Biofunctionalization of Water-Soluble poly(Phenylene Ethynylene)s

Abstract

We report the use of amide coupling chemistry to covalently link five different biofunctional groups onto an anionic water-soluble poly(phenylene ethynylene) (PPE) polymer. Two of the biofunctionalized PPEs are used in prototype applications, including pH sensing and flow cytometry labeling. The PPE is functionalized with carboxylate (R-CO₂⁻) and sulfonate (R-SO₃⁻) ionic groups. By using an activated ester, the amine-functionalized groups are covalently linked to the PPE polymer via amide linkages. The reaction chemistry is optimized using biotin-ethylene diamine, making it possible to control the loading of the biotin functionality on the PPE chains. Using the optimized approach, a family of five PPEs were prepared that contain biotin, rhodamine, cholesterol, mannose, or folic acid moieties appended to the polymer backbones. The rhodamine- and biotin-modified PPEs were further applied for pH response and flow cytometry applications. The reported approach can be utilized for other classes of water-soluble conjugated polymers, allowing facile development of a variety of new functionalized water-soluble conjugated polymers for a range of applications including sensing, bioimaging, and flow cytometry analysis.

Graphical Abstract

INTRODUCTION

Conjugated polymers are comprised of delocalized π-conjugated backbones and modifiable side chains. Through rational design of the backbone and side-chain structures, the physical and materials properties of conjugated polymers can be tuned on demand, enabling them to be widely used as active components of optoelectronic devices,^1,2 including organic solar cells,^3-5 organic light-emitting diodes,^6,7 and organic field-effect transistors.^8,9 The introduction of water-solubilizing groups (such as amino groups, quaternary ammonium groups, imidazolium groups, carboxylic groups, sulfonic groups, and phosphate groups) to the side chains enables application of conjugated polymers to problems in the biomedical area.¹⁰ The ionic side chains not only allow the polymers to disperse well in aqueous solution under physiological conditions, but they also provide electrostatic binding sites for a wide variety of biomolecules, including proteins and nucleic acids. Furthermore, introduction of functional groups into conjugated polymers can lead to materials that are perfectly designed for specialty applications. From small functional molecules, such as biotin,^11,12 rhodamine,^13,14 and folic acid,^15,16 to biomacromolecules including DNA,¹⁷ antibodies,¹⁸ and lipids,¹⁹ various molecules have been linked to or otherwise incorporated within conjugated polymer structures, for applications such as biosensing, imaging, and therapy.^20,21 Many methods for the functionalization of nanoparticles, quantum dots, carbon dots, graphene, carbon nanotubes, and other water-soluble materials were developed in the last few years.^22-26 The commonly utilized chemistries range from classical bioconjugation reactions, such as carbodiimide-mediated condensation and maleimide-mediated conjugation, click reactions including 1,3-dipolar cycloaddition, Diels–Alder reactions, and Staudinger ligation. Although many achievements have been made, the linking or attachment of biofunctional units onto conjugated polymers in a facile, controlled, and mild manner remains a challenge. In addition, most previous studies have targeted a specific application. A general approach to introducing a variety of functionalities onto water-soluble conjugated polymer platforms has not been reported. There are two methods to prepare functionalized conjugated polymers, monomer functionalization prior to polymerization and postpolymerization functionalization of the polymer. Postpolymerization functionalization is generally advantageous because it enables the synthesis of polymer libraries with consistent molecular weight and polydispersity, and it also avoids the possible negative influence of reactive functional groups on the polymerization reactions. The amidation reaction is a fundamental reaction in organic synthesis. Compared to the azide-alkyne and other “click” reaction chemistries,^27-29 which require catalysts, difficult to access precursor compounds, and nonaqueous reaction conditions, amidation is carried out under mild conditions and is compatible with aqueous solution. More importantly, amino groups are widely present in drugs, intermediates, natural products, and other biologically active compounds, which makes these compounds readily accessible for amidation.

