Targeted cancer theranostics using a plasmonic gold nanohybrid assembly of chiral ligand stabilized nanorods and protein nanoclusters

Abstract

The integration of discrete nanomaterials into a single platform enables the synergistic combination of their exceptional physicochemical properties, which is particularly advantageous for biomedical imaging and therapy. However, retaining the intrinsic optical features of individual components during such integration remains a major challenge. In particular, understanding the interactions between metallic nanostructures with distinct functionalities continues to be of significant scientific interest. In this study, we report the design of a novel hybrid gold nanoplatform, PEG–PGNC@HA–GNR, comprising DSPE–PEG coated protein gold nanoclusters (PGNCs) and hyaluronic acid (HA) wrapped anisotropic gold nanorods (GNRs). The PGNCs exhibit tunable near-infrared (NIR) photoluminescence, while GNRs provide efficient photothermal property. The hybrid nanoarchitecture was meticulously engineered to preserve these complementary optical features, enabling dual functionality for imaging and photothermal therapy. HA, a chiral anionic polysaccharide with high affinity for CD44 receptors, simultaneously enhanced GNR biocompatibility and induced plasmon driven chiroptical activity via helical wrapping. DSPE–PEG coating on PGNCs improved stability and facilitated their conjugation with HA–GNRs, yielding a hybrid nanostructure validated by UV-Vis and fluorescence spectroscopy, zeta potential analysis and TEM. The hybrid retained significant PGNC fluorescence despite conjugation with GNRs, largely due to deliberate spatial separation minimizing energy transfer. The resulting construct displayed distinct circular dichroism (CD) signals and enhanced photothermal performance, offering a multifunctional platform for targeted cancer recognition and light-triggered therapy. Preliminary in vivo studies further demonstrated its imaging potential in mice. This work underscores the utility of naturally derived chiral ligands in engineering multifunctional plasmonic nanomaterials for precision oncology.

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1. Introduction

Ultrasmall fluorescent nanomaterials with unique physicochemical properties are highly beneficial for biomedical applications.^1–3 In particular, atomically precise noble metal nanoclusters exhibit a high fluorescence quantum yield, large Stokes shift, excellent photostability, inherent biocompatibility,^4,5 which collectively contribute to their effectiveness as imaging agents, biosensors and drug delivery platforms. Recent advancements in synthetic strategies have further enhanced their tunability and functional versatility, positioning gold nanoclusters (GNCs) as promising candidates for clinical translation in diagnostics and therapeutics.^6–10 Among various synthesis approaches, green synthesis using protein-based stabilizers such as Bovine Serum Albumin (BSA), horseradish peroxidase (HRP), papain, soy protein, eggshell protein and egg white protein has gained significant attention.^11,12 Proteins offer abundant functional groups that serve as scaffolds for nanocluster formation.¹³ Ovalbumin, a glycoprotein with a molecular mass of approximately 45 kDa, contains one disulfide bond and four free sulfhydryl groups embedded within a hydrophobic core, making it a promising stabilizing and reducing agent for gold nanocluster synthesis.¹⁴

It has been reported that highly fluorescent gold nanoclusters synthesized using protein as a dual-function reducing and stabilizing agent, display excellent performance as a selective and sensitive sensor for Hg²⁺ ions.¹⁵ Recently, bright-red-emitting ovalbumin-protected gold nanoclusters were applied as a luminescent probe for highly sensitive determination of cyanide ions (CN⁻ ions).¹⁶ Li et al. reported the use of ovalbumin-stabilized gold nanoclusters modified with polyethyleneimine for the fluorescent detection of tetracyclines.¹⁷ Protein derived gold nanoclusters (PGNCs) exhibit low toxicity, stable photoluminescence, long fluorescence lifetime, strong anti-bleaching properties, and excellent water solubility.^18,19 Despite these favorable properties, the utilization of PGNCs in bioimaging remains largely underexplored indicating a substantial opportunity for further research in this area. In this study, gold nanorods (GNRs), known for their strong NIR optical absorption and excellent photothermal properties,^20,21 were conjugated with DSPE–PEG coated PGNCs to integrate the bioimaging capabilities with photo thermal therapy (Scheme 1).

Scheme 1 Schematic representation of the synthesis and application of PEG–PGNC@HA–GNR.

This integration can further promote synergetic effects by enhancing photothermal performance, facilitating the generation of plasmonic hot spots and improving stability in biological environments thereby advancing its potential for combined therapeutic and imaging applications.²² Beyond their intrinsic optical properties, recent studies have demonstrated that GNRs can exhibit chiroptical activity when functionalized with chiral bio-macromolecules and other polysaccharide-based polymers.²³ The induced chirality arises from the asymmetric arrangement of the polymer chains around the nanorod surface, leading to plasmon-induced circular dichroism (CD) signals. The chiral assembly not only imparts additional optical functionalities, but also influences interparticle interactions and colloidal stability.²⁴

Herein, we have modified gold nanorod surface with hyaluronic acid (HA), which can reduce the toxicity of the GNRs by removing CTAB. HA is an anionic, biocompatible polysaccharide composed of D-glucuronic acid and N-acetyl-D-glucosamine.²⁵ It exhibits a high binding affinity for CD44, a cell surface receptor that is overexpressed in various cancer types and closely associated with tumorigenesis, invasion, and metastasis. This specific interaction has positioned HA as a valuable targeting ligand in the design of nanocarriers for cancer therapy.²⁶ Beyond its targeting capability, HA is a naturally chiral polysaccharide whose helical conformation can induce chiroptical effects when conjugated to plasmonic nanoparticles such as gold nanorods.²⁷ The asymmetric wrapping of HA around GNRs can enhance their colloidal stability and biocompatibility and also impart plasmon-induced circular dichroism (CD) signals, enabling the development of chiral plasmonic nanomaterials. When integrated into hybrid nanostructures with nanoclusters, these HA functionalized chiral nanomaterials contribute to enhanced photothermal properties and unique optical responses, offering a multifunctional platform for targeted, image-guided, and light-responsive cancer therapy.

