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Article Cite This: ACS Omega 2019, 4, 121−129
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Orthogonally “Clickable” Biodegradable Nanofibers: Tailoring Biomaterials for Specific Protein Immobilization Ozlem Ipek Kalaoglu-Altan,† Rana Sanyal,†,‡ and Amitav Sanyal*,†,‡ †
Department of Chemistry, Bogazici University, Bebek, 34342 Istanbul, Turkey Center for Life Sciences and Technologies, Bogazici University, 34342 Istanbul, Turkey
‡
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S Supporting Information *
ABSTRACT: Multifunctionalizable polymeric nanofibers can be tailored for various biomedical applications by selective conjugation of small molecules and bioactive ligands. This study reports the design, synthesis, and application of novel biodegradable polyester-based nanofibers bearing metal-free “clickable” handles. Polylactide-based polymers were synthesized using organo-catalyzed ring-opening polymerization to contain “clickable” chain-end functional groups that specifically react through radical or nucleophilic thiol−ene reactions. A furan-protected maleimide-containing hydroxyl-bearing initiator yielded polymers containing strained oxanorbornene unit at their chain end. In addition, postpolymerization thermal treatment provides maleimide end group-containing polymers. Solution electrospinning method was utilized to obtain bead-free nanofibers. Efficient conjugation on these nanofibers was demonstrated using metal-free conjugation reactions. It was observed that polylactide nanofibers undergo extensive biofouling, which limits their possible utilization for specific biomolecular immobilization. To alleviate this problem, polymers were modified to contain two orthogonally reactive functional groups, namely, the oxanorbornene unit and an azide group at their chain ends. The former reactive handle was used for conjugation of poly(ethylene glycol) chains to impart hydrophilicity and thus an antibiofouling ability, whereas the azide group undergoes strain-promoted azide−alkyne cycloaddition to install a protein-binding ligand such as biotin. These nanofibers were able to specifically immobilize the protein streptavidin with minimal nonspecific adsorption.
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treatment.1−3 Such treatments often compromise the mechanical stability of the polymeric material and do not introduce functional groups that allow efficient transformation directly without the need of additional wet chemical activation steps. An alternative to this approach involves utilization of reactive and clickable polymers for fabrication of nanofibers.4 This approach has gained a lot of interest since the advent of click chemistry, which utilizes transformations that allow efficient postpolymerization functionalization. Importantly, it is the high efficiency of click reactions under mild reaction conditions that enables facile functionalization of heterogenous interfaces as encountered during surface functionalization of bulk materials. For example, multifunctionalization of crosslinked polymeric films have been reported by Hawker et al. by employing combination of several click-based transforma-
INTRODUCTION Polymeric nanofibers have emerged as an attractive class of nanomaterials in biomedical sciences over the past decade because of their unique properties. In particular, the high surface area and porosity of nanofibrous materials make them suitable for immobilization and encapsulation of ligands, biomolecules, and cells. Additional properties such as their resemblance to the fibrillar structure of the extracellular matrix makes them suitable for adhesion and proliferation of cells, which govern cellular organization, survival, and their function. Furthermore, the availability of a wide pool of polymers enables tuning the chemical composition of nanofibers. Until recently, most polymeric nanofibers were fabricated using commercially available polymers which often lack functional groups that would enable immobilization of ligands and biomolecules in a specific manner with high efficiency under mild conditions. The lack of reactive sites on the surface of parent fibers is addressed through their surface modification using plasma treatment and wet chemical methods such as acid © 2019 American Chemical Society
Received: October 31, 2018 Accepted: December 18, 2018 Published: January 3, 2019 121
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Scheme 1. General Illustration of Utilization of Dual-Functionalizable Nanofibers To Introduce Bioactive Ligand against an Anti-biofouling Matrix