Taking advantage of the amidation reaction, we have developed a platform for postpolymerization functionalization of a water-soluble poly(phenylene ethynylene) (PPE) type conjugated polyelectrolyte. Herein, we demonstrate the strategy by synthesis of PPEs that are functionalized with five different bio- or photo-active functional groups in a controlled manner in aqueous solution (Scheme 1). The precursor PPE is an alternating co-polymer featuring carboxylate (─R─COO⁻) and sulfonate (─R─SO₃⁻) solubilizing groups. The carboxylate groups are activated with appropriate reagents and react with various amine-containing molecules giving rise to the functional polymers. The sulfonate groups in the side chains give rise to good water solubility of the resulting functionalized conjugated polyelectrolytes. In this study, we have determined optimized reaction conditions for the amidation reaction and then utilized the method to prepare a family of five functionalized PPE-type conjugated polyelectrolytes. Conjugated polyelectrolytes modified with rhodamine and biotin were further studied with respect to their pH response and in an example flow cytometry study of biotin–avidin recognition. This work enables the facile introduction of a variety of functional groups on water-soluble conjugated polymers, which allows creation of new materials with diverse applications in biosensing, diagnostics, and therapy.

Scheme 1.

Functionalization of PPE

EXPERIMENTAL SECTION

Materials.

Organic solvents were purchased from Sigma-Aldrich and dried by elution using an MBraun MB-SPS-800 solvent purification system. Compounds 1 and 2 were synthesized according to the literature.^30,31 The synthesis and characterization of the polymers are described in the Supporting Information. Neutravidin-coated beads were purchased from Spherotech, Inc. Other chemicals were purchased from Sigma-Aldrich Chemical Company and Fisher Scientific and used as received. Deionized water (18.2 MΩ·cm) was obtained from a Milli-Q system (Millipore, Bedford, MA). 6~8 kDa molecular weight cutoff dialysis membranes were purchased from Fisher Scientific.

Measurements.

¹H NMR spectra were recorded on a Bruker Advance 500 MHz spectrometer. UV–visible absorption spectra were measured using a Shimadzu UV-2600 spectrophotometer. FT-IR spectra were measured using a Shimadzu IR Affinity-1S spectrophotometer. Corrected steady-state emission spectra were measured using an Edinburgh FLS 1000 photoluminescence spectrometer. The absolute fluorescence quantum yield was recorded on an integrating sphere. Gel permeation chromatography analysis was performed using an EcoSEC GPC system using polystyrene standards with THF as the eluent. Flow cytometry data were collected by using a BD Biosciences LSR-II instrument. Fluorescence lifetimes were measured using a PicoQuant FluoTime 300 fluorescence lifetime spectrophotometer by time-correlated single photon counting with a 405 nm diode laser. Confocal fluorescence microscope images were taken using a MicroTime 200 time-resolved fluorescence microscope (PicoQuant). The excitation was provided by a 405 nm diode laser, and detection was selected by a 425 nm long pass filter.

Flow Cytometry Analysis.

In a series of 5 mL polystyrene tubes, a portion of phosphate-buffered solution (PBS) (268 μL, pH = 7.4) was added followed by the addition of PBS of neutravidin-coated polystyrene beads (32 μL, 2.86 × 10⁷ particles/mL, 6.48 × 10⁶ neutravidin/particle, the concentration of neutravidin is 0.31 μM in the stock solution). The appropriate volume of 10 μM stock solution of PPE-biotin was added into the tube accordingly. The same procedure was repeated for a 10 μM stock solution of PPE. The solutions were allowed to sit for 2 h before being processed by a flow cytometer. On a BD Biosciences LSR-II flow cytometer, a 405 nm laser was used as excitation light and a Pacific Blue detection channel (450 ± 50 nm) was selected to collect the fluorescence of the polymer. Each tube was inserted, and the solution was drawn at a medium flow rate until 30,000 events were recorded in each sample. Gating and other statistics were obtained through FlowJo v10.

