsinki, and the Micronova Nanofabrication Centre, Aalto University, for providing access to the scanning electron microscope and the cleanroom facilities, respectively. Dr. Markus Haapala and Dr. Anu Vaikkinen are thanked for their help with mass spectrometry instrumentation.
Open access funding provided by University of Helsinki including Helsinki University Central Hospital. The research received funding from the European Research Council (ERC) under the European Union’s Seventh Framework Programme (FP/2007–2013) / ERC Grant Agreement number 311705 (CUMTAS). The work was also financially supported by the Academy of Finland (grant numbers 304400, 307466, 309608, and 297360), the University of Helsinki Research Funds, and the Doctoral Programme in Chemistry and Molecular sciences, University of Helsinki, and the Business Finland (grant number 211679).
Compliance with ethical standards
The research involves neither human participants and animals.
Conflict of interest
The authors declare that they have no conflict of interest.
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