2. Results and discussion

2.1 Synthesis and characterization

Protein stabilized gold nanoclusters (PGNCs) were synthesized initially as per the protocol given in Section 4.2. To improve biocompatibility and to enable further surface functionalization, PGNCs were encapsulated with amphiphilic polymer DSPE–PEG with a terminal NH₂ group. Furthermore, in order to develop a nanohybrid system of PGNCs and GNRs entitled as PEG–PGNC@HA–GNR, GNRs were synthesized as per the optimized protocol (see Section 4.3). The growth mechanism and growth rate of the GNRs vary on different facets, mediated by the difference in the stability of adsorbed CTAB resulting in a diverse array of morphologies from slender sphero-cylinders formed at early reaction stages to more evolved structures such as rods with a dumbbell profile, flattened end facets and octagonal prismatic structures.²⁸ The GNRs were then coated with HA to reduce the toxicity and to impart CD44 targeting ability to the integrated system. Furthermore, HA is also expected to contribute to the preservation of fluorescence of the hybrid system to some extent by serving as an effective spacer thereby restricting the energy transfer interactions between the GNRs and PGNCs, so that the final system will retain the fluorescence.²⁹ Finally, the hybrid nanomaterial was synthesized by EDC coupling between the amino group of the DSPE–PEG coated PGNCs with the –COOH terminal of the HA-coated GNRs leading to a covalent amide linkage (–CO–NH–) to form a stable hybrid system PEG–PGNC@HA–GNR (Scheme 2).

Scheme 2 EDC/NHS coupling reaction between HA and DSPE–PEG.

The UV-Vis spectral profile of the PGNCs exhibited distinct absorption features characteristic of atomically precise gold nano clusters, while notably lacking the surface plasmon absorption band around 520 nm corresponding to the oscillations of conduction electrons responsible for SPR in larger nanoparticles (Fig. S1A, SI). Moreover, the fluorescence arising from the quantization of energy state at the atomic size of the clusters further substantiated the successful formation of the same. The emission spectra of the PGNCs upon excitation at 400 nm displayed two prominent peaks, a moderate emission around 460 nm along with an intense band around 668 nm (Fig. 1(A)). Unlike bulk metals or larger nanoparticles, nanoclusters possess quantized energy states and the excitation wavelength selectively promotes electrons to different excited states or energy manifolds. These excited states can undergo radiative relaxation via distinct emissive pathways, each associated with different emission energies. Additionally, factors such as ligand-to-metal charge transfer, metal-centered transitions, and surface state emissions can contribute to multiple emissive channels within the same cluster framework. The relative population of these states depends on the excitation energy, leading to variations in the emission profile. The appearance of the emission peak around 460 nm and its excitation-dependent shift reflects the complex electronic structure of the PGNCs and the wavelength-selective activation of specific emissive states within the cluster. The excitation-dependent shift in the emission spectra is common for several protein or peptide-based metal nanoclusters.³⁰ This observation is attributed to the influence of excitation energy on the electronic states and relaxation pathways, leading to ligand to metal energy transfer and associated emission peaks. As we are interested in the imaging potential of the metal nanocluster, the stable red emission centered at 668 nm was deemed most relevant for this study; hence the emission band around 450 nm was not considered. The Quantum yield (QY) corresponding to the 680 nm peak was calculated to be approximately 10%, using Rhodamine 6G as a reference standard (SI). The emission of the PGNC under 365 nm UV light irradiation was also visible to the unaided eye (Fig. S1B, SI).

Fig. 1 (A) Fluorescence emission spectra of PGNCs recorded at different excitation wavelengths. (B) UV-Vis NIR spectra of GNRs, HA–GNR and PEG–PGNC@HA–GNR. (C) Fluorescence emission spectra of PGNCs, PEG–PGNC, and PEG–PGNC@HA–GNR. (D) Photograph showing the fluorescence from (1) water, (2) PGNCs, (3) PEG–PGNC, and (4) PEG–PGNC@HA–GNR, respectively. (E) Zeta potential values of different materials and (F) IR spectra of HA–GNR and PEG–PGNC@HA–GNR.

The GNRs showed characteristic longitudinal and transverse absorption peaks around 830 and 530 nm (Fig. 1(B)), respectively. On HA coating, the longitudinal absorption band of GNR was significantly red shifted from 830 nm to 850 nm, which is attributed to a reflection of the ligand exchange process (Fig. 1(B)). When HA is introduced, it can partially displace the loosely bound outer CTAB layer, especially on the less tightly bound end facets. According to Gans theory, the longitudinal surface plasmon resonance (LSPR) of a metal nanorod is highly sensitive to the refractive index of its immediate surroundings. Adding a HA layer effectively increases the refractive index around the GNR, which reduces the energy of their longitudinal plasmon oscillations, causing the LSPR band to red-shift.³¹ Furthermore, after conjugation with DSPE–PEG encapsulated GNCs, the LSPR absorption band of HA–GNR undergoes a hypsochromic shift with considerable broadening (Fig. 1(B)). This spectral alteration is attributed to the combined effects of changes in the local dielectric environment, plasmonic coupling between the GNRs and proximal GNCs and surface modification induced electron density perturbations. The emission maximum of PEG–PGNC showed a small blue shift of 4 nm with an additional shoulder band around 632 nm (Fig. 1(C)). Meanwhile, the fluorescence intensity of PEG–PGNC was slightly decreased after conjugation with HA–GNR, though largely preserving the properties (Fig. 1(C)). This preservation of fluorescence can be attributed to the DSPE–PEG and HA coating, which act as a spacer maintaining an appropriate distance between the PGNCs and GNRs. This spatial separation minimizes the energy transfer interactions, thereby preventing significant quenching of fluorescence, as generally expected for fluorophores like GNCs in the proximity of GNRs. The quantum yield of the hybrid nanomaterial was calculated to be approximately 3.5%, using Rhodamine 6G as a reference standard (SI).