RESULTS AND DISCUSSION

Polymer Derivatization.

The synthesis of water-soluble PPE is outlined in Scheme 2. An AA/BB type polymerization under Sonogashira conditions of monomers 1 and 2 afforded the conjugated polymer PPE-ester in a 67% yield with a number average molecular weight (M_n) of 16,000 and a polydispersity index (Ð = M_w/M_n) of 2.1 determined by gel permeation chromatography analysis. The number average degree of polymerization is 16 (there are two phenylene ethynylene units per repeat). The water-soluble polymer PPE was obtained by base-promoted hydrolysis of PPE-ester carried out in a mixed 1,4-dioxane/water solution in a 95% yield. With the R─SO₃⁻ and R─CO₂⁻ groups on the side chains, PPE was readily soluble in water at room temperature. As shown in Figure S1, the UV–visible absorption spectrum of PPE in water features a maximum at 434 nm with a molar absorption coefficient of 3.6 × 10⁴ L·mol⁻¹·cm⁻¹ and the fluorescence spectrum exhibits a maximum at 460 nm in water with an absolute fluorescence quantum yield of 19%.

Scheme 2.

Synthetic Route to PPE

Coupling of PPE with biotin-ethylenediamine (Biotin-EDA, Scheme 3) was employed as a model reaction for optimization studies. Biotin was selected as the initial biofunctional group, given that it has an extraordinarily high affinity to avidin and has been widely used in proteomics, assembly, detection, labeling, and drug delivery.^32-35 A variety of coupling reagents and reaction conditions were used to explore the efficiency of the coupling reactions. Aminium salts (HATU; HBTU) and a carbodiimide (EDCI) with different promoters (DMAP, HOBT, and NHS) were assessed as coupling reagents (Table 1, see the footnote for acronyms). After each reaction, the product solution was diluted with water and dialyzed for 2 days to remove the small molecules and reaction byproducts. The extent of reaction of the available carboxyl units (m%) was assessed by analysis of the ¹H NMR of the derivatized polymer. (Note that m = 50% corresponds to on average one biofunctional group added per repeat unit). Most of the conditions afforded amidation except for EDCI and EDCI/DMAP in aqueous solution. The amidation reaction using EDCI/NHS as coupling reagents exhibited the highest m values (Table 1). We further optimized the reaction conditions by performing the EDCI/NHS amidation in different buffered aqueous solutions, and the results show that the reaction in MES buffer with a tunable pH (see Table 1) achieved the highest loading amount, m = 53%, corresponding to just over one biotin per repeat unit. This may be because that a relatively low pH value is beneficial for the formation of the NHS ester in step I, and in step II, the pH was tuned to 8 after Biotin-EDA was added, and the resulting slightly basic medium is beneficial for the nucleophilic addition of Biotin-EDA with the NHS ester.

Scheme 3.

Synthesis of PPE-biotin

Table 1.

Screening of Reaction Conditions

entry	coupling reagenta	solvent	% loading (m)
1	HBTU, iPr2NEt	H2O/DMF	15
2	HATU, iPr2NEt	H2O/DMF	22
3	EDCI	H2O	0
4	EDCI, DMAP	H2O	0
5	EDCI, HOBT	H2O	7
6	EDCI, NHS	H2O	37
7	EDCI, NHS	PBS (pH 7.4)	31
8	EDCI, NHS	MESb	53

^aHBTU = 1-[(dimethylamino)(dimethyliminio)methyl]-1H-benzo-[d][1,2,3]triazole 3-oxide hexafluorophosphate, HATU = 1-((dimethylamino)(dimethyliminio)methyl)-1H-[1,2,3]triazolo[4,5-b]pyridine 3-oxide hexafluorophosphate, EDCI = 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride, DMAP = 4-dimethylaminopyridine, HOBT = 1-hydroxybenzotriazole, NHS = N-hydroxysuccinimide.

