Direct covalent attachment of fluorescent molecules on plasma polymerized nanoparticles: a simplified approach for biomedical applications

Abstract

Polymeric nanoparticles surface functionalised with fluorescent molecules hold significant potential for advancing diagnostics and therapeutic delivery. Despite their promise, challenges persist in achieving robust attachment of fluorescent molecules for real-time tracking. Weak physical adsorption, pH-dependent electrostatic capture, and hydrophobic interactions often fail to achieve stable attachment of fluorescent markers. While covalent attachment offers stability, it often entails laborious multi-step wet-chemistry processes. This work demonstrates that plasma polymerised nanoparticles (PPNs) can directly and covalently attach fluorescent molecules with no need for additional interim treatment processes. For the first time, we provide evidence indicating the formation of covalent bonds between the fluorescent molecules and PPN surfaces. Two model fluorescent molecules, fluorescein isothiocyanate (FITC) and Nile blue (NB), were attached to PPNs in a one-step process. The attached molecules remained on nanoparticle surfaces even after detergent washing, as confirmed by a combination of X-ray photoelectron spectroscopy (XPS), fluorescence spectroscopy, flow cytometry, and time-of-flight secondary ion mass spectrometry (ToF-SIMS) data. The robust attachment of fluorescent molecules on PPNs ensures their stability and functionality, enhancing the potential of these fluorescently labelled nanoparticles for diagnostic, therapeutic, and imaging applications.

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Behnam Akhavan

Dr Behnam Akhavan, an Australian Research Council (ARC) DECRA Fellow and an Associate Professor of Biomedical Engineering at the University of Newcastle, Australia, heads the Plasma Bio-engineering Laboratory at the School of Engineering and the Hunter Medical Research Institute (HMRI). Since obtaining his PhD in Advanced Manufacturing from the University of South Australia in 2015, he has held postdoctoral and academic positions at the Max Planck Institute for Polymer Research and Fraunhofer Institute of Microtechnology in Germany, and the University of Sydney. AProf. Akhavan's pioneering work in plasma surface bio-engineering, published in over 80 journal articles, has led to innovative applications in healthcare and beyond. He is recognised by Engineers Australia as one of the nation's Most Innovative Engineers.

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1 Introduction

Surface-functionalized nanoparticles have become extensively popular over the past few decades as a versatile platform for a variety of applications ranging from environmental remediation^1,2 and gas sensing³ to printed electronics⁴ and numerous applications in the ever-growing biomedical field.^5–10 Where bioactive molecules and fluorescent molecules are coupled, unprecedented functionalities can be achieved, making them attractive for applications in disease treatment,^11,12 diagnostic assays,¹³ delivery of therapeutic agents,^14,15 or a combination of the above.^16,17 In particular, their surface modification with fluorescent markers has amplified their functional capabilities for in vivo multi-modal imaging^18–20 and hybrid sensing techniques.^21,22 The development of fluorescent nanoparticles provides the additional benefit of enabling their real-time tracking with a higher stability and better resistance to photobleaching than that of the common fluorescent dyes.²³

Numerous methods for coupling fluorescent molecules with nanoparticles have been explored. Physical adsorption between fluorescent molecules and polymer matrices is considered a relatively minimally complicated procedure; however, the resultant association force is weak, and the fluorescent molecules tend to leach into the surrounding environment of the particle.^24,25 Electrostatic capture, involving the attraction between charged fluorescent dyes and oppositely charged nanoparticles, is another commonly applied method for fluorescent molecule attachment onto polymeric nanoparticles. Despite its prevalence in chemical research, it yields pH-dependent bonding, which can readily deteriorate with minor changes in pH.^26,27 An alternative method for attaching fluorescent molecules onto polymeric nanoparticles involves relying on hydrophobic interactions,^28,29 yet its major downsides arise from the need for numerous wet-chemistry steps and the variable and often weak nature of such interactions.^30,31

Covalent attachment provides higher robustness and bond stability compared to physical attachment methods, but the process is often tedious and time-consuming.³² For example, the covalent labelling of FITC onto chitosan-coated nanoparticles required a 3-hour reaction in the dark, precipitation using a high pH solvent, and dissolution in acetic acid before being dialyzed in distilled water for 3 days in darkness.^33–36 The chemicals and reagents utilised in the covalent binding process may also be toxic, posing challenges for regulatory approvals. As such, there is a growing demand for a simple, yet versatile, approach that offers the advantages of covalent attachment but without these limitations.

Plasma polymerization is an emerging technology for producing reactive and cytocompatible organic nanoparticles for biomedical applications.³⁷ In this technology, a precursor monomer is introduced into a vacuum chamber and excited into plasma state by an applied electric field. The fragmentation of monomer into polymer-forming species leads to the nucleation of clusters or ‘proto-particles’ approximately 10 nm in size.^38–40 The proto-particles grow as radicals and ions accrete onto their surfaces, eventually accumulating a negative charge and being expelled from the plasma volume once they reach a critical mass. The stability and cytocompatibility of the resulting nanoparticles are attributed to their highly cross-linked structure, controlled surface chemistry, solvent-free synthesis, and the use of organic monomers, which polymerize into non-toxic materials.^38,41 This technique has been used to create nanoparticles enriched with specific functional groups, such as carboxyl-enriched particles using acrylic acid monomer.^42,43 In previous work, plasma-polymerized nanoparticles (PPNs) using mixtures of acetylene, argon, and nitrogen were synthesised.^37,44,45 The highly radical activated surfaces of such PPNs make them promising candidates for direct covalent linking of fluorescent molecules without the need for additional linkers or reagents.

Here, we demonstrate that the synthesis of surface-active polymeric nanoparticles via the plasma polymerization technology facilitates the covalent attachment of fluorescent molecules, while providing evidence validating the covalent nature of the bonds. As a proof of concept, we use a single incubation step to conjugate each of model fluorescent molecules: fluorescein isothiocyanate (FITC) and Nile blue (NB). FITC is a traditional fluorescein molecule widely prevalent for its fluorescent properties,^46–48 while NB is a lipid-targeting fluorescent molecule, most popular in photodynamic tumour therapy.⁴⁹ Both fluorescent molecules have relatively large extinction coefficients, high quantum yields, and solvent-dependent optical properties that make them ideal for the development of nanoprobes. We anticipate that this work will broaden the scope of covalently surface-functionalised nanoparticles, paving the way for creating the next generation of robust fluorescently labelled polymeric nanoparticles with modern applications in biomedicine and beyond.