Successful PEGylation of PGNCs was also confirmed by changes in zeta potential analysis. While the unmodified PGNCs exhibited a negative surface charge with a zeta potential of −21.0 mV, with DSPE–PEG encapsulation, the zeta potential decreased to −25.1 mV, indicating effective surface modification (Fig. 1(E)). The GNRs displayed a positive zeta potential of +31.3 mV, attributed to the positively charged CTAB bilayer on the surface. Therefore, HA coating on the GNR surface could be achieved based on the electrostatic attraction between the positively charged CTAB and the negatively charged HA. A corresponding decrease of zeta potential from +31.3 mV to −22.2 mV was observed on HA coating. PEG–PGNC@HA–GNR exhibited a zeta potential of −15.8 mV, confirming an efficient surface modification of the GNRs. In the IR spectra, HA–GNR showed all the characteristic bands of HA. The –OH stretching vibration is around 3500 cm⁻¹ and the sharp C=O stretching band at 1600 cm⁻¹ of HA is present in the IR spectra of GNR after HA coating (Fig. 1(F)). In the IR spectra of PEG–PGNC@HA–GNR a strong sharp band at 1660 cm⁻¹ confirms the amide bond formation.

The PEGylation of the PGNCs hasn’t altered the dispersed pattern of the clusters in the colloid as visible in the TEM image. However, well dispersed individual clusters are seen within the polymer matrix (Fig. 2(A)).³² The TEM images of the PGNCs revealed that the gold nanoclusters are well-dispersed and predominantly spherical in shape, with an average diameter ranging from 2 to 3 nm (Fig. S2, SI). The GNRs showed rod-like to mildly dumbbell-shaped particles in the TEM image, with a minor fraction of off-shaped particles (including occasional spherical seeds). These are likely due to the result of different growth phases during synthesis and are not expected to significantly influence the overall properties (Fig. 2(B) and Fig. S3). HA-coated gold nanorods confirmed that the structural integrity of the nanorods was preserved,³³ as evidenced by the unaltered aspect ratio following hyaluronic acid functionalization (Fig. 2(C)). In the TEM image of the final hybrid material, the PGNCs were clearly visible around the GNRs, which confirms the successful formation of the hybrid material. A distinct separation of the PEG–PGNCs from the HA–GNRs is also very clear from the TEM image, which explains the preservation of the fluorescence of GNCs in the hybrid system. In the hybrid system, the PGNCs maintain their well disbursed appearance and do not show a direct interaction with the HA–GNRs, which further confirms the EDC/NHS coupling between the ligands HA and DSPE–PEG (Fig. 2(D)).

Fig. 2 TEM images of (A) PEG–PGNC, (B) GNRs, (C) HA–GNR and (D) PEG–PGNC@HA–GNR. Red arrows indicate the well dispersed PGNCs (scale bar of A is 5 nm, and that of B, C and D is 10 nm). (E) Raman spectra of HA–GNR and GNRs. (F) XRD pattern of GNR and HA–GNR.

The successful functionalization of GNRs with HA was further validated by Raman spectroscopy (Fig. 2(E)). The HA–GNR spectrum displayed a series of well-defined and intense peaks in contrast to the GNR spectrum, which shows only weak and broad signals. The HA–GNR spectrum exhibits prominent peaks that are characteristic of hyaluronic acid. Specifically, the band at 752 cm⁻¹ is attributed to C–C or C–O skeletal vibrations associated with the polysaccharide backbone. The amide I band, typically observed around 1600 cm⁻¹, results from C=O stretching. The band at 1135 cm⁻¹ is indicative of C–C stretching and CH₂ twisting modes. The peaks observed at 1384 and 1424 cm⁻¹ are assigned to symmetric CH₃ bending and CH₂ deformation modes, respectively. GNRs exhibit distinct XRD peaks at ∼38°, 44° and 64° corresponding to the (111), (200), and (220) planes of fcc Au (Fig. 2(F)). HA–GNR shows changes in the peak intensities, retaining the peak positions, suggesting the successful surface modification without altering the core crystal structure of the GNRs.

Dynamic light scattering analysis was performed to evaluate the hydrodynamic size distribution of the synthesized nanostructures. Fig. S6, SI represents the size distribution of the GNRs, showing a narrow size distribution (z-average hydrodynamic diameter 207.4 d.nm) with polydispersity index (PDI) 0.442 indicating good colloidal stability and uniformity of the GNRs. The Fig. S7, SI corresponds to the DLS profile of the PEG–PGNC@HA–GNR. A noticeable shift in the hydrodynamic diameter is observed (z-average hydrodynamic diameter 215.2 d.nm), with PDI 0.417, which can be attributed to successful conjugation and possible aggregation or surface interaction between the nanorods and nanoclusters. This increase in size confirms the formation of a hybrid nanostructure, which is consistent with the expected outcome of the conjugation process. The results collectively support the structural transformation and functionalization of GNRs with nanoclusters.

2.2 Photothermal therapy

The strong absorption of the GNRs in the IR region was explored for temperature generation efficacy of the hybrid nanomaterials for its use in photothermal therapy application. For this, GNRs, HA–GNR, PEG–PGNC@HA–GNR were irradiated with an 808 nm laser (2 W cm⁻²) and the corresponding temperature changes with time were recorded (Fig. 3(A)).