tions.5,6 Nanofibers provide an alternative surface morphology due to their nanosize dimensions. Initial reports of nanofiber modifications were based on the Huisgen-type Cu(I)-catalyzed azide−alkyne cycloaddition (CuAAC) reactions,7,8 a methodology which is now being employed with reservation because of the likelihood of residual toxic metal impurities in the material. Because of this reason, metal-free click reactions such as nucleophilic and radical thiol−ene reactions, strainpromoted azide−alkyne cycloaddition (SPAAC), and the Diels−Alder cycloaddition have gained popularity for the surface modification of nanofibers.9,10 For example, in a recent work by Becker et al., chain-end reactive poly(lactide) (PLA) nanofibers were prepared using 4-dibenzyocyclooctynol as the initiator for the ring-opening polymerization (ROP) of Llactide and were postfunctionalized with N3-GYIGSR peptide via SPAAC reaction.11 The same group also studied multifunctionality of nanofibers by sequentially functionalizing the poly(ε-caprolactone) (PCL) nanofibers with Gly−Arg−Gly− Asp−Ser (GRGDS), BMP-2 peptide, and dopamine via SPAAC, oxime ligation, and CuAAC, respectively.12 Other recent examples include the radical thiol−ene and Diels−Alder reactions for metal-free functionalization of polymeric nanofibers.13−15 Interestingly, there is lack of electrospun fibers that can undergo either nucleophilic thiol−ene or inverse electron demand Diels−Alder (iedDA) cycloaddition, both known to be very efficient click-type transformations. Interestingly, the oxanorbornene cycloadduct provides a molecular structure that can undergo rapid iedDA and radical thiol−ene addition, whereas the thermal treatment of cycloadduct yields the maleimide unit, a well-known Michael acceptor that undergoes rapid thiol−ene addition. Incorporation of these reactive handles in biodegradable nanofibers will certainly expand the toolbox of functionalizable biopolymers for fabrication of functional materials. Reactive biodegradable polymers are the frequently used materials for various biomedical applications that benefit from multiple attachment of biologically active materials such as
drugs, proteins, enzymes, growth factors, and so forth. In recent years, environmentally friendly biomaterials based on renewable resources have gained increasing attraction.16,17 In particular, aliphatic polyesters, such as poly(glycolide), PLA, and PCL, are among the commonly used polymeric materials employed in biomedical applications as a result of their biocompatibility and biodegradability properties.18,19 PLA has received particular interest as its raw material, L-lactic acid, can be obtained through the fermentation of renewable resources such as corn and sugars.20 Besides these advantages, PLA also possesses good mechanical properties. Consequently, PLA is widely used for biomedical applications such as drug delivery, absorbable sutures, medical implants, scaffolds for tissue engineering, and so forth.21,22 Owing to the wide range of applications, in this study we chose to utilize PLA for demonstrating the functionalization strategy, which can be readily adapted to other biodegradable polymers obtained using ROP. PLA can be obtained via the ROP of lactide, initiated by an alcohol in the presence of a catalyst. In our recent study, a furan-bearing PLA was synthesized via organocatalyzed ROP and electrospun into Diels−Alder clickable nanofibers.23 These nanofibers were modified with RGD peptide and found to promote cell attachment and proliferation. Although the Diels−Alder reaction with a furan-based system offers several advantages, it is limited in its ability to conjugate molecules bearing maleimide groups. In this study, biodegradable nanofibers that undergo facile functionalization using metal catalyst-free “click” reactions are fabricated, and their utility toward specific ligand-mediated protein immobilization is demonstrated (Scheme 1). A chainend functional PLA was synthesized via organo-catalyzed ROP using a furan-protected maleimide functional group containing alcohol as an initiator. After polymerization, the furanprotecting group was removed to obtain the thiol-reactive maleimide group. These polymers were electrospun from chloroform solution to yield nanofibers. Although these nanofibers could be readily functionalized with small thiol122
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Figure 1. Synthesis of chain-end reactive PLA polymers.
Figure 2. SEM images and histograms of (A) FM-PLA and (B) M-PLA nanofibers.
Figure 3. Thiol−ene-based functionalization reactions and fluorescence microscopy images of SAMSA dye conjugation on (A) FM-PLA nanofibers via radical thiol−ene and (B) M-PLA nanofibers via Michael reaction. Images from control experiments are shown as insets.
containing molecules using various “click” reactions, they failed as supports for selective biomolecular immobilization because of their high disposition toward biofouling. To expand the utility of these nanofibers for specific bioimmobilization, the hydroxyl end group of the PLA was converted to an azide group to provide dual-functionality. Ability of nanofibers to undergo dual-functionalization was investigated using the SPAAC and iedDA reactions with appropriate fluorescent
dyes. At this point, dual-functionalization was used to undertake biotinylation and PEGylation of nanofibers, which enabled immobilization of fluorescein isothiocyanate (FITC)labeled streptavidin in a specific manner.