2 Materials and methods

2.1 Materials

Argon (99.99%), nitrogen (99.99%), and acetylene (98%) gases, used in the plasma process, were supplied from BOC Australia. Fluorescein isothiocyanate (FITC), Nile blue A (NB), and all other buffer reagents, such as sodium dodecyl sulfate (SDS) and phosphate-buffered saline (PBS), were purchased from Sigma Aldrich and used without further processing, unless stated otherwise.

2.2 Plasma polymerization of nanoparticles

Nanoparticles were produced through plasma polymerization inside a stainless-steel chamber equipped with a ring-shaped gas dispenser tube positioned above the primary electrode powered by a 13.56 MHz (RF) generator. This configuration is sketched in Fig. 1. The chamber was evacuated to a base pressure of 5 × 10⁻⁵ Torr to eliminate impurities and moisture. Argon, nitrogen, and acetylene gases of high purity were introduced into the reaction chamber at specific flow rates of 3, 10, and 6 sccm (standard cubic centimeters per minute), respectively. The flow rate of each gas was controlled using computer software (FlowVision) and mass flow controllers (Allicat Scientific). Prior to nanoparticle generation, the pressure was set at a working level of 1.5 × 10⁻¹ Torr by adjusting an inlet valve connecting the chamber to the vacuum pumping system. To ensure the correct desired power of 100 W is supplied to the reactor, a matching network was used to adjust the ‘reverse power’ to 0 W. The plasma polymerized nanoparticles (PPNs) were collected in a 24-well cell culture plate placed on the lower electrically floating electrode (Fig. 1). All particle characterisation experiments were performed within 24 hours of PPN production, unless otherwise mentioned.

Fig. 1 Sketch of the plasma polymerization system depicting the nanoparticle collection process. In the sketch, the primary and floating electrodes can be seen as well as the top gas dispenser tube through which argon, nitrogen, and acetylene gases are let in at 3, 10, and 6 sccm respectively.

2.3 Optical emission spectroscopy

During the nanoparticle generation process, non-intrusive optical emission spectroscopy (OES) was utilized to monitor the polymerization process and the intensities emitted by excited species in the plasma phase. Emission spectra were captured using an Acton SpectraPro 2750 spectrometer from Princeton Instruments, featuring a grating with 1200 grooves per mm. The spectrometer was configured with an exposure time of 100 μs, 2000 consecutive acquisitions were averaged to maximize the signal-to-noise ratio, and an intensifier gain of 250 in shutter mode. An optical fibre, aligned perpendicularly to the chamber window was connected to the spectrometer for spectral acquisition, and emission intensities across specific wavelengths were recorded on a computer screen using the Winspec-32 software.

2.4 pH-dependent hydrodynamic size and ζ potential measurements

The nanoparticle size and zeta potential in solution were determined using a Malvern Zetasizer Nano series optical unit. Unless otherwise specified, all measurements were performed in de-ionized MilliQ® water, with a pH value of approximately 5.5, as confirmed by a pH meter.

For experiments involving varied pH levels, different pH buffer solutions were prepared while maintaining a consistent ionic strength with 0.001 M NaCl solutions. Six 50 mL 0.001 M NaCl solutions were adjusted to different pH values using titration methods. Specifically, 0.1 mL of 0.05 M HCl solution was added to each NaCl solution, followed by the addition of 0 mL, 0.45 mL, 0.695 mL, 0.8 mL, 0.9 mL, and 1.15 mL of 0.01 M NaOH solution into samples 1 to 6 respectively. The actual pH values of these solutions were measured using a pH meter, resulting in values of 4, 5, 6, 7, 8 and 9 for samples 1 to 6 respectively. Next, 8 × 10⁸ nanoparticles were suspended in 2 mL of each pH-adjusted solution, achieving a concentration of 4 × 10⁸ particles per mL. Absorbance at 405 nm excitation light was measured using a Biotek plate reader, yielding a value of 0.44 ± 0.05 with MilliQ® water as the control. The nanoparticles were then transferred to disposable folded capillary cells (Malvern, DTS1070) from polystyrene vials under sterile conditions using 1 mL syringes. Zeta (ζ) potential measurements were conducted, and the average ζ potential values for specific pH levels, along with standard deviations representing errors, were reported. Hydrodynamic size measurements and the average size were determined based on three measurements obtained from at least three independent runs.

2.5 Attenuated total reflection – Fourier transform infrared spectroscopy (ATR-FTIR)

The study of functional groups was conducted using a Bruker ALPHA portable FTIR system (Bruker Optik GmbH, Germany) equipped with a platinum attenuated total reflection (ATR) single reflection diamond attachment. Background scans were performed before each sample scan. All background and sample spectra were collected with a resolution of 4 cm⁻¹, spanning the spectral range from 400 to 4000 cm⁻¹ with an average of 128 scans. Subsequently, all FTIR spectra were subjected to processing and baseline correction using OPUS 8.7 software (Bruker Optik GmbH, Germany).

2.6 X-Ray photoelectron spectroscopy (XPS)

The chemical state and composition of the PPN sample surfaces were investigated using XPS with a Thermo Fisher K-Alpha+ XPS/UPS system (Thermo Fisher Scientific, USA). The XPS system utilized a monochromated, microfocused Al K-Alpha X-ray source and a 180° double focusing hemispherical analyzer equipped with a 128-channel detector. The PPNs collected in the 24-well plate were suspended post-synthesis in 10 mL Milli-Q water and transferred to a 15-mL centrifuge tube, followed by centrifugation using an Eppendorf 5810 centrifuge at 8000 rpm for 10 minutes. The PPNs were freeze-dried using an Alpha 2–4 LSCbasic Freeze Dryer for 24 hours then arranged on an indium foil affixed into a double-sided conductive carbon tape on the XPS sample holder. The survey spectra were collected at a pass energy of 200 eV and a resolution of 1 eV, while the high-resolution C 1s spectra were recorded at a pass energy of 50 eV and a resolution of 0.1 eV. The base pressure was below 5 × 10⁻⁸ mbar for all the measurements. A total of 10 scans were conducted for each sample. Elemental composition calculations and peak fitting for high-resolution peaks were performed using the Avantage software (Thermo Fisher Scientific, USA).