Fig. 3 (A) Photothermal heating profiles of GNRs, HA–GNR and PEG–PGNC@HA–GNR. (B) Thermographic images of PEG–PGNC@HA–GNR at different concentrations with time. (C) Thermal stability profile of PEG–PGNC@HA–GNR during five consecutive cycles of laser exposure. (D) Circular dichroism (CD) spectra of the GNRs, HA and HA–GNR.

The GNRs exhibited a steady rise in temperature over time reaching a maximum of 46 °C within 12 minutes of irradiation. Upon HA coating, a minor enhancement in photothermal performance was observed, with an increase of 3 °C for the same duration. Furthermore, after the hybrid formation with PGNC conjugation, PEG–PGNC@HA–GNR retained the photothermal properties and demonstrated a concentration-dependent increase in temperature upon laser irradiation (Fig. 3(A), (B) and Fig. S8, SI). Notably, the system also exhibited excellent photothermal stability. Even after five consecutive irradiation cycles, no significant reduction in the photothermal performance was observed, underscoring the thermal robustness and reusability of the composite material (Fig. 3(C)). The photothermal conversion efficiency (η) of the GNRs and HA–GNR was calculated as 25.6% and 27.86%, respectively (Fig. S9 and S10, SI). The observed enhancement in the photothermal performance of HA–GNR compared to the bare GNRs was unexpected, as surface coatings are typically anticipated to attenuate efficiency by introducing additional thermal resistance. To explore this anomaly, we investigated whether the intrinsic chiroptical properties of HA contributed to the chiral behaviour of the final hybrid material. Accordingly, circular dichroism (CD) spectra of HA, GNRs, and HA–GNR were recorded to assess the effect of biopolymer functionalization on the nanostructures. As shown in Fig. 3(D), the CD spectrum of the GNRs displayed no significant ellipticity across the measured spectral range, consistent with the expected achirality of the colloidal gold nanorods. However, HA exhibited a pronounced negative peak at 220 nm, characteristic of its intrinsic molecular chirality. This signal arises from the helical conformation adopted by its ordered sugar backbone in aqueous solution, which gives rise to a distinct CD signature in the far-UV region.³⁴ Upon surface functionalization with HA, the GNRs exhibited a distinct negative CD peak at 220 nm, along with a weak but discernible positive CD signal at 745 nm (ellipticity +0.9 mdeg), corresponding to the longitudinal localized surface plasmon resonance (LSPR) band. This observation suggests the emergence of induced chiroptical activity, attributed to the asymmetric, stereoselective adsorption of the chiral HA macromolecules onto the anisotropic GNR surface. Such interactions likely generate a chiral electromagnetic field around the nanorods, thereby imparting optical activity to an otherwise achiral system. Based on these results, we hypothesize that the enhanced photothermal efficiency observed in HA–GNR may be an indirect consequence of this induced chirality, potentially influencing the light–matter interactions and energy dissipation mechanisms within the system.^35–37

2.3 Cellular uptake and imaging potential of PEG–PGNC@HA–GNR

The high fluorescence quantum yield of PGNCs, along with the retention of their fluorescence properties in the final hybrid nanomaterial, enabled their use as an intrinsic imaging indicator without the need for additional dyes. The imaging potential of the material was demonstrated in MDA-MB-231 breast cancer cells, with the targeted uptake of PEG–PGNC@HA–GNR due to the presence of HA, capable of binding with CD44, a known HA receptor overexpressed in cancer cells.^38,39 In this study, the cellular uptake and imaging potential of PEG–PGNC@HA–GNR were evaluated at a concentration of 80 µg mL⁻¹ over incubation periods of 2 and 6 hours. Cellular internalization of nanomaterials depends on various factors such as size, surface charge, and surface properties of the material, as well as the biological characteristics of the cell type. The PEG–PGNC@HA–GNR nanohybrid possesses a net negative surface charge, which typically poses a barrier to cellular uptake due to electrostatic repulsion with the negatively charged cell membrane. However, in this case, the presence of HA appears to play a critical role in promoting membrane adhesion and initiating receptor-mediated endocytosis. Fig. 4 illustrates that the cell nuclei were properly stained, exhibiting blue fluorescence, while the internalized PEG–PGNC@HA–GNR displayed varying red fluorescence at different time points. After 2 h of incubation, the uptake was observed by weak, scattered red fluorescence predominantly in the perinuclear cytoplasm, corresponding to early endosomal compartments. After 6 h, there was a substantial increase in red fluorescence intensity, reflecting enhanced endocytic uptake and vesicular trafficking of PEG–PGNC@HA–GNR. This pattern of internalization was also observed after 4 h (Fig. S11, SI). These observations suggest that the increased uptake occurs with longer incubation times, which is attributed to CD44 receptor-mediated endocytosis, specifically through clathrin-/caveolae-independent endocytosis, a known mechanism for the internalization of negatively charged nanoparticles.⁴⁰ It was observed that PEG–PGNC@HA–GNR exhibited red fluorescence signals from the nuclei as well.⁴¹ The underlying mechanism remains unclear and requires further investigation. These findings highlight the importance of surface functionalization with biologically relevant ligands in enhancing the cellular internalization efficiency of nanomaterials, particularly for applications in targeted cancer imaging and therapy.

Fig. 4 Cellular uptake of PEG–PGNC@HA–GNR in MDA-MB-231 cells. Fluorescence microscopy images showing PEG–PGNC@HA–GNR (red) localized within cells, with nuclei stained by Hoechst 33342 (blue). Corresponding brightfield images are included. Uptake was observed after 2 h (upper panel) and 6 h (lower panel) of incubation, observed at 40× magnification.