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RESULTS AND DISCUSSION The furan-protected maleimide group containing PLA (FMPLA) which bears a strained alkene unit was synthesized using 123
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Figure 4. Evaluation of antibiofouling on nonPEGylated and PEGylated nanofibers and fluorescence microscopy images of FITC-BSA conjugation on (A) FM-PLA and (B) PEGylated FM-PLA nanofibers.
analysis of the nanofibers revealed that the thiol-containing fluorescence probe reacted with the alkene groups on the nanofibers (Figure 3a). No dye attachment was observed in a control experiment carried out using the similar procedure but no UV-irradiation, thus supporting the attachment of dye through radical thiol−ene conjugation. Although the radical thiol−ene reaction provides an effective method for conjugation of thiol-containing molecules, the method involves utilization of a photoinitiator and UVirradiation. Conjugation of thiol-containing molecules using the nucleophilic thiol−ene Michael-type reaction, on the other hand, proceeds without such requirements. It proceeds with high efficiencies under mild and benign conditions. Hence, the thiol-containing fluorescent dye 5-((2-(and-3)-S-(acetylmercapto)succinoyl)amino)fluorescein (SAMSA) was immobilized on the maleimide-containing M-PLA nanofibers in the dark for 6 h. The unbound dye was rinsed off with ample water, and the fibers were dried under a gentle stream of nitrogen. On investigation using fluorescence microscopy, green bright-green fluorescence of the functionalized nanofibers indicated successful conjugation through the Michael reaction (Figure 3B). As a control experiment, SAMSA-treated masked maleimide FM-PLA nanofibers did not show any significant attachment of the dye. After successful conjugation of thiol-containing molecules in a specific manner, we sought to expand the utility of these nanofibers for biomolecular immobilization of proteins. Ability to detect a protein selectively with nanofibers conjugated with specific ligands will also show their potential as viable platforms for detection. Specific attachment of growth factors and antibodies can modulate the interaction of these nanofiber scaffolds with various cells. However, to conjugate a biomolecule with specificity, the nanofibers should be inherently antibiofouling. Hence, we evaluated the suitability of nanofibers with masked and unmasked maleimide groups toward nonspecific binding of proteins. To our dismay, we noticed that when these nanofibers were treated with FITC bovine serum albumin (FITC-BSA), a high level of protein absorbed onto these nanofibers through nonspecific binding (Figure 4A). It is well known that incorporation of hydrophilic polymers such as poly(ethylene glycol) (PEG) into nanofibers reduces nonspecific absorption of biomolecules. Hence, we
a masked maleimide-containing alcohol as an initiator using organocatalyzed ROP, using a previously reported procedure.24 In our case, the catalyst system was replaced with a combination of 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) and 1-(3,5-bis(trifluoromethyl)phenyl)-3-cyclohexylthiourea (TU) and polymerizations were carried out in anhydrous dichloromethane (DCM) at room temperature under nitrogen atmosphere (Figure 1). The organo-catalyzed polymerization was chosen because it proceeds at ambient temperature and thus avoids the risk of unmasking the maleimide group via a retro Diels−Alder reaction and does not leave any residual metal impurities in the resulting polymer. The thus-obtained polymer was characterized using 1H NMR and size exclusion chromatography (SEC), the former characterization confirming the presence of the oxabicyclic reactive group in the polymer. The parent copolymer was modified to obtain the polymer M-PLA containing a maleimide group at the chain end by removing the furan protection through retro Diels− Alder reaction at 110 °C. This would enable conjugation of thiol-containing molecules through the nucleophilic thiol−ene reaction that does not require any thiyl radical formation. Nanofibers from the FM-PLA and M-PLA polymers were obtained using a solution electrospinning method. Optimum conditions for electrospinning were established by varying experimental parameters such as solvent, polymer solution concentration, applied voltage, solution injection flow rate, and the needle to collector distance. A solution of the polymer in chloroform (35 wt %) provided uniform nanofibers using 14 kV applied voltage with a needlecollector distance of 12 cm. Scanning electron microscopy (SEM) images indicated formation of bead-free nanofibers with average diameters of 278 ± 112 and 277 ± 131 nm, for FM-PLA and M-PLA nanofibers, respectively (Figure 2). Initial functionalization of FM-PLA nanofibers was examined by immobilization of an activated thiol-containing fluorescein dye via radical thiol−ene reaction. An aqueous solution containing the thiol-containing dye and 2-hydroxy-4′(2-hydroxyethoxy)-2-methylpropiophenone as the photoinitiator was dropped on the glass slide covered with FM-PLA nanofibers, which was then placed under UV (365 nm) irradiation for 5 min. The surface was rinsed with excess water to remove the unreacted dye. Fluorescence microscopy 124
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Figure 5. (A) Synthesis of the dual-functional PLA and (B) SEM and size-distribution histogram of FM-PLA−N3 nanofibers.
