Styrene/BSA-Br, I /EY/AscA | Reaction time (min) | BSA-Br (I ) consumption | |
---|---|---|---|
8 | 4000/1/1/0.5 | 120 | High |
Styrene/BSA-Br, I /EY/TEMED | Reaction time (min) | BSA-Br (I ) consumption | |
---|---|---|---|
Without irradiation. Formation of polystyrene nanoparticles. | |||
9 | 4000/1/1/10 | 10–15 | Quantitative |
10 | 4000/1/0.5/5 | 30–45 | Quantitative |
11 | 4000/1/0.2/2 | 120 | Near quantitative |
12 | 4000/1/0.02/0.2 | 240 or 480 | Low |
13 | 4000/0/1/10 | 240 | n.a. |
14 | 4000/0/0.2/2 | 240 | n.a. |
When control experiments were performed in the absence of a selected reaction component such as the catalyst ( Table 1 , entry 5), the monomer or irradiation ( Table 1 , entry 6 and Fig. S3 ‡ ), no biohybrid formation could be detected while in all cases, the macroinitiator was recovered unaffected. On the other hand, polystyrene nanoparticles were formed when styrene was subjected to emulsion polymerization conditions in the absence of the protein macroinitiator BSA-Br (I o ) ( Table 1 , entry 7 and Fig. 2 ). The produced polystyrene was isolated and characterized with 1 H-NMR spectroscopy (Fig. S5 ‡ ).
(A) FE-SEM micrographs and (B) TEM micrographs of BSA-poly(styrene) ( , entry 9) observed as two distinct populations of spherical nanoparticles with diameters between 100 and 130 nm and between 20 and 40 nm; (C) FE-SEM micrographs of poly(styrene) ( , entry 13) observed as spherical nanoparticles with diameters between 10 and 40 nm; and (D) FE-SEM micrographs of BSA coated poly(styrene) nanoparticles with diameters between 10 and 50 nm. |
Throughout this study, quantitative macroinitiator consumption was targeted as it minimizes the effort required to isolate the biohybrids by rendering only a simple dialysis step necessary. We evaluated grafting in the presence of AscA since it was elegantly employed in recent protocols as a means to deoxygenate the reaction mixtures. 63,65 Under the conditions used herein, grafting of styrene was found to proceed yet without quantitative macroinitiator consumption ( Table 1 , entry 8 and Fig. S3, S7 ‡ ). For this reason, we proceeded to investigate the effect of a tertiary amine sacrificial electron donor since it has been previously shown in EY mediated PET RAFT polymerizations that the stability of the generated amine radical cation enhanced both the efficiency of the reduction of excited-state EY and oxygen tolerance. 80–82 Indeed, when N , N , N ′, N ′-tetramethylethylenediamine (TEMED) was added as a sacrificial electron donor, 48 quantitative macroinitiator consumption could be attained after merely 10 to 15 minutes of blue LED irradiation at a feed molar ratio of styrene/BSA-Br, I o /EY/TEMED = 4000/1/1/10 ( Fig. 1 , Table 1 , entry 9, Fig. S3 and S8 ‡ ). When a reduced EY feed molar ratio was used, quantitative macroinitiator consumption could again be achieved, albeit at increased irradiation times (between 30 minutes and 2 hours depending on the photoredox catalyst loading, Table 1 , entries 10–12, Fig. S8 ‡ ).
Imaging of the products ( Table 1 , entry 9) with FE-SEM revealed two distinct populations of spherical nanoparticles with defined diameters between 100 and 130 nm and between 20 and 40 nm ( Fig. 2A and B ). Poly(styrene) formed in the absence of a macroinitiator appeared as spherical nanoparticles with significantly smaller diameters between 10 and 40 nm ( Fig. 2C , Table 1 , entries 13 and 14, Fig. S5 ‡ ). Hence, the spherical assemblies observed using FE-SEM can be most possibly attributed to hybrid polymer/bioconjugate nanoparticles ( Fig. 2A and B ). The nature of the nanoparticles was further elucidated through the synthesis of BSA-responsive polymer bioconjugates ( vide infra ). Taking into account the amphiphilicity of the bioconjugates and thus the lack of a solvent that would both dissolve them and preserve the conformation of the protein, the free polymer could not be efficiently removed from the product assemblies.