2.4 In vitro cytocompatibility

The cytocompatibility of GNR, HA–GNR, and PEG–PGNC@HA–GNR was assessed in both triple-negative breast cancer cells (MDA-MB-231) and normal fibroblast cells (L929) using the MTT assay (Fig. 5(A) and Fig. S12, SI).

Fig. 5 Relative cell viability of MDA-MB-231 cells after material incubation for 24 hours (A) without laser irradiation and (B) PEG–PGNC@HA–GNR with and without laser irradiation. (C) Live/dead cell images of MDA-MB-231 incubated with GNRs and PEG–PGNC@HA–GNR without laser and with laser irradiation. The cells were irradiated with an 808 nm laser at a low power density of 0.25 W cm⁻² for 10 minutes using a fiber optic cable.

In this study, the GNRs showed the least cell viability or maximum cytotoxicity. This is attributed to the presence of either surface-bound or free CTAB, as both are known to be highly cytotoxic due to their nonspecific interactions with cells and biomolecules in biological systems.⁴² Modification of GNRs with hyaluronic acid (HA) improved cell viability from 68% to 70% in MDA-MB-231 cells, likely due to the partial replacement of toxic CTAB with biocompatible HA at a concentration of 120 µg mL⁻¹. Surface modification with PEG–PGNC comprising DSPE–PEG-conjugated PGNCs resulted in a further increase in cell viability to 72%. Cytocompatibility was also assessed in L929 fibroblasts treated with GNR, HA–GNR, and PEG–PGNC@HA–GNR, with observed cell viabilities of 69%, 77%, and 80%, respectively (Fig. S12, SI). The reduced viability observed in MDA-MB-231 cells compared to L929 cells may be attributed to the enhanced cellular uptake of the nanomaterials, as MDA-MB-231 cells overexpress CD44 receptors, the primary binding target of HA, which is absent in L929 cells,⁴³ as in the case of MDA-MB-231 cells.

2.5 In vitro photothermal therapeutic effect and cell death analysis

An MTT assay was performed to evaluate the cell death/cytotoxicity of PEG–PGNC@HA–GNR in MDA-MB-231, with and without laser irradiation (Fig. 5(B)). The results showed that the PEG–PGNC@HA–GNR formulation exhibited marked cell death in the presence of a laser due to the PTT effect, demonstrating its superior therapeutic efficacy. A cell viability assay was also performed with GNR, HA–GNR, and PEG–PGNC@HA–GNR in the presence of laser irradiation. Laser exposure led to a reduction in the viability of MDA-MB-231 cells, with GNRs, HA–GNR, and PEG–PGNC@HA–GNR showing cell viabilities of 51%, 47%, and 41%, respectively (Fig. S13, SI). This reduction can be attributed to the photothermal properties of the materials, which generate localized heating upon laser activation, leading to cellular damage and death. These findings support the potential of the materials as effective photothermal therapeutic agents for cancer treatment. The enhanced photothermal effect of PEG–PGNC@HA–GNR is demonstrated by the maximum cell death, highlighting its potential as an efficient PTT agent.

Furthermore, a live/dead assay was performed on MDA-MB-231 cells treated with GNRs and PEG–PGNC@HA–GNR to assess their photothermal therapeutic potential. Following 24 h of incubation, the cells were exposed to 808 nm laser irradiation for 10 minutes. As shown in Fig. 5(C), both treatment groups exhibited a marked reduction in cell viability post-irradiation, confirming the photothermal activity of GNRs in both formulations. Notably, the PEG–PGNC@HA–GNR-treated group demonstrated a higher level of cell death, which correlates with the elevated temperature generated by this formulation upon laser exposure (Fig. 3(A)). This enhanced thermal response is attributed to the contribution of the chiral plasmonic properties of GNRs within the PEG–PGNC@HA–GNR complex. These findings suggest that the inherent fluorescence and photothermal properties of PEG–PGNC@HA–GNR make it a promising candidate for fluorescence imaging-assisted photothermal therapy (PTT) in cancer treatment.

2.6 In vivo imaging: a preliminary study

A preliminary evaluation of the in vivo imaging potential of the developed hybrid material, PEG–PGNC@HA–GNR, was carried out by subcutaneous injection into Swiss Albino mice. Following administration, real-time fluorescence imaging revealed a clearly detectable emission signal from the site of injection, demonstrating the material's capacity for in vivo optical tracking (Fig. S14, SI). This visible fluorescence signal confirms the ability of the hybrid system to function effectively as an optical imaging probe in a biological environment. The observed in vivo signal intensity is primarily attributed to the comparatively high quantum yield of the developed nanohybrid, which ensures efficient photon emission even under physiological conditions. The incorporation of PGNCs within the PEGylated and HA-functionalized structure not only enhances the biocompatibility but also contributes to the sustained bright fluorescence. These findings suggest that the PEG–PGNC@HA–GNR construct holds significant promise for non-invasive imaging applications, particularly in fluorescence-guided diagnostics and image-assisted therapeutic interventions.

3. Conclusions

In this study, we successfully engineered a chiral plasmonic nanohybrid system comprising gold nanoclusters and gold nanorods, carefully designed to retain the intrinsic red fluorescence of GNCs while imparting the photothermal properties of GNRs. Strategic spatial separation within the hybrid architecture effectively minimized Förster resonance energy transfer (FRET), thereby preserving the optical fidelity of each component. Functionalization with hyaluronic acid enabled selective targeting of CD44-overexpressing MDA-MB-231 breast cancer cells, resulting in enhanced in vitro therapeutic efficacy via fluorescence imaging-assisted photothermal therapy. Preliminary in vivo investigations further demonstrated the optical imaging potential of the hybrid system. Overall, this work highlights the feasibility of integrating distinct gold nanostructures into a single multifunctional platform without compromising their individual functionalities. The synergistic enhancement in optical and photothermal properties, combined with tumor-targeting capability and preliminary in vivo validation, positions this hybrid nanomaterial as a promising candidate for future image-guided cancer diagnostics and therapy.

