Suppressing Cell Migration through Discoidal Bottlebrush Copolymer Nanocarriers

Abstract

Precise control over two-dimensional (2D) polymer nanostructures remains a fundamental challenge, as polymer self-assembly overwhelmingly favors spherical morphologies. Here, we introduce a topology-driven design strategy that overcomes this limitation, enabling the predictable and modular formation of amorphous polymer nanodiscs. Our strategy decouples nanodisc diameter from bottlebrush chemistry. By systematically varying the length of hydrophobic poly­(ethoxyethyl glycidyl ether) (PEE) side chains in the bottlebrush segment, we obtain precise control over nanodisc diameter while maintaining uniform thickness. This tunability allows investigation of size-dependent cellular interactions using MDA-MB-231 cancer cells. Importantly, we show that nanodiscs can serve as pH-responsive carriers that disassemble under acidic conditions and release ICAM-1 inhibitors (A-205804), resulting in effective suppression of cancer cell migration.

Introduction

Two-dimensional (2D) nanomaterials have transformed electronics, catalysis, and energy technologies, − yet their translation into biological environments remains limited. − Conventional inorganic 2D materials, although structurally precise, are typically rigid, persistent, and poorly suited to soft, dynamic biological systems. This necessitates the development of a soft-matter analogue: a biocompatible, tunable, and structurally anisotropic 2D nanomaterial capable of programmed interactions with cells and tissues.

Polymeric 2D nanodiscs might offer an attractive soft-matter solution for drug delivery, promising enhanced therapeutic outcomes driven by their unique shape-mediated cellular interactions, predictable hydrodynamic behavior, and capacity for controlled therapeutic release. , Yet, their development has been hindered by a fundamental challenge of polymer self-assembly: to reduce interfacial free energy, amphiphilic systems overwhelmingly form low-energy spherical aggregates. As a result, the ∼50 polymer nanodisc systems reported over the past three decades have largely been serendipitous, dependent on restrictive solvent conditions, crystallinity, top-down fabrication, or specialized chemistries that limit scalability and biological adaptation. We recently summarized the international progress on polymer nanodisc formation and identified the lack of a reliable platform that enables 2D assembly of amphiphilic copolymers and consequently their assessment for nanomedicine applications.

In 1998, Stupp highlighted the potential of “rodcoil molecules” in self-assembly. Molecules that feature a rigid (rod) and a flexible (coil) segment were predicted to assemble into disc-like structures, among others. Reducing the flexibility of a macromolecule to increase its rigidity is typically done through chemistry (e.g., via conjugation). An alternative strategy is the use of topology whereby the introduction of branching within a polymer reduces its flexibility. A prominent example is a molecular polymer bottlebrush (MPB) (or bottlebrush polymer), − where side chains are densely grafted on a polymer backbone, leading to stiffening and a stretched conformation. Over the past decade, bottlebrush-type polymers have seen a striking increase in application-focused research, especially on nanomedicines , and polymer networks, , in additive manufacturing , and photonics. ,

Recently, we reimagined Stupp’s “rodcoil” concept and designed amphiphilic bottlebrush block copolymers (BBCs) with a flexible linear (coil) and a semiflexible bottlebrush (rod) segment. Leveraging the advantages of polymer topologyrather than chemical composition alonethis architecture enabled the reliable fabrication of soft, amorphous nanomaterials directly in aqueous media. Because the rigidity of the bottlebrush segment is encoded in the architecture, the platform is modular and adaptable, which enabled the assembly of nanodiscs made from inert and stimuli-responsive polymers. − While nanodiscs can be reliably generated through this approach, controlling their dimensions independently of chemical function has not been possible.

Herein, we introduce a strategy that decouples nanodisc diameter from bottlebrush chemistry. By systematically varying the length of hydrophobic poly­(ethoxyethyl glycidyl ether) (PEE) side chains in the bottlebrush segment, we obtain precise control over nanodisc diameter while maintaining uniform thickness (Scheme A). This result was unexpected because in an earlier BBCs example (albeit using a grafting-from approach), altering the side chain length allowed nanodisc formation only in a very narrow window or not at all. ,, Our new approach overcomes this limitation by successfully decoupling the nanodisc diameter from the chemical functionality, enabling robust formation of pH-responsive nanodiscs across a tunable size range. This capability allows now a direct investigation of diameter-dependent cellular interactions within MDA-MB-231 cells, a widely used triple-negative breast cancer (TNBC) model. Given the critical role of the intercellular adhesion molecule-1 (ICAM-1), a key regulator of cancer cell adhesion, motility, and metastasis frequently overexpressed in aggressive cancers such as TNBC, − we incorporated its selective inhibitor (A-205804 − ) into the nanodiscs to showcase their therapeutic potential (Scheme B). Upon release, A-205804 led to targeted inhibition of cancer cell migration in vitro, demonstrating how precision molecular design and stimuli-responsive 2D nanomaterials can be combined to enable next-generation therapeutics.