Next, intermittent light exposure was investigated to assess the possibility of activating and deactivating polymerization. Rapid macroinitiator consumption was observed after 2–3 minutes of irradiation, as can be observed in Fig. 1C (styrene/BSA-Br, I o /EY/TEMED = 4000/1/1/10, Table 1 , entry 9 and Fig. S9 ‡ ). This short induction period can be ascribed to the time required to in situ remove the oxygen from the polymerization solution and the time required for the EY radical anion to form and in turn interact with the amine co-initiator to kick-start the polymerization. 83–85 Under these conditions, the polymerization could be activated and deactivated by switching on and off the irradiation source until fully consuming BSA-Br (after 10 minutes of total ON irradiation time); nevertheless, temporal control was poor. We reasoned that the concentrations of EY and the tertiary amine would influence both the induction period and polymerization control, and therefore, to attain better temporal control, we lowered the concentration of the catalyst, i.e. , styrene/BSA-Br, I o /EY/TEMED = 4000/1/0.5/5 ( Table 1 , entry 10 and Fig. 1C ). The reaction could be again triggered or halted by turning the blue LEDs on and off with improved temporal control.
To further exploit this photoinduced methodology, we synthesized BSA-poly(styrene) on a larger scale using the same experimental setup. SEC verified that the macroinitiator consumption was quantitative on a 6 times larger scale without the need for further optimization (Fig. S9 ‡ ).
Characterization of amphiphilic, hydrophilic, and responsive protein–polymer conjugates. (A) IR spectra of the bioconjugates. (B) H-NMR spectra acquired for hydrophilic BSA-polymer conjugates. (C) Transmittance vs. time curve at different temperatures showing the rapid response of BSA-poly(NIPAM). (D) Transmittance vs. pH curves of BSA-poly(DPA). Two cycles are shown for the same sample in which the response was induced by changing the pH with the addition of HCl (pH decrease) or NaOH (pH increase). (E) SEM and FE-SEM micrographs of amphiphilic protein–polymer conjugate nanoparticles. |
Entry | Monomer | Monomer/BSA-Br, I /EY | BSA-Br (I ) consumption |
---|---|---|---|
Addition of 5% v/v toluene. | |||
1 | MA | 4000/1/1/10 | Quantitative |
2 | MMA | 4000/1/1/10 | Quantitative |
3 | VAc | 8000/1/5/50 | High |
4 | VP | 4000/1/0.5/5 | Quantitative |
5 | NAM | 4000/1/1/10 | Near quantitative |
6 | HEA | 4000/1/1/10 | Quantitative |
7 | HEMA | 4000/1/1/10 | Quantitative |
8 | NIPAM | 1000/1/1/10 | Quantitative |
9 | NIPAM | 1000/1/0.2/10 | Quantitative |
10 | NIPAM | 100/1/0.2/10 | Quantitative |
11 | DMAEMA | 4000/1/0.2/10 | Quantitative |
12 | DPA | 4000/1/0.2/10 | Quantitative |
As seen with styrene, several other monomers used in this study (MMA, DPA and NIPAM vide infra ) were also shown to polymerize in the absence of a macroinitiator (Fig. S6 ‡ ).
Grafting of the less activated monomer vinyl acetate (VAc) proved to be more demanding. In general, ATRP of VAc is considered highly challenging because the homolytic bond dissociation energy of the dormant poly(VAc) chains makes reactivation difficult while at the same time the VAc propagating radical is not stabilized either. 86,87 Indeed, neither addition of an organic cosolvent nor increased catalyst loadings or grafting times could significantly increase biomacroinitiator consumption. The lowest amount of unreacted macroinitiator was detected at a feed molar ratio of VAc/BSA-Br, I o /EY/TEMED = 8000/1/5/10 (Fig. S14 ‡ and Fig. 3E ).