group and DBCO group at room temperature was confirmed from the green fluorescence of the nanofibers. As expected, the dual functionalizable FM-PLA−N3 nanofibers were also prone toward high nonspecific binding of proteins, as revealed from the green fluorescence of nanofibers on treatment with aqueous solution containing FITCstreptavidin (Figure 7A). Hence, the FM-PLA−N3 nanofibers were PEGylated to impart antibiofouling property using tetrazine−mPEG via the iedDA reaction. Nanofibers were treated with an aqueous tetrazine−mPEG solution for 6 h. On removal of unbound tetrazine−PEG and drying the nanofibers under nitrogen, they were treated with a solution of FITCBSA. To our satisfaction, presence of negligible green fluorescence confirmed that PEGylation induced antibiofouling (Figure 7B). Attachment of hydrophilic PEG chains was also evident from the change in the water contact angle that decreased from 120° ± 4.44° to 85° ± 10.17° after PEGylation (Figure 7C,D). Next, we aimed to examine if these PEGylated nanofibers containing reactive handles could undergo ligand-directed biomolecular attachment in a specific manner. PEGylated nanofibers were biotinylated using a DBCO-appended biotin via SPAAC reaction. The biotin−streptavidin pair was chosen because of the high biological interaction between biotin and avidin molecules. Furthermore, streptavidin contains four binding sites for biotin which allows the utilization of the remaining three vacant sites for further immobilization of other biotinylated biomolecules such as peptides and oligonucleotides to fabricate sensing platforms. The PEGylated nanofibers were treated with an aqueous solution of DBCO−PEG4− biotin at room temperature. After 6 h, the fibers were washed with ample water to remove the unbound ligands, followed by their incubation in an aqueous solution of FITC-streptavidin at room temperature in the dark. After 30 min, the nanofibers were rinsed with water to remove any nonspecifically bound streptavidin. The bright green fluorescence of the nanofibers indicated successful protein immobilization (Figure 8A). To verify the availability of the vacant ligand binding sites on the streptavidin, immobilization of a biotinylated dye Cy5-biotin was attempted. Streptavidin-immobilized nanofibers were
decided that attachment of PEG chains onto these nanofibers will decrease such undesirable biofouling. Treatment of FMPLA nanofibers with a PEG chain containing a tetrazine unit led to the attachment of hydrophilic short chains on the fibers through the tetrazine norbornene iedDA cycloaddition reaction. It was observed that such PEGylation of the FMPLA nanofibers imparted them with antibiofouling characteristic, which prevented the nonspecific FITC-BSA conjugation (Figure 4B). To realize nanofiber functionalization without sacrificing the reactive groups required for ligand attachment, we decided to introduce another reactive handle on the parent polymer. The presence of a chain-end hydroxyl functional group in the parent polymers provided a simple route to install an azide unit, a moiety well known to undergo highly efficient conjugation with cyclo-octyne-bearing molecules through the SPAAC reaction. The dual functionalizable polymer (FMPLA−N3) was prepared by converting the terminal hydroxyl group of FM-PLA polymer to an azide group by esterification with 4-azidobutanoicacid in the presence of 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (EDCI) and 4-(dimethylamino)pyridine (DMAP) (Figure 5). Presence of the azide group at the polymer chain end was confirmed from 1 H NMR spectroscopy and from the strong azide stretching vibration in the Fourier transform infrared spectroscopy spectrum of the polymer (see Supporting Information). Uniform bead-free nanofibers with average diameter of 297 ± 128 nm were obtained using electrospinning of a 35 wt % chloroform solution of the FM-PLA−N3 