4. Experimental section

4.1 Materials

Hydrogen tetrachloroauratetrihydrate (HAuCl₄), hexadecyltrimethylammonium bromide (CTAB), silver nitrate (AgNO₃), L-ascorbic acid, sodium borohydride, hydrochloric acid, hyaluronic acid (HA), chicken egg white (CEW), sodium hydroxide, 1-ethyl-3-(-3-dimethylaminopropyl)carbodiimide hydrochloride (EDC), N-hydroxysuccinimide (NHS), and DSPE–PEG were used. The water used throughout the study was ultrapure. All glassware was cleaned with aqua regia, rinsed with distilled and ultrapure water, and dried in an oven. All chemicals were commercially available and used without further purification unless specified.

4.2 Synthesis of PEG-coated protein-stabilized gold nanoclusters

The ovalbumin stabilized gold nanoclusters were synthesized as per the described protocol.⁴⁴ Briefly, 0.1 M HAuCl₄ was added to a 10 mL aqueous solution containing CEW under vigorous stirring, which is followed by the addition of NaOH under pH 10 and is incubated at 37 °C under dark conditions for 6 hours. The clusters were lyophilized for further use. The PEG-coated P-GNCs were prepared by mixing both at a 5 : 1 ratio and stirring continuously at room temperature for 10 h.

4.3 Synthesis of HA gold nanorods

The GNRs were prepared using the seed-mediated synthesis method.^45–47 Initially, the seed solution was prepared by treating 0.1 M hexadecyltrimethylammonium bromide (CTAB) solution with 50 mM chloroauric acid (HAuCl₄) followed by 10 mM sodium borohydride (NaBH₄) under vigorous stirring. Next, GNR growth solution was prepared by mixing CTAB, 1 M HCl and 50 mM HAuCl₄ with a pH range of 1.5. This was followed by the addition of 10 mM AgNO₃ and 100 mM ascorbic acid, and to this seed-CTAB is added and the solution is kept undisturbed for 24 h under dark conditions. Excess CTAB was removed from the GNR solution by centrifugation at 10 000 rpm for 30 minutes.

Then, the CTAB-GNR collected were converted to HA–GNR by a ligand exchange process. Briefly, 2 mL of HA solution in water (8 mg mL⁻¹) was added to 60 mL of CTAB-GNR suspension (20 µg Au mL⁻¹) and stirred mildly for 24 h. The solution was centrifuged for 13 000 rpm for 15 minutes and the collected precipitate was ultrasonically dispersed in 15 mL of standard PBS (7.4) and stored at 4 °C for use.

4.4 Synthesis of HA–gold nanorod with PEG–protein gold nanocluster hybrid nanomaterials

The synthesis of PEG–PGNC@HA–GNR was done by maintaining an acidic pH of around 5.5. Initially, the HA functionalized GNR was treated with 5 µL 1-ethyl-3-(-3-dimethylaminopropyl)carbodiimide hydrochloride (EDC) (15 mg mL⁻¹) and 5 µL N-hydroxysuccinimide (NHS) (20 mg mL⁻¹) in MES buffer in order to activate the carboxylate group. This was kept for half an hour with gentle agitation at room temperature followed by the addition of PEG–PGNC according to the concentration. The reaction was carried out under dark conditions for 12 h with gentle agitation. Then, BSA 2% was added to the final concentration in order to prevent further non-selective binding and kept for an hour. The material was purified by centrifugation at 14 000 rpm.⁴⁸

4.5 In vitro cytocompatibility study

Cell viability of the materials was evaluated using an MTT assay in both MDA-MB-231, triple negative breast cancer cell lines and mouse fibroblast cell lines, L929. Briefly, the cells were seeded in a 96 well plate at a density of 4 × 10⁴ cells per well. After 24 h of incubation, the culture media was replaced with material containing media. After 48 h of incubation, the media were removed and 100 µL of fresh DMEM containing 3-(4, 5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT-10 µL, 5 mg mL⁻¹) was added to it. Insoluble MTT formazan crystals were formed after 4 h of incubation. For solubilization of the MTT crystals, 100 µL of DMSO was added to the wells and they were placed on a shaker for 30 min for complete solubilization of the crystals and then the optical density at 570 nm of each well was determined by using a multi-well plate reader (Synergy H1 hybrid multi-mode microplate reader, Bio-Tek, Agilent). The absorbance is directly proportional to the number of living cells in the culture.

In vitro cytotoxicity after laser irradiation was also assessed using the MTT assay. The assay was performed with varying concentrations of the materials, and absorbance was measured at 570 nm using a Synergy H1 hybrid multimode microplate reader (Bio-Tek). The percentage of cell viability at each concentration, relative to the control cells, was calculated and plotted.

4.6 In vitro cellular uptake

The cellular uptake and imaging potential of PEG–PGNC@HA–GNR were evaluated using the MDA-MB-231 cell line, seeded at a density of 4 × 10⁴ cells per well in a four-well plate containing coverslips of 12 mm diameter, and cultured for 24 h. Upon reaching confluency at 24 h, the cells were incubated with the materials at a concentration of 80 µg mL⁻¹ for 2, 4, and 6 h. After incubation, the cells were washed twice with PBS, fixed with 3.7% paraformaldehyde, and counterstained with Hoechst 33342 for 2–5 minutes. The coverslips were then mounted onto glass slides, and the cells were imaged using an IX83 inverted fluorescence microscope (Olympus) equipped with a custom-made filter set of 425/50 and 642/80 nm for excitation and emission, respectively.