^bIn step I, the pH value of MES buffer is 5. In step II, after adding Biotin-EDA, the pH of MES buffer is tuned to 8 by adding NaOH solution.

In order to test if Biotin-EDA could be introduced onto the PPE side chains with control over the degree of functionalization (m%), we performed five separate reactions under the same reaction conditions with varying molar equivalents (0.4, 0.8, 1.2, 1.6, and 2.0) of biotin relative to the concentration of the PPE repeat units. The ¹H NMR spectra demonstrated the coupling of biotin onto PPE and allowed quantification of the degree of functionalization. In particular, as shown in Figure 1, the methine proton signals (δ 4.32 ppm) on biotin were clearly observed in the spectra of PPE-Biotin. Integration of these peaks relative to the signals of the protons from the methylene unit adjacent to the PPE carboxylate units (δ 4.71 ppm) established the degree of biotin functionalization. As the number of equivalents of biotin used in the reaction increased, the loading amount of biotin increased from m = 18% to m = 53%, corresponding to approximately 1 biotin per 3 repeat units, to just over 1 biotin per repeat. This result clearly shows that it is possible to vary the loading of the biofunctional group on the polymer scaffold in a controlled manner by varying the reaction stoichiometry.

Figure 1.

Analysis of the loading of biotin onto PPE. ¹H NMR was performed at 75 °C in DMSO-d₆.

Having established the utility of the amidation reaction of PPE with Biotin-EDA, we further investigated the versatility of this postpolymerization method by introducing several other different functional molecules. As shown in Table 2, rhodamine, folic acid, mannose, and cholesterol were employed to functionalize PPE. The loading level (m%) of these functional groups was determined from the ¹H NMR spectra of the derivatized polymers (Figures S19-S26). The formation of the amide was also confirmed by FT-IR (Figure S27). The photophysical properties of the PPE derivatives are summarized in Table S1. The introduction of folate and rhodamine into PPE decreased the fluorescence quantum yield and lifetime of the polymer. PPE-biotin showed a higher fluorescence quantum yield and a longer lifetime than PPE. The fluorescence quantum yields of PPE-mannose and PPE-cholesterol slightly decreased.

Table 2.

Versatility of the Developed Functionalization Method^a

^a2 eq Rhodamine-NH₂, 1.2 eq Cholesterol-NH₂, 2 eq Mannose-NH₂, 0.8 eq Folate-NH₂. (The stoichiometry of the molecules was varied in order to obtain good water solubility of PPE-R’).

Five novel functionalized PPE derivatives were prepared by using the developed method, demonstrating the versatility of both the PPE-based platform and the postpolymerization functionalization reaction. The ability to prepare such functionalized, water-soluble conjugated polyelectrolytes in a straight-forward manner with good control of the loading level of functional molecules is desirable for many applications. For example, PPE-biotin could be used as a fluorescence label for recognizing biotin-binding proteins,¹² PPE-rhodamine could serve as a pH sensor (vide infra), and PPE-folic acid could selectively bind to a series of cancer cells for cell imaging and photodynamic therapy.^16,36 Finally, PPE-cholesterol could be used for cell membrane anchoring, and PPE-mannose could bind to lectin and specific bacteria for antibacterial studies.^37,38

Applications.