2.7 Time of flight secondary ion mass spectrometry (ToF-SIMS)

ToF-SIMS was performed using a Physical Electronics Inc. PHI TRIFT V nanoTOF® instrument, with a pulsed liquid ⁷⁹⁺Au primary ion gun, operated at 30 kV. Dual charge neutralization was utilized in the form of an electron flood gun and Ar⁺ ions (both 10 eV). The collection time per spot was 3 minutes, and the raster size was 10⁻⁴ cm², with 7 spots per sample. All resulting spectra were processed using WincadenceN® software (version 1.8.1.3 Physical Electronics Inc.).

2.8 Water contact angle measurements

The hydrophobicity of the nanoparticles was assessed by measuring the contact angle formed by a water droplet on a silicon substrate coated with a dense layer of nanoparticles. The measurements were performed using a Biolin Scientific Theta Tensiometer in the sessile drop configuration. 10 water droplets with volumes of 3 μL were deposited onto the particle-coated substrate. The interaction behavior with water was closely observed and the average minimum contact angle was measured to be (23.8 ± 0.6)°. The reported contact angle value of each sample is the average of at least three measurements.

2.9 Scanning electron microscopy (SEM)

The surface morphology of plasma polymerized nanoparticles was investigated using a Zeiss Ultra Plus scanning electron microscope (SEM). The experimental setup involved configuring the microscope with an aperture size of 30 μm, operating at a low voltage of 5 kV, and maintaining a working distance of 5 mm, using the in-lens detector. Subsequently, the SEM micrographs were analyzed and processed using the Smart Tiff software. Nanoparticles were stored in a dry state for one week then dispersed in MilliQ® water at a concentration of 4 × 10⁸ particles per mL. They were then dropped onto a clean silicon wafer and left to naturally air dry before capturing scanning electron microscopy (SEM) images.

2.10 Electron paramagnetic resonance (EPR) spectroscopy

A Bruker EMX X-band EPR spectrometer was utilized to measure spin density in the PPNs two hours after synthesis. The spectrometer operated at room temperature with a microwave frequency of 9.8 GHz, a central magnetic field of 3510 G, sampling time of 90 ms, modulation amplitude of 3 G and a microwave power of 25 mW. The reported EPR spectra represent the average of 10 scans.

2.11 Conjugation of FITC to plasma polymerized nanoparticles

FITC (1 μg mL⁻¹) was incubated with nanoparticles (5 × 10⁷ nanoparticles per mL) in 100 μL of MilliQ® water at room temperature for 30 minutes. The conjugated particles were extracted by centrifugation at 1 × 10⁵ rpm for 5 minutes. 80 μL of the supernatant was removed and replaced with 80 μL of MilliQ® water. This washing procedure was carried out three times unless otherwise specified, and the fluorescence intensity of supernatant and re-suspended conjugated particles was measured using a CLARIOstar Plus plate reader after each wash. The excitation and emission wavelengths were set to 405 nm and 504 nm, respectively.

2.12 Detergent washing of FITC-conjugated PPNs

To confirm that the FITC molecules were covalently immobilized onto the nanoparticle surfaces, washing protocols were enlisted between measurements. Nanoparticles, initially washed with water for three times as described above, were then incubated for 1 hour in 100 μL of 5% sodium dodecyl sulfate (SDS) or 5% Tween20 detergent at room temperature. The solutions were centrifuged at 10 000 rpm for 5 minutes, after which the supernatants were decanted and reserved for analysis. The remaining conjugated particles were then washed with water as specified in each section followed by measurements of the immobilized fluorescent molecule.

2.13 Flow cytometry of FITC-conjugated PPNs (flow nanometry)

Nanoparticles (5 × 10⁷ nanoparticles per mL) were incubated in distilled water, DMSO or FITC (1 μg mL⁻¹) as described above. Nanoparticles were analysed using a Gallios Flow Cytometer (Beckman Coulter, Brea, CA, USA) located in the Bosch Institute Live Cell Analysis Facility (University of Sydney). The machine parameters were adjusted to reduce the size threshold, enabling the detection of PPNs. FITC fluorescence was assessed using fluorescent channel 1. Flow cytometry data was analysed using Flowjo version 10 and data analysed using Graphpad Prism.

2.14 Conjugation of Nile blue to plasma polymerized nanoparticles

Nile blue (N0766, Sigma-Aldrich) was dissolved in MilliQ® water to a stock concentration of 0.25 mM. Fresh nanoparticles were suspended in 10 mL MilliQ® water at an initial concentration of 3.76 × 10¹¹ PPN per mL. For NB conjugation, 3.8 mL of the diluted 0.25 mM NB solution were added to 3 mL of nanoparticles stock, achieving approximately 30 pg of NB per 10⁵ nanoparticles. The mixture was left to incubate on a rocker, covered in foil, at room temperature for 30 minutes. The samples were then centrifuged at 8000 rpm for 10 minutes and washed with 10 mL MilliQ® water. Detergent washing was achieved by mixing with 10 mL of 5% SDS solution for 30 minutes, followed by three water washes. After each step, the samples were centrifuged, and the supernatant was saved for XPS, ATIR-FTIR, or fluorescence measurements Nile blue (Ex: 624/13 nm, Emi: 678/17) using fluorescent black plate UV Greiner box with clear-bottom 96 well plates. Finally, for cellular uptake imaging, the above steps were repeated and the pellets were dispersed in 1 mL sterile PBS.

2.15 Cell culture assay

MCF-7 human breast cancer cells were maintained at 37 °C in an atmosphere with 5% CO₂ in minimum essential medium, 10% (v/v) fetal bovine serum (FBS), and 0.01 mg mL⁻¹ recombinant human insulin. Cell density was determined using a trypan blue dye exclusion assay. For the process of passaging and plating, detachment of the cells was carried out using 0.05% trypsin-EDTA (Invitrogen).