polymer. Ability to undergo dual-functionalization was probed by treatment of nanofibers with an aqueous solution of a tetrazine-appended Cy5-dye, via iedDA ene−tetrazine click chemistry. After incubation at room temperature, the unbound dye was rinsed off with excess water and successful conjugation was confirmed from their bright red fluorescence (Figure 6). Thereafter, the Cy5-conjugated FM-PLA−N3 nanofibers were treated with an aqueous solution of a DBCO-appended green fluorescent dye. Successful conjugation of the dye on the nanofiber via the SPAAC reaction between the azide-end 125
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Figure 7. Fluorescence microscopy images of FITC-streptavidin immobilization on (A) nonPEGylated and nonbiotinylated and (B) PEGylated and nonbiotinylated FM-PLA−N3 nanofibers. Water contact angles of (C) non-PEGylated and (D) PEGylated FMPLA−N3 nanofibers.
that can be functionalized either using radical thiol−ene or iedDA reaction. Furthermore, modification of the chain-end reactive group in the parent polymer to a nucleophilic thiolreactive form is achieved by simple unmasking of the maleimide group. Importantly, the available hydroxyl group at the other extremity of the polymer could be readily modified to incorporate an azide group, which undergoes catalyst-free conjugation through the SPAAC reaction. Multifunctionalization of nanofibers obtained using these polymers highlights the utility of orthogonal reactivity, where the azide group is used to incorporate the ligand necessary for specific protein immobilization and the oxanorbornene unit is used to integrate the antibiofouling PEG segment which enables the selective ligandmediated protein immobilization. One can envision that the facile fabrication and functionalization of these biodegradable “clickable” nanofibers would make them attractive candidates for various biomedical applications.
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EXPERIMENTAL SECTION Materials and Characterization. L-Lactide (Aldrich) was recrystallized from toluene twice, dried in vacuo at room temperature for 24 h, and stored in a nitrogen filled glovebox. DBU (Fluka) was dried over activated 4 Å molecular sieves and stored in a nitrogen filled glovebox. Furan-protected maleimide-containing alcohol,25 TU26 and 4-azido butyric acid27 were synthesized according to the literature procedure. Polymerizations were done in a glovebox using anhydrous DCM obtained from a dry solvent system. Toluene (VWR chemicals), DMAP (Aldrich), and EDCI (Alfa Aesar) were used as received. SAMSA (fluorescein; Invitrogen) was activated according to the literature procedure prior to use.28 Albumin FITC-BSA was obtained from Sigma. Cy5-tetrazine, DBCO−PEG4−carboxyrhodamine 110, tetrazine−mPEG (5 kDa), DBCO−PEG4−biotin, and Cy5-biotin were purchased from Click Chemistry Tools. FITC-streptavidin was purchased from Pierce. 1H NMR spectra of the polymers were recorded on a Varian 400 MHz spectrometer using trimethylsilane as an internal reference and deuterated chloroform as a solvent. The molecular weights of the polymers were estimated by SEC analysis using a PSS-SDV (length/i.d. 8 × 300 mm, 10 μm particle size) Linear M column calibrated with polystyrene
Figure 6. Orthogonal functionalization of nanofibers using SPAAC and iedDA reactions. Fluorescence microscopy images of (A) Cy5tetrazine-immobilized FM-PLA−N3 nanofibers via iedDA, (B) DBCO-appended dye conjugation on Cy5-tetrazine-immobilized FM-PLA−N3 nanofibers via SPAAC, and (C) merged image showing the dual-functionalization of the FM-PLA−N3 nanofibers.
treated with an aqueous solution of Cy5-biotin at room temperature for 30 min. The bright red fluorescence confirmed attachment of the Cy5-biotin molecule, thus demonstrating that the streptavidin-coated nanofibers could be further conjugated with biotinylated materials (Figure 8B).