In order to provide examples of the utility of the functionalized conjugated polyelectrolytes, PPE-rhodamine and PPE-biotin were used in preliminary work to investigate the applications. First, fluorescence studies of PPE-rhodamine were carried out to determine whether energy transfer occurred from the PPE backbone to the rhodamine side groups. However, carrying out this work, we uncovered an interesting pH dependence of the optical spectra, which is primarily associated with the combined effects of pH on the rhodamine structure and a consequent effect on energy transfer from PPE to rhodamine. First, as shown in Figure 2a, the rhodamine chromophore exists in two forms: (1) a spirolactam that is colorless and nonfluorescent and is predominant at high pH; (2) a ring-opened form that absorbs in the mid-visible and is strongly fluorescent at low pH.³⁹ This effect is well-known and has been used to develop colorimetric and fluorescence-based sensors for a variety of applications including intracellular pH sensing.^40,41 As shown in Figure 2c, as pH is decreased from 8 to 3, the absorption and fluorescence of PPE-rhodamine undergo significant changes. In particular, at high pH, the absorption is dominated by the PPE backbone, which exhibits a single absorption at ~430 nm. As the pH decreases, a second band emerges at 540 nm, which is due to the ring opened form of the rhodamine units.⁴² More interestingly, the fluorescence undergoes significant changes with pH. At high pH, the fluorescence is dominated by the PPE fluorescence, with λ_max = 480 nm; as the pH decreases, the PPE fluorescence is quenched and replaced by a fluorescence at 570 nm, which is characteristic of the rhodamine chromophore. The quenching of the PPE emission signals the turn-on of energy transfer from the backbone to the covalently attached rhodamine chromophore, and it is confirmed by emission excitation spectra (see Figure S2). The overall changes in the fluorescence lead to a pronounced change in the emission color from blue to pink over the pH range 8–3. Due to the high sensitivity to pH, PPE-rhodamine is expected to be used as a potential probe to track the pH distribution in cells.

Figure 2.

(a) Structure conversion of PPE-rhodamine in basic and acidic conditions and the changes of emission colors from blue to pink as the pH values decrease from 8 to 3. (b) Absorption and (c) emission spectra of PPE-rhodamine at different pH values. The excitation wavelength was 360 nm. The concentration of PPE-rhodamine is 5 μM; the solvent is citrate–phosphate buffer.

In a second line of work, the use of PPE-biotin as a fluorescent label to signal the avidin–biotin interaction was explored. PPE-biotin is anionic in PBS (pH = 7.4) due to the presence of the R-CO₂⁻ and R-SO₃⁻ side groups. To investigate the interaction of PPE-biotin with a biotin-binding protein, neutravidin (NA), a deglycosylated native avidin from egg whites, was employed because of its low isoelectric point (pI ~6.3), which suppresses nonspecific electrostatic binding to the anionic polyelectrolyte. The loading of biotin on PPE-biotin used in this study was 53%, which means that each repeat unit of PPE-biotin has on average approximately one biotin unit. Thus, the concentration of PPE-biotin (as polymer repeat units) is approximately equal to the biotin concentration.

As shown in Figure 3a, the emission of PPE-biotin in the absence of neutravidin is broad, and the absorption shows a red-shifted band max at 470 nm, indicating that the polymer is aggregated in the aqueous PBS.^43,44 Interestingly, with the increasing concentration of neutravidin, the absorption maximum of PPE-biotin blue-shifts from 470 to 430 nm and the fluorescence maximum also shifts from 570 to 480 nm (Figure 3b). Taken together, these effects suggest that neutravidin leads to de-aggregation of the PPE-biotin. It is likely that the binding of the biotin units to the neutravidin leads to a “protein coating” on the polymer, which reduces the interchain interactions that are characteristic of the aggregated polymer. In contrast, the absorption and fluorescence spectra of PPE did not change much with the increasing concentration of neutravidin (Figure 3c,d), consistent with only a weak interaction between PPE and the biotin-binding protein.

Figure 3.

UV–visible absorption and emission spectra of (a, b) PPE-biotin (m = 53%) and (c, d) PPE with addition of different concentrations of neutravidin in PBS (containing 10% glycerol). The concentration of PPE-biotin and PPE was 2 μM. The concentrations of neutravidin increased from 0 to 1.50 μM (namely, 0, 0.12, 0.25, 0.37, 0.5, 0.62, 0.75, 1.00, and 1.50 μM). The excitation wavelength was 440 nm.