2.16 Confocal laser scanning microscopy

The internalization and intracellular behaviour of NB-PPNs were examined using confocal laser scanning microscopy. MCF-7 cells were seeded onto 1.5 cm coverslips in 24-well plates at a 250 000 cells per well in 500 μL. Nile blue conjugated PPNs were produced as mentioned above. After removing the medium, the cells were treated with NB-PPNs at a final concentration of 7500 PPNs per cell in the culture medium for 4 hours. Untreated cells served as the control. Subsequently, the solution was aspirated, and cells were washed with 200 μL PBS. Nuclei were stained with 200 μL of NucBlue™ live reagent (Hoechst 33342) (ThermoFisher Scientific, R37609) for 3 minutes, washed, and fixed with 10% neutral buffered formalin (Sigma-Aldrich) at room temperature. After a 10-minute incubation, cells were washed again and stained with ActinRed™ (ThermoFisher Scientific) followed by a 10-minute incubation protected from light. The solution was removed, cells were washed twice with PBS, stained cultures were stored at 4 degrees celsius in PBS until being mounted on slides for imaging. Confocal images were captured using a Nikon C2 confocal microscope (lasers: 405 nm, 488 nm, 561 nm, 640 nm. Detectors: 3 standard PMTs using filter blocks: DAPI/Cy5, FITC, TRITC). Image analysis was performed using ImageJ (version 1.53c, NIH) maximum intensity projections were generated, and mean fluorescence intensity of Nile blue associated with cells was determined by applying a binary selection mask of the actin cytoskeleton.

2.17 Statistical analysis

The data is presented as mean ± standard deviation (SD), unless otherwise stated, and was analyzed using Prism (version 9.5.1, Graphpad) or Excel (Microsoft 365 MSO, Version 2309 Build 16.0.16827.20278). Flow cytometry data was processed using Flowjo (TreeStar, Ashland, OR, USA), with statistical analysis conducted via Prism (version 9.5.1, Graphpad). Data shown is the mean ± standard error of the mean (n = 6 from 2 independent experiments) and data points were compared using a one-way ANOVA. Mean fluorescence intensity (MFI) at specific channels was compared using an ordinary one-way ANOVA with a Tukey post hoc test. Significance levels in figures are indicated as ns if P > 0.05, *P ≤ 0.05, **P ≤ 0.01, ***P ≤ 0.001, and highly significant if ****P ≤ 0.0001.

3 Results and discussion

3.1 Plasma polymerization of nanoparticles

Plasma polymerized nanoparticles (PPNs) were generated using a mixture of argon, nitrogen, and acetylene gases as the precursor (Fig. 1). The generated particles were collected in a 24-well cell culture plate positioned on the lower electrically floating electrode. To provide insights into the molecular species present during the plasma polymerization, optical emission spectroscopy was employed for real-time diagnostics. The observation of transitions in the spectral range of 385–395 nm unveiled molecular species like excited N₂ neutral molecules, N₂⁺ molecular ions, and CN radical molecules (Fig. 2a). These observations within the nitrogen-based spectrum highlight the role of the gas mixture in shaping the spectral signature and influencing the biocompatibility and formation of nanoparticles.^37,45

Fig. 2 PPN physical and surface chemical characterization panel showing (a) optical emission spectrum (OES) of the plasma, (b) XPS survey scan, (c) atomic concentrations of carbon, nitrogen and oxygen, (d) FTIR, (e) XPS C 1s high resolution spectra, (f) XPS area percentage of carbon bonding components, (g) EPR spectrum and (h) particle size distribution of the synthesised nanoparticles.

XPS was employed to examine the elemental composition of PPNs post-synthesis, showing strong signals from carbon (C 1s) and nitrogen (N 1s) (Fig. 2b), with atomic concentrations of 64.9%, and 29.9% respectively (Fig. 2c). Despite no oxygen in the initial gas mix, around 5.2 atm% was detected, likely due to oxidation after production when exposed to air, a process commonly known as autoxidation. This phenomenon occurs as reactive radicals interact with atmospheric oxygen, and it has been previously observed in plasma polymers deposited from similar gases in the form of coatings^50–53 and others generated from other precursor monomers.^50,54–56

Functional group analysis through ATR-FTIR was employed to further understand the molecular composition and surface functionalities of the synthesized nanoparticles. The FTIR spectra obtained for PPNs (Fig. 2d) revealed distinctive peaks associated with bending and stretching vibrations of C=C, C–O, C–N, and C–H from alkene, primary alcohol, amine, and methyl groups at 980–1630 cm⁻¹, along with C≡N stretching from nitrile functional groups at 2000–2500 cm⁻¹. Additionally, peaks corresponding to stretching vibrations of C–H, N–H, and O–H were observed in the 2800–3600 cm⁻¹ spectral region. The presence of these functional groups correlate well with the bonding species determined by peak fitting the C 1s XPS high resolution peak (Fig. 2e), indicative of the formation oxygen and/or nitrogen-containing species within the samples. Three primary carbon environments were identified with respective binding energies of 284.6 eV for C1: C–C/C–H, 286.5 eV for C2: C–N/C–O, and 288.0 eV for C3: C=O/N–C=O,⁵⁷ with approximately 68%, 26%, and 6% of the total carbon content, respectively (Fig. 2f). The identification of these bonds and functional groups reaffirms our earlier conclusions regarding the role of molecular nitrogen in the synthesis process, influencing both the chemical composition and surface characteristics of PPNs.^37,50 These results are in good agreement with the FTIR data (Fig. 2d), which indicate the presence of alkenes, alcohols, amines, methyl, and nitrile groups.

EPR spectroscopy examines materials with unpaired electrons like free radicals, unveiling details about their structure, bonding, and electronic properties through analysis of electron spin. The EPR spectrum of the PPNs affirms the presence of radicals within the PPNs (Fig. 2g). It exhibits a singular symmetric resonance peak positioned at 3510 G, featuring a calculated g-factor of 2.004, indicating unpaired electrons linked with radical compounds. During the plasma polymerization process, the fragmentation of the organic precursor leads to the initial formation of small nanometer-sized clusters through gas-phase chemical reactions.^58,59 These clusters grow into larger entities by coagulating and adhering together. Within this dynamic environment, radicals generated from the precursor fragmentation that are unable to form bonds with neighbouring species due to steric constraints become entrapped within the growing nanoparticles. This encapsulation of radicals is a key feature of the plasma polymerization process, contributing to the unique properties of the PPNs.

To assess the hydrophobicity of nanoparticles in solution, a tensiometer was employed to measure the water contact angle formed between water droplets and layers of nanoparticles deposited on silicon wafers. The minimal contact angle was measured at (23.8 ± 0.6)° which indicates low hydrophobicity and reflects the affinity of the nanoparticle surfaces to water. This observation aligns with the FTIR and XPS data, showing the presence of polar groups such as C–O and C–N, rendering the surfaces hydrophilic.