4.7 In vitro photothermal therapeutic effect and cell death analysis

Cell death analysis of the material was performed on the MDA-MB-231 cell line using the LIVE/DEAD Cell Imaging Kit (Invitrogen, Thermo Fisher Scientific). The cells were seeded into a 96-well plate at a density of 1 × 10⁴ cells per well and incubated for 24 hours. Upon reaching confluency, the cells were treated with GNRs and PEG–PGNC@HA–GNR at a concentration of 120 µg mL⁻¹ for 24 h. Cells without any treatment were used as the control group. Following material treatment, the cells were irradiated with an 808 nm laser at a power density of 0.25 W cm⁻² for 10 minutes using a fiber optic cable. After irradiation, the cells were stained with Calcein AM/PI and incubated for 30 minutes at 37 °C. The stained cells were then immediately observed under a fluorescence microscope (Olympus IX 83).

4.8 In vivo imaging: preliminary study

The in vivo imaging potential of PEG–PGNC@HA–GNR was evaluated in Swiss albino mice weighing 30–32 g. All experimental procedures were conducted in accordance with the guidelines of the Committee for the Purpose of Control and Supervision of Experiments on Animals (CPCSEA), Government of India, and were approved by the Institutional Animal Ethics Committee of Sree Chitra Tirunal Institute for Medical Sciences and Technology (IAEC approval No-SCT/IAEC-399/MARCH/2021/109). A 2 mg mL⁻¹ suspension of the material was administered subcutaneously at the dorsal neck region of anaesthetized mice, and imaging was performed 20 minutes post-injection using the IVIS Spectrum animal imaging system (Xenogen) at excitation and emission of 430 nm and 780 nm, respectively. The control animal received saline injection at the same site.

Author contributions

Nisha N.: writing – original draft, methodology, investigation, and conceptualization. Dhujana N. S.: writing – original draft, methodology, investigation, and conceptualization. Remyachand R.: writing – original draft, methodology, investigation, and conceptualization. M. E. Dhushyandhun: methodology, and investigation. Ramapurath S. Jayasree: conceptualization, funding acquisition, resources, writing – review and editing, and supervision.

Conflicts of interest

There are no conflicts to declare.

Data availability

Data of the manuscript are available upon request from the authors.

The data supporting this article have been included as part of the supplementary information (SI). SI includes the results of absorption and emission spectra of PGNC, DLS results, quantum yield calculations, TEM images of PGNC, PEG-PGNC and PEG-PGNC@HA-GNR, Photothermal profile of PEG-PGNC@HA-GNR, Photothermal conversion efficiency, in vitro cytotoxicity assay and in vivo images. See DOI: https://doi.org/10.1039/d5tb01497b.

Acknowledgements

The authors R. S. J. and N. N. acknowledge the Indian Council of Medical Research (ICMR) Government of India, for the financial support of the project IIRP-2023-0782/F1. The author D. N. S. acknowledges the Department of Biotechnology, Government of India, for the DBT-JRF fellowship (DBT/2022-23/SCTMIST/2102). M. E. D. acknowledges the financial support provided by the Science and Equity, Empowerment and Development (SEED) Division, Department of Science and Technology, Government of India, through the project SEED/SCSP/2019/117, and its PI of SCTIMST, Dr Roy Joseph.