Flow cytometry is frequently used to analyze the physical and chemical characteristics of cells or particles. Although a variety of fluorescent labels have been developed for flow cytometry and some are commercially available, more novel functional agents are still needed. Conjugated polymers with large molar absorption coefficients, high fluorescence quantum yield, and excellent light stability are promising materials for application as high brightness fluorescence labels for flow cytometry. However, there are few reports of their use as flow cytometry labels or probes.^45,46 Herein, we report a proof of concept study that relies on the affinity of PPE-biotin to bind to neutravidin-coated polystyrene beads (Figure 4a). In this study, commercial neutravidin-coated polystyrene microspheres that are 6–8 μm in diameter were used. Each surface-bound neutravidin has on average two accessible biotin binding sites, while the other two sites are used to bind the protein to the polystyrene bead surface. The microspheres were premixed with aliquots of PPE-biotin (m = 53%), and then, the mixtures were analyzed by flow cytometry. The excitation was at 405 nm, and fluorescence detection was at 450 nm. As seen in Figure 4b, as the PPE-biotin concentration increased, the detected fluorescence intensity of the beads increased by up to an order of magnitude (Figure 4b). By contrast, addition of PPE (without biotin groups) to the beads did not lead to a significant change in the fluorescence intensity (Figure 4c). The difference in the behavior of the PPE-biotin is clearly due to the binding of the polymer to the beads via the biotin–neutravidin interaction.

Figure 4.

(a) Illustration of the binding of PPE-biotin with neutravidin-coated beads. (b) and (c) Fluorescence intensity of neutravidin-coated beads with the incubation of different concentrations of PPE-biotin and PPE. The concentration of the neutravidin was 33 nM. The concentration ratios of [PPE-biotin]/[neutravidin] were 4, 3, 2, 1, 0.5, and 0.25, respectively. The laser wavelength was 405 nm, and the fluorescence detection channel was 450 ± 50 nm. Thirty thousand events were recorded for each sample. (d) Fluorescence image of neutravidin-coated beads with the treatment of PPE-biotin. The concentration ratio of [PPE-biotin]/[neutravidin] was 4. The laser wavelength was 405 nm, and the detection was selected by a 425 nm long pass filter.

Careful inspection of the data in Figure 4b shows that the fluorescence from the PPE-biotin/neutravidin beads exhibits several peaks, suggesting that a heterogeneous distribution of beads is present in the samples. We hypothesize that this behavior is due to bead “aggregates” formed by crosslinking due to the fact that the PPE-biotin chains have multiple biotin units available for binding to more than a single bead. Inspection of the forward-scattering plots from the flow cytometry data (Figure S3) reveals the presence of particles that are larger, likely due to particle dimers and trimers. Confocal microscope images of a PPE-biotin/neutravidin bead sample (PPE-biotin/neutravidin 4:1) were obtained, and it is possible to observe the bright fluorescence characteristic of the polymer on the surface of the beads (Figure 4d). Interestingly, it is possible to observe three distinct bead dimers in the image, supporting the assignment of the multiple peaks in the flow cytometry data to particle dimers or aggregates. Taken together, these results show the promise of biofunctionalized PPE to be used as an effective fluorescent label in flow cytometry studies.

CONCLUSIONS

We have designed and synthesized a novel water-soluble PPE-based conjugated polymer with carboxylate group and sulfonate group side chains. The polymer was used as a platform for postpolymerization functionalization through the amidation of the carboxylate group with amine-bearing functional molecules via an optimized procedure. Biotin, rhodamine, folic acid, mannose, and cholesterol were incorporated into PPE side chains to demonstrate the versatility of the developed functionalization method. PPE-biotin and PPE-rhodamine were further employed to study their applications in pH sensing and flow cytometry. This work provides a method for postpolymerization functionalization of PPE derivatives with desired functional groups in a simple, general, and controlled manner. This approach can also be potentially utilized for other types of water-soluble conjugated polymers, which allows for the facile development of many new functionalized water-soluble CPEs and exploring new applications.