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CONCLUSIONS In summary, we demonstrate that multifunctional biodegradable nanofibers can be readily tailored for particular applications through facile conjugation of (bio)molecules of interest. A furan-protected maleimide group containing alcohol-initiated ROP of lactide provides a parent polymer 126
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Figure 8. Illustration of ligand-directed protein immobilization on nanofibers and subsequent protein functionalization. Fluorescence microscopy images of (A) FITC-streptavidin immobilization on biotinylated FM-PLA−N3 nanofibers, (B) Cy5-biotin conjugation on FITC-streptavidinimmobilized FM-PLA−N3 nanofibers, and (C) merged image showing Cy5-biotin binding on FITC-streptavidin-immobilized nanofibers.
4-azidobutanoicacid (13.65 mg, 1.05 × 10−1 mmol), DMAP (4.155 mg, 3.4 × 10−2 mmol), and EDCI (6.52 mg, 3.4 × 10−2 mmol) were placed in a round bottom flask with a stir bar at room temperature using anhydrous DCM as the solvent. After 16 h, the solution was extracted with water, saturated aqueous sodium bicarbonate, 0.1 M HCl, and again with water. The organic solution was dried with Na2SO4 and concentrated in vacuo. ([M]0/[I]0 = 150, Mn,SEC = 14.0 kg/mol, Mw/Mn = 1.38). Electrospinning of PLA Polymers. Polymer solutions were prepared by dissolving polymers (250 mg) in CHCl3 (0.3 mL). Using a syringe pump, the solution was delivered at a constant flow rate to a blunt needle connected to a highvoltage power supply. For a typical experiment, the tip to collector distance was 12 cm, the applied voltage was 14 kV, and the flow rate was 0.01 mL min−1. Fibers were collected on 22 × 22 mm glass slides attached on a grounded aluminum collector. Fluorescent Dye Conjugation on FM-PLA Nanofibers via Thiol−Ene Reaction. SAMSA was activated according to the literature procedure prior to use. 5 μL of the activated SAMSA solution (1 mg/120 μL) in distilled water was dropped on FM-PLA nanofiber-containing glass slide using 1.1 μL 2-hydroxy-4′-(2-hydroxyethoxy)-2-methylpropiophenone (1.3 mg/mL) as the photoinitiator and exposed to UV light (365 nm) for 5 min. The slides were washed with ample water to remove the unbound dye. The conjugation was confirmed using fluorescence microscopy. Fluorescent Dye Conjugation on M-PLA Nanofibers via Michael Reaction. Activated SAMSA solution (5 μL, 1 mg/120 μL) in distilled water was dropped on M-PLA nanofiber-containing glass slide. The reaction proceeded for 6 h at room temperature in the dark. Thereafter, the slide was washed with ample water to remove the excess dye. The dye conjugation was confirmed using fluorescence microscopy.