References

1. Z. Li, Q. Sun, Y. Zhu, B. Tan, Z. P. Xu and S. X. Dou, J. Mater. Chem. B, 2014, 2, 2793–2818.
2. B. A. Kairdolf, A. M. Smith, T. H. Stokes, M. D. Wang, A. N. Young and S. Nie, Annu. Rev. Anal. Chem., 2013, 6, 143–162.
3. L. Han, J. Zhao, X. Zhang, W. Cao, X. Hu, G. Zou, X. Duan and X. J. Liang, ACS Nano, 2012, 6, 7340–7351.
4. J. Xie, Y. Zheng and J. Y. Ying, J. Am. Chem. Soc., 2009, 131, 888–889.
5. W. Fei, S. Y. Tang and M. B. Li, Nanoscale, 2024, 16, 19589–19605.
6. S. Santhoshkumar, M. Madhu, W. Bin Tseng and W. L. Tseng, 2023.
7. A. Nag, P. Chakraborty, A. Thacharon, G. Paramasivam, B. Mondal, M. Bodiuzzaman and T. Pradeep, J. Phys. Chem. C, 2020, 124, 22298–22303.
8. A. Baghdasaryan and H. Dai, Chem. Rev., 2025, 125, 5195–5227.
9. Nonappa, Beilstein-Institut Zur Forderung der Chemischen Wissenschaften, 2020.
10. C. Zhang, X. Gao, W. Chen, M. He, Y. Yu, G. Gao and T. Sun, iScience, 2022, 25(10), 105022.
11. S. Iravani and R. S. Varma, Green Chem., 2020, 22, 612.
12. B. A. Russell, B. Jachimska, I. Kralka, P. A. Mulheran and Y. Chen, J. Mater. Chem. B, 2016, 4, 6876–6882.
13. D. Hao, X. Zhang, R. Su, Y. Wang and W. Qi, J. Mater. Chem. B, 2023, 11, 5051–5070.
14. Y. Yang, Y. Zhang, A. Thakur, R. Li, H. Xu, Z. Wang, M. Ghavami, Z. Tu and H. Liu, Int. J. Pharm., 2019, 571, 118704.
15. X. J. Li, J. Ling, C. L. Han, L. Q. Chen, Q. E. Cao and Z. T. Ding, Anal. Sci., 2017, 33, 671–675.
16. R. Rajamanikandan and M. Ilanchelian, ACS Omega, 2018, 3, 14111–14118.
17. Y. Guo, H. T. N. N. Amunyela, Y. Cheng, Y. Xie, H. Yu, W. Yao, H. W. Li and H. Qian, Food Chem., 2021, 335, 127657.
18. Y. Wang, Y. Wang, F. Zhou, P. Kim and Y. Xia, Small, 2012, 8, 3769–3773.
19. K. Selvaprakash and Y. C. Chen, Biosens. Bioelectron., 2014, 61, 88–94.
20. B. Khlebtsov, V. Zharov, A. Melnikov, V. Tuchin and N. Khlebtsov, Nanotechnology, 2006, 17, 5167–5179.
21. A. Koulali, P. Radomski, P. Ziółkowski, F. Petronella, L. De Sio and D. Mikielewicz, Sci. Rep., 2025, 15, 9543.
22. S. Liao, W. Yue, S. Cai, Q. Tang, W. Lu, L. Huang, T. Qi and J. Liao, Front. Pharmacol., 2021, 12, 664123.
23. B. Ni, G. González-Rubio, K. Van Gordon, S. Bals, N. A. Kotov and L. M. Liz-Marzán, Adv. Mater., 2024, 36(47), e2412473.
24. S. Wang, X. Liu, S. Mourdikoudis, J. Chen, W. Fu, Z. Sofer, Y. Zhang, S. Zhang and G. Zheng, ACS Nano, 2022, 16, 19789–19809.
25. K. T. Dicker, L. A. Gurski, S. Pradhan-Bhatt, R. L. Witt, M. C. Farach-Carson and X. Jia, Acta Biomater., 2014, 10(4), 1558–1570.
26. M. Yusupov, A. Privat-Maldonado, R. M. Cordeiro, H. Verswyvel, P. Shaw, J. Razzokov, E. Smits and A. Bogaerts, Redox Biol., 2021, 43, 101968.
27. P. Spaeth, S. Adhikari, W. Heyvaert, X. Zhuo, I. García, L. M. Liz-Marzán, S. Bals, M. Orrit and W. Albrecht, ACS Photonics, 2022, 9, 3995–4004.
28. K. Park, L. F. Drummy, R. C. Wadams, H. Koerner, D. Nepal, L. Fabris and R. A. Vaia, Chem. Mater., 2013, 25, 555–563.
29. A. Chakraborty, H. Dave, B. Mondal, Nonappa, E. Khatun and T. Pradeep, J. Phys. Chem. B, 2022, 126, 1842–1851.
30. C. Brownlee, ACS Nano, 2023, 17, 16287–16290.
31. N. Savage, Nature, 2013, 495, S8–S9.
32. F. Xiao, Y. Chen, J. Qi, Q. Yao, J. Xie and X. Jiang, Adv. Mater., 2023, 35, 2210412.
33. J. S. Mohanty, A. Maity, T. Ahuja, K. Chaudhari, P. Srikrishnarka, V. Polshettiwar and T. Pradeep, Nanoscale, 2021, 13, 9788–9797.
34. A. J. Miles and B. A. Wallace, Chem. Soc. Rev., 2016, 45, 4859–4872.
35. W. Cai, Y. Shi, N. Liu, Z. Z. Yin, J. Li, L. Xu, D. Wu and Y. Kong, Spectrochim. Acta, Part A, 2024, 318, 1386–1425.
36. J. Lu, Y. X. Chang, N. N. Zhang, Y. Wei, A. J. Li, J. Tai, Y. Xue, Z. Y. Wang, Y. Yang, L. Zhao, Z. Y. Lu and K. Liu, ACS Nano, 2017, 11, 3463–3475.
37. T. Mantso, S. Vasileiadis, I. Anestopoulos, G. P. Voulgaridou, E. Lampri, S. Botaitis, E. N. Kontomanolis, C. Simopoulos, G. Goussetis, R. Franco, K. Chlichlia, A. Pappa and M. I. Panayiotidis, Sci. Rep., 2018, 8(1), 10724.
38. K. Kim, H. Choi, E. S. Choi, M. H. Park and J. H. Ryu, Pharmaceutics, 2019, 11, 301.
39. X. Yan, Q. Chen, J. An, D. E. Liu, Y. Huang, R. Yang, W. Li, L. Chen and H. Gao, J. Mater. Chem. B, 2018, 7, 95–102.
40. J. Dausend, A. Musyanovych, M. Dass, P. Walther, H. Schrezenmeier, K. Landfester and V. Mailänder, Macromol. Biosci., 2008, 8, 1135–1143.
41. D. H. Lin and A. Hoelz, Annu. Rev. Biochem., 2019, 88, 725–783.
42. E. E. Connor, J. Mwamuka, A. Gole, C. J. Murphy and M. D. Wyatt, Small, 2005, 1, 325–327.
43. G. E. S. Chaudhry, A. Akim, M. N. Zafar, N. Safdar, Y. Y. Sung and T. S. T. Muhammad, Adv. Pharm. Bull., 2021, 11, 426–438.
44. D. Joseph and K. E. Geckeler, Colloids Surf., B, 2014, 115, 46–50.
45. Z. S. Mbalaha, P. R. Edwards, D. J. S. Birch and Y. Chen, ACS Omega, 2019, 4, 13740–13746.
46. A. Gole and C. J. Murphy, Chem. Mater., 2004, 16, 3633–3640.
47. M. Z. Wei, T. S. Deng, Q. Zhang, Z. Cheng and S. Li, ACS Omega, 2021, 6, 9188–9195.
48. R. V. Nair, M. F. Puthiyaparambath, R. Chatanathodi, L. V. Nair and R. S. Jayasree, Nanoscale, 2022, 14, 13561–13569.

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Footnote

† These authors contributed equally.

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