1. (A) Design and Self-Assembly of Amphiphilic BBCs into Functional Nanodiscs. (B) Co-Assembly of BBCs with A-205804 to form pH-Responsive Nanodiscs that Degrade at Acidic pH and Release the Payload for Therapeutic Effects.

Results and Discussion

Synthesis and Characterization of Cy5-Labeled PEG₁₁₄-block-[(PGMA-N₃)₂₀-graft-(PEE _n )₁₅] (PE_nC)

The pH-responsive, rod–coil-like BBCs were synthesized through a grafting-to strategy. Briefly, mPEG₁₁₄-CDTPA macro-RAFT agent was chain-extended with glycidyl methacrylate (GMA) via photoinduced electron/energy transfer reversible addition-fragmentation chain-transfer (PET-RAFT) polymerization, followed by azidation to afford the clickable PEG₁₁₄-block-(PGMA-N₃)₂₀ backbone. Three alkyne-terminated PEE side chains with degrees of polymerization (DP = 14, 28, and 42) were synthesized by anionic ring-opening polymerization, followed by end-group functionalization (Scheme S1). These alkyne-functionalized PEE side chains were subsequently cografted with a small fraction of Cy5 fluorophores onto the azide-containing polymer backbone via Cu­(I)-catalyzed azide-alkyne cycloaddition (CuAAC) to afford the final Cy5-labeled amphiphilic BBCs (Scheme S2).

The resulting BBCs were characterized by proton nuclear magnetic resonance (¹H NMR) spectroscopy, size exclusion chromatography (SEC) and Fourier-transform infrared spectroscopy (FTIR) to confirm their chemical structures (Figures S1–S18). Across all three compositions, the side chain grafting efficiencies were approximately 75%, regardless of PEE side chain length. Such grafting densities are consistent with reported grafting-to efficiencies for densely packed bottlebrushes to ensure stretched conformation and decreased flexibility. Structurally, all BBCs feature a common molecular architecture, with a flexible linear PEG block serving as the hydrophilic segment, and a rigid PEE bottlebrush acting as the hydrophobic segment. The reduced flexibility of the bottlebrush segment arises from the densely grafted and sterically crowded side chains. Noteworthy, while the backbone and grafting density were fixed across all variants, systematic variation of the side chain DP has allowed us to tune the overall hydrophobic volume of the brush segment without altering its chemical identity. Thus, this design enabled us to investigate the effect of bottlebrush volume on the self-assembly process, while the PEE block introduces the opportunity for a pH-responsive hydrophobic-to-hydrophilic transition, which is favorable in drug release applications. ,

Self-Assembly of PE_nC Nanodiscs

To investigate the influence of side chain length on self-assembly, the BBCs were dissolved in N,N-dimethylformamide (DMF) (good solvent for BBCs) and dialyzed into water (selective solvent for the bottlebrush). Interestingly, all three BBC variants (PE₁₄C, PE₂₈C and PE₄₂C) spontaneously self-assembled to well-defined nanodiscs in aqueous solution, as confirmed by dynamic light scattering (DLS), transmission electron microscopy (TEM), and atomic force microscopy (AFM) (Figures and S19). These findings were surprising given that previous attempts to vary side chain length (albeit using grafting-from approaches) failed to yield nanodiscs reliably. , The likely reason seemed to be the reduced grafting densities when grafting methacrylic monomers via ATRP which made it difficult to find a balance between bottlebrush flexibility and amphiphilic character of the BBC. The current results highlight the robustness of our grafting-to approach-based polymer design using polyether-based side chains, wherein 2D self-assembly is possible even across a range of side chain lengths. A systematic correlation between side chain length and nanodisc diameter was observed. Specifically, the hydrodynamic diameter (D _h) of the nanodiscs, as measured by DLS, increased progressively with their side chain length, yielding ∼140 nm for PE₁₄C, ∼230 nm for PE₂₈C, and ∼450 nm for PE₄₂C (Table ). Note, DLS assumes an equivalent sphere and does not truly measure our planar nanodisc dimensions. However, the size evolution can be attributed to enhanced hydrophobic interactions among longer PEE side chains, which promote stronger interchain associations and more extensive hydrophobic domain formation. Despite their differences in diameter, all nanodiscs exhibited a uniform thickness of ∼8 nm, as observed by AFM (Figure C,G,K). This consistent thickness suggests that the stacking or layering of the bottlebrush segments remains constant, while disc diameter is modulated laterally through the side chain DP. The ability to adjust nanodisc diameter in a predictable and controlled manner through side chain engineering thus represents a straightforward strategy to investigate their structure–property relationships, particularly in the context of size-dependent cellular uptake and biological interactions of 2D soft nanomaterials. Notably, when we used a grafting-from approach to design rod–coil building blocks, achieving nanodiscs was less straightforward, as variations in bottlebrush side chain length drastically altered the bottlebrush and amphiphilic character and led to morphological transitions.