standards (1−150 kDa) using a refractive-index detector. Tetrahydrofuran was used as the eluent at a flow rate of 1 mL min−1 at 30 °C. The morphology of electrospun nanofibers was observed with Jeol Neoscope JCM-5000 SEM. Fluorescence images of the samples were recorded at room temperature on a Zeiss Observer Z1 fluorescence microscope. UV light (365 nm) was used for thiol−ene reactions. Static water contact angles were measured in air via the sessile-drop method using a goniometer (CAM 101 KSV instruments). Approximately 5 μL of deionized water was deposited on the surface, and images were taken by an integrated digital camera. The software CAM2008 was used for image processing to determine the contact angle. The contact angle value for each sample was independently measured at three different locations and average values were measured. Synthesis of Alkene-Ended PLA (FM-PLA) Polymers. L-Lactide (400 mg, 2.78 mmol), TU (17.04 mg, 0.046 mmol), and furan-protected maleimide-containing alcohol (4.1 mg, 0.018 mmol) were placed in a vial under nitrogen atmosphere in a glove box. Anhydrous DCM (3 mL) was added as the solvent and DBU (13.88 μL, 0.092 mmol) was added to the reaction mixture. The reaction was left to stir for 24 h at room temperature in the glovebox. The reaction was quenched with benzoic acid and the resulting polymer was purified by precipitation in 1:1 methanol/ether. The polymer was dried in vacuo for 24 h at room temperature. ([M]0/[I]0 = 150, Mn,SEC = 14.7 kg/mol, Mw/Mn = 1.31). Synthesis of Maleimide-Ended PLA (M-PLA) Polymers. Furan protection of FM-PLA was removed by heating the copolymer at 100 °C in toluene. After 20 h, toluene was evaporated under reduced pressure and the resulting polymer was dried in vacuo at room temperature. ([M]0/[I]0 = 150, Mn,SEC = 13.8 kg/mol, Mw/Mn = 1.32). Synthesis of Dual-Functional PLA (FM-PLA-N3) Polymers. The copolymer FM-PLA (500 mg, 3.4 × 10−2 mmol), 127
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Notes
FITC-BSA Conjugation on FM-PLA Nanofibers. FITCBSA solution (10 μL) in 1× phosphate buffered saline (1 mg/ mL) was dropped on the FM-PLA nanofiber-coated glass slide in the dark at room temperature. After 6 h, surfaces were washed and dried under a nitrogen stream. As a control experiment, PEGylated FM-PLA nanofibers were treated with FITC-BSA under the same conditions. Dual Fluorescent Dye Conjugation on FM-PLA-N3 Nanofibers. Cy5-tetrazine solution (10 μL, 1 mg/mL) in distilled water was dropped on an FM-PLA−N3 nanofibercoated glass slide. After 6 h at room temperature in the dark, the glass slide was washed with water to remove any unbound dye. Then, an aqueous solution of DBCO−PEG4−carboxyrhodamine 110 (10 μL, 1 mg/mL) was dropped on the Cy5tetrazine-immobilized nanofibers. The reaction was carried out for 6 h in the dark at room temperature. The slide was washed with ample water to remove the excess dye and dried under nitrogen stream. The conjugations were analyzed using fluorescence microscopy. PEGylation and Enzyme Immobilization on FM-PLAN3 Nanofibers. At first, FM-PLA−N3 nanofibers were PEGylated using an aqueous solution of tetrazine−mPEG (5 kDa). Tetrazine−mPEG solution (10 μL, 1 mg/mL) was dropped on the FM-PLA−N3 nanofiber-containing glass surface. After 6 h, the slide was washed with water and dried under nitrogen stream. Then, the fibers were biotinylated by applying 20 μL of aqueous DBCO−PEG4−biotin solution (1 mg/mL) on the nanofibers for 6 h at room temperature. The surface was washed with ample water to remove excess biotin and dried under a stream of nitrogen. An aqueous FITCstreptavidin solution (10 μL, 0.1 mg/mL) was dropped onto the biotinylated nanofibers at room temperature in the dark. Excess streptavidin was rinsed off with water after 30 min, and the surface was dried under a stream of nitrogen. Finally, 10 μL of aqueous Cy5-biotin solution (0.1 mg/mL) was dropped on the streptavidin-immobilized nanofibers at room temperature. After 30 min in the dark, the fibers were rinsed with ample water and dried under a stream of nitrogen. All conjugations steps were analyzed using fluorescence microscopy.
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The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors acknowledge financial assistance for this research by The Scientific and Technological Research Council of Turkey (TUBITAK project no. 114Z139) and Ministry of Development of Turkey for grant nos. 2009K120520 and 2012K120480. O.I.K.-A. acknowledges TUBITAK BIDEB 2211-A for financial support.
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ASSOCIATED CONTENT
* Supporting Information S
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsomega.8b03041. 1
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REFERENCES
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HNMR Spectra and GPC traces in THF of FM-PLA, M-PLA, and FM-PLA-N3, and FT-IR spectra of FMPLA and FM-PLA-N3 (PDF)
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Phone: +90-212-3597613. Fax: +90-212-287-2467 (A.S.). ORCID
Rana Sanyal: 0000-0003-4803-5811 Amitav Sanyal: 0000-0001-5122-8329 Author Contributions
The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. 128
DOI: 10.1021/acsomega.8b03041 ACS Omega 2019, 4, 121−129
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