We also sought to graft hydrophilic monomers from BSA-Br (I o ), and for this reason, vinyl pyrrolidone (VP), N -acryloyl morpholine (NAM) and 2-hydroxyethyl acrylate (HEA) were selected since all produce polymers useful in a variety of pharmaceutical and biomedical applications ( Fig. 3 ). 88,89 1 H-NMR spectroscopy provided an additional means to characterize hydrophilic protein–polymer conjugates while dialysis was sufficient to remove both unreacted monomers and the produced polymers from the bioconjugate solution. It should be noted that for hydrophilic monomers, the optimum conditions of emulsion polymerization did not result in macroinitiator consumption which was more difficult to attain. A feed molar ratio of VP/BSA-Br, I o /EY/TEMED = 4000/1/0.5/5 was found to be sufficient to yield BSA-poly(vinyl pyrrolidone) biohybrids ( Fig. 3 and Fig. S15 ‡ ). For the synthesis of BSA-poly( N -acryloyl morpholine), near quantitative macroinitiator consumption was observed after optimization with NAM/BSA-Br/EY/TEMED = 4000/1/1/10 ( Fig. 3 and Fig. S16 ‡ ). At the same molar loading, both 2-hydroxyethyl acrylate (HEA) and 2-hydroxyethyl methacrylate (HEMA) led to the formation of BSA-poly(HEA) ( Fig. 3 and Fig. S187 ‡ ) and BSA-poly(HEMA) ( Fig. 3 and Fig. S18 ‡ ), respectively.
Targeting the synthesis of responsive bioconjugates, N -isopropylacrylamide (NIPAM), 2-(dimethylamino)ethyl methacrylate (DMAEMA) and 2-(diisopropylamino)ethyl methacrylate (DPA) were grafted from the protein macroinitiator BSA-Br ( Fig. 3 ). The conditions identified for full macroinitiator consumption are summarized in Table 2 (Fig. S19–S24 ‡ ). To get insight into the kinetics of this oxygen-tolerant approach, the photoinduced grafting of NIPAM from BSA-Br (I o ) was further studied. In time course experiments performed under the conditions identified to be optimal (NIPAM/BSA-Br, I o /EY/TEMED = 2000/1/0.2/10, Fig. S19 ‡ ), the formation of biohybrids was apparent within the first 5 minutes of irradiation and full macroinitiator consumption could be achieved within 30 minutes. Importantly, when samples of the reaction mixture were withdrawn at fixed time points and studied with 1 H-NMR spectroscopy without purification, full monomer consumption was also seen after 60 minutes (Fig. S19 ‡ ). The lower critical solution temperature (LCST) of BSA-poly(NIPAM) was determined to be between 32.8 and 33 °C at sufficiently dilute concentrations and was found to be reversible ( Fig. 3 and Fig. S20 ‡ ). The spherical assemblies formed at temperatures higher than the LCST were imaged by SEM (Fig. S20 ‡ ). The response of BSA-poly(DPA) was also found to be reversible with the turning point determined to be at pH 5.8 ( Fig. 3 and Fig. S21 ‡ ). 90 Taking advantage of their response, both BSA-poly(NIPAM) and BSA-poly(DPA) could be effectively isolated from independently formed polymer chains, i.e. , by performing dialysis after phase transition while retaining the conditions required for the biopolymer to be hydrophilic (at 20 °C for BSA-poly(NIPAM) and at pH below 5.8 for BSA-poly(DPA)). BSA-poly(DPA) samples were collected before and after dialysis performed at pH 5.5 and analyzed with native PAGE (Fig. S22 † ). The dialysate was collected and the released polymer was isolated and characterized with 1 H-NMR spectroscopy (Fig. S22 ‡ ). Similar trends were observed in the synthesis of BSA-poly(DMAEMA) (Fig. S23 and S24 ‡ ).
Protein-coated polymer nanoparticles.