The size and charge of nanoparticles critically impact their function and stability. Smaller particles enhance cellular entry but face instability challenges, prone to aggregation. PPN size distribution shown in Fig. 2h indicates a uniform nanoparticle size (∼230 nm) with a polydispersity index of 0.094. SEM micrographs and a representative size distribution plot are shown in Fig. 3a. The SEM micrographs confirmed the PPNs' spherical shape, revealing smaller particles forming chains. Particle interaction with the environment was explored by varying pH, impacting size and charge. Size increased with pH (∼4.5–9.0), showing a slight drop from pH 3.7 to 4.5 (Fig. 3b), while the zeta potential gradually decreased from ∼47 mV to ∼35 mV (Fig. 3c). Due to the dissolution of CO₂ from the surrounding air, the MilliQ® water remains generally acidic. Consequently, the size and zeta potential of nanoparticles in MilliQ® water deviated slightly from the trend observed for nanoparticles in an NaCl solution, exhibiting a size of approximately 220 nm and a zeta potential of around 50 mV. Agglomeration can be explained by the classical Derjaguin–Landau–Verwey–Overbeak (DLVO) theory, which considers the interplay of van der Waals (vdW) attractive forces and electrostatic double layer (EDL) forces.⁶⁰ Agglomeration occurs when EDL forces are diminished relative to vdW forces. The surfaces of most nanoparticles contain titratable surface chemical groups (e.g., N₂, CH, COOH), which can be protonated by H⁺ (resulting in a positively charged surface at low pH) or deprotonated by OH⁻ (yielding a negatively charged surface at high pH). The isoelectric point (IEP), where particles become electrically neutral, is expected above pH 7, driving agglomeration at higher pH due to reduced repulsion. The stability of the particles depends on the storage medium and conditions. Our preliminary findings show that storing PPNs dry or in Milli-Q deionized water maintains a monodispersed size distribution with an average hydrodynamic size of <360 nm for up to 6 months.

Fig. 3 (a-i–iii) SEM images of PPNs, showing various populations of PPNs including singular unit spheres, coagulates of PPNs and long chain-like aggregates. (a-iv) A histogram plot, obtained from the SEM image shown in (a-iii) with n = 104, showing the size distribution of the PPNs. (b) Hydrodynamic size of PPNs as a function of pH obtained by DLS and (c) PPN zeta potential changes as a function of pH.

3.2 Covalent attachment of fluorescent molecules to plasma polymerized particles

In evaluating the capacity of PPNs to covalently bind fluorescent molecules, we used fluorescein isothiocyanate (FITC) and Nile blue (NB), well-established and widely recognized fluorescent compounds. FITC is reporter fluorescent molecule with reactive group (–N=C=S) known for its hydrophilic nature, its bright fluorescence intensity, and its prevalence over the past several decades for labelling proteins, drugs, cells, molecules, and polymers. NB is a targeting fluorescent molecule with high affinity to unsaturated free fatty acids with chain lengths of more than 16-carbon atoms,⁶¹ especially in tumour therapy.⁴⁹ Their attachment to nanoparticles is of paramount significance as it enhances particle functionality by enabling precise tracking, labelling, and real-time monitoring in various biomedical applications. Here, the radical-containing nanoparticles serve as a conducive nanocarrier for the direct binding of FITC and NB molecules, enabling their immobilization through a straightforward, one-step incubation process at room temperature.

To verify that FITC was covalently bonded to nanoparticles’ surfaces, nanoparticles incubated in FITC solution were rigorously washed with SDS or Tween20 and characterised using XPS and ToF-SIMS before and after the detergent wash. SDS and Tween20 are detergents that have a well-documented ability to remove physisorbed molecules, while retaining those robustly attached to a surface.^62–64 SDS, in particular, is an ionic detergent that disrupts physical interactions to unfold proteins⁶⁵ while leaving covalent bonds unaffected, rendering it suitable for assessing the attachment of macromolecules onto surfaces.⁶⁶ SDS disrupts the physical forces that underpin the physisorption of proteins and hence it is used to identify biological molecules that are covalently attached to surfaces.^67–69

XPS aimed to identify and quantify FITC bound to the surface by analyzing its sulfur component (Fig. 4a). The absence of sulfur signals in the XPS data (Fig. 4b) of plasma polymer nanoparticles before exposure to FITC (Fig. 4b(i), <0.15 at%, the XPS signal sensitivity) served as a robust baseline for comparison. Following incubation in FITC solution and subsequent water washing thrice, a sulfur atomic concentration of 0.56 ± 0.13 at% was measured (Fig. 4b(ii)). After Tween20 washing, a detection of 0.48 ± 0.12 at% sulfur was observed (Fig. 4b(iii)), suggesting covalent attachment of FITC molecules to the surface. SDS washing was excluded from XPS measurements due to its sulfur content, rendering sulfur ineffective for detecting residual FITC.

Fig. 4 (a) Chemical structure of FITC used in these experiments. (b) XPS sulphur S 2p high resolution spectra obtained from (i) plasma poylmerized nanoparticles and FITC-incubated nanoparticles after washing with (ii) water and (iii) Tween20. (c) ToF-SIMS spectra obtained from (a) plasma poylmerized nanoparticles (top) and FITC-incubated nanoparticles after washing with water (middle) and Tween20 (bottom). (d) Fluorescence intensities obtained from FITC-incubated nanoparticles and their corresponding supernatant solutions. (i) Samples were washed with MilliQ water and SDS. The SDS washing was carried out following the third water wash. (ii) Samples were washed for up to 6 times with MilliQ® water. (iii) Samples were washed 6 times with water and stored in MilliQ® water for 2 and 3 weeks before the fluorescence measurements. The measurement of samples stored for 2 and 3 weeks was conducted after they were washed three times with water. (e) Flow cytometric analysis of FITC-conjugated nanoparticles. Nanoparticles were incubated in the presence of distilled water, DMSO or FITC and washed with distilled water, SDS or Tween-20 and analysed using a Gallios flow cytometer. (i) Shows representative flow cytometric profiles of FITC fluorescence of nanoparticles from each treatment group. (ii) Shows the mean fluorescence intensity (arbitrary units) of nanoparticles from each treatment group. NS: not significant.