We therefore performed styrene polymerization in the presence of native BSA without adding the ATRP initiator BSA-Br, I o . In PAGE electrophoresis, a band not migrating past the stacking gel front was predominant upon completion of the reaction while native BSA could also be detected (Fig. S28 ‡ ). After the dialysis step, the presence of both BSA and poly(styrene) was confirmed in the product mixture with FT-IR. The nanoparticles were visualized via FE-SEM imaging to be spherical with diameters varying between 10 and 50 nm ( Fig. 2D ). Despite numerous efforts, the protein could not be fully detached from the nanoparticles by simple means that would allow further characterization post polymerization. We therefore proceeded to synthesize responsive poly(DPA) nanoparticles in the presence of native BSA ( Fig. 4 and Fig. S23 ‡ ). To determine the nature of the produced nanoparticles, the product was characterized after synthesis and was then subjected to dialysis against phosphate buffer, pH 5.0, i.e. , below the turning point (5.8, Fig. 3 ). Both the product and the dialysate were characterized. As seen in native PAGE analyses of the samples collected before and after the phase transition, native BSA (pI 4.8–5.0) was liberated, leaving no trace of the band attributed to the nanocarrier (Fig. S22 ‡ ). Our initial assumption was further supported through the detection of poly(DPA) obtained by acquiring a 1 H-NMR spectrum of the dialysate (Fig. S22 ‡ ).
Proposed pathways to produce BSA-polymer and BSA-coated polymer nanoparticles. |
(A) Left: imaging of BSA coated poly(styrene) with internal reflection fluorescence (TIRF) microscopy. Right: 3D intensity plot of one nanoparticle. (B) Activity of CALB-poly(styrene), polystyrene nanoparticles coated with CALB and native CALB. The graph depicts the slopes of the activity kinetics recorded at 20, 25 and 37 °C. |
CALB catalyzes the hydrolysis of esters, converting triglycerides into glycerol and fatty acids, while being also one of the most used enzymes in biocatalysis with widespread applications. 94 5-(6)-Carboxyfluorescein diacetate (CFDA) was used to test in vitro the catalytic activity of the CALB-poly(styrene) biohybrids by monitoring the formation of the hydrolysis product carboxyl fluorescein (CF) at 453 nm. The biohybrids were shown to retain part of the catalytic activity of the parent enzyme.
The CALB-coated polystyrene nanoparticles were also found to retain part of the esterase activity of native CALB ( Fig. 5 ). Notably, the coated nanoparticles were proven to be remarkably stable as they retained their activity after storing for one year at 4 °C (Fig. S33, ‡ activity data not shown). To the best of our knowledge, this is the first time that such significant stability of such nanocarriers is being reported.
Materials and methods, general polymerization protocol for the oxygen tolerant, ey/temed mediated grafting of monomers from protein macroinitiators, conclusions, author contributions, data availability, conflicts of interest, acknowledgements.
† Dedicated to the memory of Prof. R. J. M. Nolte, whose seminal contributions to the field and mentorship shaped our academic journeys. |
‡ Electronic supplementary information (ESI) available. See DOI: . Dedicated to the memory of Prof. R. J. M. Nolte, whose seminal contributions to the field and mentorship shaped our academic journeys. |
§ These authors contributed equally to this work. |
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The green synthesis of copper nanoparticles (CuNPs) using a leaf extract from Jatropha curcas (JC) has been documented in our present research work. The existence of flavonoids, tannins, glycosides, and alkaloids was confirmed by the phytochemical analysis of the plant extract and these chemicals can be used as reducing, stabilizing and capping agents.
2. Green synthesis of nanoparticles. Green synthesis can be defined as the derivation of materials from green or eco-friendly resources by the use of solvent, good reducing agent, and harmless material for stabilization (Citation 37).Additionally, this synthesis route is straightforward, cost-effective, dependable, sustainable, and relatively repeatable, and results in more stable compounds.
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Top-Down Synthesis breaks intermolecular connections to reduce bulk materials to nanoparticles. Physical or chemical erosion or biological disintegration may do this (Bashir and Liu, 2015).Bottom-Up Synthesis assembles nanostructures atom-by-atom or molecule-by-molecule, frequently by self-assembly (Majumder et al., 2007).Top-down procedures can make nano and microscale materials, whereas ...
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