ToF-SIMS was used to detect molecular signatures unique to FITC, providing strong evidence on their covalent attachment to the nanoparticle surfaces. ToF-SIMS spectra of the PPNs before and after incubation in FITC solution are shown in Fig. 4c. The observed peak at m/z 333 corresponds to FITC molecules, while the other peaks originate from the plasma polymer structure. The nanoparticle surfaces retained a dominant m/z 333 peak after the Tween20 wash. The presence of fragments associated with FITC molecules on the surface after the Tween wash further validates the XPS results and presents additional evidence of robust immobilization.

To further validate the covalent conjugation of FITC fluorophores to PPNs, a comprehensive suite of fluorescence spectroscopy analyses was conducted. The fluorescence emission from both the supernatant and the resuspended FITC-conjugated PPNs was quantified in solution (excitation/emission: 405 nm/504 nm). PPNs conjugated with FITC (FITC-PPNs) were then subject to a systematic series of water washes followed by SDS detergent treatment (Fig. 4d). Following the SDS treatment, there was a decrease in fluorescence intensity observed in the resuspended FITC-conjugated PPNs. However, a signal, significantly higher than the control group remained, affirming the covalent binding of FITC molecules. The decrease in fluorescence intensity post-SDS washing is attributed to the removal of physically attached FITC molecules from the PPN surface, while the discernible signal that remains is attributed to a persistently attached covalent monolayer. Furthermore, Fig. 4d(ii) shows the fluorescence intensities of nanoparticles following a succession of water washes. The appearance of fluorescence in the supernatant after each wash indicates the stepwise removal of consecutive layers of physically-adsorbed FITC. Despite this removal, the fluorescence signals from the nanoparticles remained unwavering, substantiating the robustness of the covalently bound FITC molecules to the PPN surfaces. Moreover, Fig. 4d(iii) illustrates the enduring stability of FITC-labeled nanoparticle fluorescence during extended storage in water for up to 3 weeks following 6 water washes. This extended stability corroborated the enduring covalent attachment of FITC to the nanoparticle surfaces, underscoring its potential for sustained performance in various applications requiring steadfast and persistent fluorescence properties.

The fluorescence intensity of FITC-conjugated nanoparticles was also assessed by flow cytometry. Adjusted to a detection threshold suitable for PPNs, flow cytometry allows for large-scale analysis of single nanoparticles, referred to as nanometry.⁴⁵ FITC is typically dissolved in aqueous buffer solutions or organic solvents, such as dimethyl sulfoxide (DMSO). PPNs were incubated in FITC, distilled water or DMSO to provide a variety of carrier solutions for evaluation. The conjugated particles were then washed in distilled water, SDS or Tween-20 and analysed for fluorescence with 488 nm excitation using a Gallios flow cytometer. Fig. 4e(i) shows representative flow cytometry data from this study. Nanoparticles incubated in distilled water or DMSO showed no fluorescent signal above background. Only FITC-conjugated nanoparticles demonstrated significant increase in fluorescence in the FITC detection range. Fig. 4e(ii) illustrates that while the FITC-conjugated nanoparticles showed a statistically significantly higher FITC fluorescence than distilled water or DMSO-treated nanoparticles, there was no significant difference in mean fluorescence intensity between FITC-conjugated nanoparticles washed with distilled water, Tween-20 or SDS, which demonstrates the covalent attachment of FITC molecules that are not removed with a simple water wash. The ease of removal of physically adsorbed FITC can be attributed to its small molecular size which limits the combined strength of the physical interactions relative to large physiosorbed molecules such as proteins. These results align with the conclusion of FITC covalent bonding obtained from XPS, ToF-SIMS and fluorescence spectroscopy by demonstrating consistent fluorescence intensity emitted from FITC-conjugated nanoparticles after different detergent washing procedures.

To ensure that the results observed for FITC are not specific to this particular molecule, we also attached Nile blue, a cationic benzophenoxazine fluorescent molecule with affinity to unsaturated free fatty acids, onto the PPNs. Nile blue comprises carbon, hydrogen, nitrogen, and oxygen elements intricately arranged within its phenoxazine ring (Fig. 5a). The molecule incorporates diverse substituents and functional groups strategically positioned to impart its characteristic blue hue. These structural features, including side chains, contribute to the fluorescent molecule's solubility, reactivity, and suitability for applications such as fluorescence and biological staining. This sequential approach allowed us to comprehensively understand the covalent binding process, shedding light on the distinct characteristics and applications of each functionalized nanoparticle variant.

Fig. 5 Surface characterization of the PPN and PPN-NB showing successful conjugation of Nile blue into the PPN. (a) NB chemical structure. (b) Spectral analysis of the XPS survey spectra indicating changes in carbon, nitrogen, and oxygen atomic percentages, (c) peak fitted high-resolution C 1s peak of PPN and PPN-NB attributing three bonding components (C–C/C–H, C–O/C–N, and C=O/N–C=O), (d) percentage of each C 1s components based on peak area, and (e) Fourier transform infrared (FTIR) spectra highlighting changes in the functional groups present in the PPN surface after conjugation.

The attachment of Nile blue onto the PPN surface was investigated through XPS and ATR-FTIR. Changes in the elemental composition of PPN-NB were evident following the conjugation and detergent treatment process as observed in Fig. 5b. Specifically, the oxygen content increased from 5.4% to 13.4%, whereas nitrogen content decreased from 26.3% to 22.9%. Post-conjugation, XPS analysis shows a decrease in nitrogen levels, likely due to Nile blue's lower nitrogen-to-carbon ratio compared to the as-synthesized nanoparticles. These changes in surface composition can be primarily attributed to the immobilisation of NB and the autoxidation of the PPN surface as discussed in Section 3.1.

The high-resolution C 1s peaks of the PPNs before and after conjugation were investigated to provide further evidence of NB attachment. In terms of peak shape, the PPN-NB exhibits broadening at the high binding energy (BE) tailing end compared to the unconjugated PPN (Fig. 5c). Three distinct carbon peaks corresponding to C1: C–C/C–H (BE ≅ 285.0 eV), C2: C–O/C–N (BE ≅ 286.5 eV), and C3: C=O/N–C=O (BE ≅ 288.2 eV) were fitted to the C 1s spectra with the best fit area percentages shown in Fig. 5d. The C2 peak, corresponding to C–O and C–N bonding species, showed an increase in area percentage from 29.2% to 41.0% after NB conjugation (Fig. 5d). This increase that accounts for the observed C 1s peak broadening is attributed to the presence of primary, secondary, and aromatic amine groups in the NB structure. The rise in C–O species originates from the oxygen atom present in the aromatic structure of NB, as well as the reaction of atmospheric oxygen with carbon-based radical species on the nanoparticle surfaces. Conversely, the predominant C1 peak (C–C/C–H) decreased in abundance from 64.5% to 52.3% which is a direct result of the C2 increase. Lastly, the C3 peak (C=O/N–C=O) exhibited a slight increase from 6.4% to 6.8%. This change may be attributed to the formation of more stable oxidation products such as carbonyl (C=O) and carboxylate (COOH) functionalities due to a chain of auto-oxidation reactions initiated at the carbon-centered radicals at the PPN surface. This is consistent with the oxidative degradation mechanisms previously observed in plasma polymers in the form of coatings of similar composition, where oxygen and water molecules diffusing into the polymer matrix react with metastable species, leading to an increase in oxygen-containing functional groups.⁵⁷ The decrease in nitrogen content observed could be explained by the volatilization and leaching of nitrogen-containing compounds from the polymer matrix as well as by the lower nitrogen to carbon ratio of the NB molecules compared to that in the PPNs. The autooxidation is likely accelerated by hydrolysis reactions targeting nitrogen-containing groups within the polymer, further exacerbated by the oxidative environment. The decrease in nitrogen content upon exposure and conjugation with Nile blue, a benzophenoxazine fluorescent molecule, could also be contributed to by the oxidative degradation of the nitrogen-containing heterocyclic structures within the fluorescent molecule itself or the plasma polymer matrix. In the presence of oxidative stress, such as exposure to atmospheric oxygen or reactive oxygen species in aqueous environments, the nitrogenous heterocyclic rings can undergo oxidative cleavage, leading to the breakdown of these structures and the consequent release of nitrogen in forms that are more volatile or soluble, thereby reducing the overall nitrogen content measurable on the surface of the plasma polymerized nanoparticles. This degradation process is consistent with the observed increase in oxygen content, suggesting that oxidative reactions are a significant pathway for the alteration of nitrogen-containing components in the polymer-molecule conjugate system. As the XPS measurements were conducted within two days of PPN production, the observed nitrogen loss and oxidative changes can be attributed to the conjugation process and early-stage oxidative reactions, rather than prolonged exposure to aqueous media.

Functional group analysis of NB conjugation was conducted using ATR-FTIR on both pre- and post-conjugated PPN samples. Distinct amine peaks emerged in the FTIR spectra of PPN-NB (Fig. 5e). A secondary shoulder peak at 1587 cm⁻¹ is identified and attributed to N–H bending from the primary amine group within the NB. Minor peaks at 1331 and 1271 cm⁻¹ are ascribed to C–N stretching originating from the aromatic amine present in the heterocyclic ring of NB. Additionally, multiple C–N stretching peaks at 1211, 1175, 1142, 1118, and 1076 cm⁻¹ are also observed, correlating with various amine groups (primary, tertiary, and aromatic) distributed across the NB structure. The other peaks found in the FTIR spectra of PPN-NB overlap with the spectra of the PPN, and all peak assignments are summarized in Table 1. Overall, these changes in the FTIR spectra of the PPN post-conjugation indicates the successful covalent attachment of NB onto the PPN surface. The successful covalent binding of NB to PPNs, supported by XPS and ATR-FTIR, aligns with the conclusions drawn from FITC conjugation. Both fluorescent molecules exhibited changes in elemental composition on the PPN surface and robust bonding post-conjugation and detergent washing. The results presented in this section highlight the robustness and versatility of the PPN surface for the covalent attachment of various fluorescent molecules.

Table 1 Functional group assignment for the ATR-FTIR spectra of the unconjugated PPN and PPN-NB

Frequency range (cm^−1)	Type of vibration	Functional group	Presence in FTIR spectra (Y/N)
			Unconjugated PPN	PPN-NB
3490	O–H stretching	Alcohol	Yes	Yes
3357	N–H stretching	Primary/secondary amine	Yes	Yes
2958	C–H stretching	Alkane	Yes	Yes
2934	C–H stretching	Alkane	Yes	Yes
2858	C–H stretching	Alkane	Yes	Yes
2195	C≡N stretching	Nitrile	Yes	Yes
1624	C=C stretching	Alkene	Yes	Yes
1587	N–H bending	Amine	No	Yes
1448	C–H bending	Methyl	Yes	Yes
1376	C–H bending	Methyl	Yes	Yes
1330	C–N stretching	Aromatic amine	No	Yes
1271	C–N stretching	Aromatic amine	No	Yes
1211	C–N stretching	Amine	Yes	Yes
1175	C–N stretching	Amine	No	Yes
1146	C–N stretching	Amine	No	Yes
1118	C–N stretching	Amine	No	Yes
1077	C–N stretching	Amine	No	Yes
1059	C–O stretching	Primary alcohol	Yes	Yes
980	C=C bending	Alkene	Yes	Yes

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3.3 Intracellular cytoplasmic uptake of Nile blue conjugated nanoparticles

To exemplify one among numerous potential applications of PPNs covalently functionalized with fluorescent molecules, we investigate Nile blue-conjugated nanoparticles for drug delivery. A significant challenge in targeted drug delivery lies in enhancing the cellular uptake of targeting molecules. Current strategies predominantly focus on extracellular delivery through cell surface receptors, where the drug is subsequently released. However, this approach is suboptimal for drugs which exert their effects on intracellular targets. To overcome this limitation, engineering nanoparticles specific to the cell and incorporating elements that promote cytoplasmic uptake becomes crucial. Here, Nile blue was used as a potent inducer for cellular uptake, with the aim of achieving the direct delivery of therapeutic payloads into the cell's interior.

We investigated the cellular uptake of Nile blue-conjugated polymeric nanoparticles (NB-PPNs) in MCF7 breast cancer cells. In our control group, untreated cells and those treated with unconjugated PPNs were examined at both low and high magnifications. No observable gross morphological differences were observed, as depicted in Fig. 6a and b. Unconjugated PPNs displayed no detectable fluorescence signal in the far-red channel. Conversely, NB-PPN treated cells exhibited a robust Nile blue signal, indicating successful cellular uptake (Fig. 6c). We also conducted a detailed qualitative analysis at higher magnification, revealing the observed uptake of Nile blue within the cells. Upon closer examination, Nile blue stains were visibly present in the cytoplasm of the cells. Notably, the fluorescence signals emanated from both small aggregates and diffusely distributed stain throughout the cells (Fig. 6d).

Fig. 6 Following incubation, fixation, staining, and imaging with confocal microscopy, MCF-7 cells internalize PPNs conjugated with the lipophilic Nile blue probe and washed with detergent. Pseudocoloring of the images is performed using ImageJ, with actin appearing yellow, representing cytoskeletal F-actin; nuclei stained by NucBlue in grayscale; and Nile blue in blue. Three conditions are compared: (a) untreated cells without PPNs, (b) cells treated with non-conjugated PPNs, and (c) cells treated with Nile blue-conjugated PPNs. Merged images of all channels under 40× magnification with 2× image zoom and 100× magnification are presented in the large panels on the right. Smaller panels on the left display individual channels for actin red, NucBlue, and Nile blue, along with a transmitted channel. Additionally, (d) exhibits two representative images from PPN-NB treated cells at 100× magnification, focusing on the 660 nm fluorescent signal for the Nile blue channel. The images reveal small aggregates (circles) and diffusely distributed stain (arrows) throughout the cells. (e) Provides quantification of mean fluorescence intensity (MFI), indicating a significant Nile blue fluorescence signal in cells treated with PPN-NB. The images were captured using a Nikon C2 confocal microscope with a 40× dry and a 100× oil immersion objective and analyzed using ImageJ.

To quantify the cellular uptake of NB-PPNs and evaluate the distribution of Nile blue within MCF7 cells, we employed a quantitative approach by calculating the mean fluorescent intensity (MFI). The resulting plot in Fig. 6e shows individual data points representing MFI in the cells, both with and without NB-PPN treatment. A significant Nile blue signal was detected following NB-PPNs treatment, contrasting the negligible signal observed in the negative control (Fig. 6e).

The confocal images revealed that the PPNs covalently conjugated with NB were successfully internalized by cells into the cytoplasm while maintaining their integrity, avoiding degradation and cellular deformation. They effectively stained the fatty acids, which served as the intracellular target and achieved intracellular localization of NB-PPNs. This observation can be well explained, given Nile blue's peak fluorescence in the presence of unsaturated free fatty acids commonly found in the cytosol. The diffuse cellular staining highlights the uptake of nanoparticles into the cytoplasm, distinguishing it from lysosomal entrapment which is typically characterized by compact compartments within the cytoplasm.⁷⁰ Moreover, the discrete NB-bright fluorescent aggregates suggest the proximity of NB-conjugated nanoparticles to accumulations of free fatty acids. By covalently conjugating a lipophilic fluorescent dye, we efficiently transported the particles into the cellular cytoplasm, avoiding lysosomal entrapment or peripheral adhesion. While untreated cells with NB alone show high internalization, the NB-PPN system offers significant advantages in targeted intracellular delivery and localization. Conjugating NB to plasma polymer nanoparticles improves control over quantification, delivery, and retention, enhancing stability and fluorescence signal over time. This is crucial for prolonged imaging or tracking studies where free dye may diffuse or degrade. The NB-PPN system effectively delivers nanoparticles into the cytoplasm, avoiding lysosomal entrapment, and targets intracellular compartments rich in unsaturated fatty acids, where NB's fluorescence is maximized, as shown in Fig. 6d. This result aligns with our previous findings, where NB-conjugated magnetic microparticles demonstrated comparable effects on cellular uptake and staining.⁷¹ While NB-conjugated microparticles can be utilized for cell filtering or isolating cancer cells in peripheral lymphatic cancers, their potential for therapeutic delivery is hindered by their metallic nature. In contrast, PPNs can not only achieve lipophilic staining though the covalent attachment of NB, but also offer a biocompatible carrier with no cytotoxic effects, as was demonstrated in our previous cytotoxicity studies.^41,45 Our preliminary results indicate that the PPNs remain reactive for covalent biofunctionalization beyond a period of one year, suggesting their potential for long-term applications in both diagnostic and therapeutic settings. Further studies are underway to fully understand the mechanisms that contribute to this prolonged surface activity. The stability of the particles depends on the storage medium and conditions. Our recent findings show that storing PPNs in Milli-Q deionized water maintains a monodispersed size distribution with a hydrodynamic size of <300 nm for up to one year. Future directions may also involve optimizing theranostic capabilities by co-immobilizing a drug and a cellular targeting agent with the lipophilic probe. This strategic approach creates a system where one element guides the particles, while the other enhances their cytoplasmic uptake for efficient drug distribution. The findings presented in this study advance plasma-polymerized nanoparticles as versatile platforms in nanomedicine, enabling direct covalent attachment of fluorescent molecules without the need for additional reagents or steps, thereby broadening their utility across medical and non-medical applications.

4 Conclusions

This study investigated plasma polymerized nanoparticles (PPNs) synthesized through radical-induced plasma polymerization, exhibiting well-defined physical and chemical characteristics. The covalent attachment of biofunctional molecules, including fluorescein isothiocyanate (FITC) and Nile blue (NB), was confirmed through its resistance to rigorous washing, validated by XPS, ToF-SIMS, and fluorescence spectroscopy. The successful intracellular uptake of Nile blue-conjugated PPNs (NB-PPNs) was demonstrated in MCF7 breast cancer cells, showing the potential for PPNs for cell entry, targeted drug delivery and their biocompatibility. The versatility and robust synthesis of plasma polymerized nanoparticles (PPNs) shows that this process not only facilitates surface functionalization but also allows robust covalent conjugations with biofunctional molecules in a single incubation step, minimizing preparation steps and eliminating the need for particle surface modification. This study sets the stage for advancements in employing PPNs as effective carriers in biomedical applications.

Data availability

The authors declare that all data supporting the findings of this study are available within the article or from the corresponding author upon request.

Conflicts of interest

There are no conflicts to declare.

Acknowledgements

The authors would like to thank the Australian Research Council (ARC) for funding this work under the DECRA (Discovery Early Career Researcher Award) (DE210100662) and Laureate (FL190100216) programs. The authors also express their appreciation for the tools, advice from scientists, and technical assistance provided by Microscopy Australia.

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