1.

(A, E, I) Intensity-weighted hydrodynamic diameter (D _h ) of self-assembled nanodiscs in water as measured by DLS (concentration: 1 mg mL⁻¹). (B, F, J) TEM micrographs and (C, G, K) AFM height images of self-assembled nanodiscs, including (D, H, L) their cross-sectional analysis across selected nanodiscs. AFM z-scale: −5 to 10 nm.

1. Characterization Data on BBCs and Their Corresponding Nanodiscs.

	bottlebrush side chains			BBCs				Nanodiscs
	Composition (NMR)	Grafting Efficiency (%)		M n,NMR (kg/mol)	M n,GPC (kg/mol)	Đ b		D h (nm)
PE14C	PEE14	75		32.5	37.9	1.15		141 ± 2
PE28C	PEE28	75		53.0	46.0	1.17		229 ± 4
PE42C	PEE42	75		73.4	51.6	1.17		446 ± 5

^aDetermined by NMR spectroscopy using end group analysis.

^bDetermined by DMAc SEC with PMMA calibrations.

^cDetermined by DLS (intensity-weighted) with the results expressed as means ± SD, n = 3 independent replicates.

Cellular Uptake and Cytotoxicity of PE_nC Nanodiscs

To assess the cytocompatibility of nanodiscs, MTT assays were performed in MDA-MB-231 cells across a range of concentrations after 24 and 48 h incubation. All three variants of nanodiscs demonstrated excellent biocompatibility and maintained cell viabilities exceeding 80% (Figures A and S20). The cellular uptake of nanodiscs was further evaluated by confocal laser scanning microscopy (CLSM) imaging. After incubating for 24 h, strong intracellular fluorescence signals were observed, indicating internalization of Cy5-labeled nanodiscs (Figures B and S21). Cell association was quantified using flow cytometry, which revealed a size-dependent trend: PE₂₈C nanodiscs displayed the highest association, followed by PE₄₂C and then PE₁₄C (Figure C). These results were consistent with the CLSM observations, where PE₂₈C exhibited the strongest and PE₁₄C the weakest intracellular fluorescence, respectively. To examine whether this size-dependent uptake behavior is specific to the MDA-MB-231 cell line, nanodisc internalization was also evaluated in HeLa cells. Notably, a similar trend was observed, with PE₂₈C again exhibiting the highest level of cellular association among our tested nanodiscs (Figure S22). The observed dependence of cellular uptake on nanodisc dimensions is consistent with general principles of nanoparticle–cell interactions, where particle size has been shown to critically influence uptake efficiency and endocytic pathways in various cell types. Mechanistically, this size dependency can be understood as an energetic balance between the interfacial adhesion energy required for membrane wrapping and the elastic cost associated with membrane deformation. , Smaller nanodiscs such as PE₁₄C may lack sufficient surface area to generate the cumulative adhesive interactions necessary to drive spontaneous membrane wrapping. Conversely, for larger nanodiscs such as PE₄₂C, the high energetic cost of deforming the cell membrane over a larger lateral area may limit the internalization rate. − Consequently, nanodiscs of intermediate size, such as PE₂₈C (∼230 nm in diameter), appear to provide a favorable balance for cellular internalization. Although further mechanistic studies on uptake pathways are ongoing, these findings underscore the importance of nanodisc dimension as a critical design parameter influencing biological interactions. Based on these results, PE₂₈C was selected as the optimized system for subsequent studies on nanodisc disassembly and cargo release, as it combined cytocompatibility with favorable cellular uptake in the MDA-MB-231 model.

2.

(A) Cell viability of MDA-MB-231 cells following treatment with PE_nC nanodiscs at various concentrations for 24 h at 37 °C. Data are presented as mean ± SD, n = 3. (B) CLSM images of MDA-MB-231 cells after 24 h incubation with PE_nC nanodiscs at 37 °C. Scale bars are 20 μm. Color code: nucleus (blue), nanodiscs (red). (C) Flow cytometry analysis of MDA-MB-231 cells after 24 h incubated with PE_nC nanodiscs at 37 °C.

pH-Responsive Behavior and Controlled Cargo Release

Nanocarriers designed for intracellular delivery should ideally be able to release their therapeutic cargo in response to biological cues. , Among these, acidic pH is particularly relevant, since nanoparticles typically encounter acidic environments in endosomes and lysosomes (pH 5.0–5.5) as well as mildly acidic tumor microenvironments. , Thus, designing carriers that remain intact at physiological pH but degrade under acidic conditions provides a strategy for site-specific drug release while minimizing premature drug leakage. In our case, we selected PE₂₈C to evaluate its stability and degradation behavior under pH 7.4 and pH 5.0, representing physiological circulation and late endosomal/lysosomal conditions, respectively. At neutral pH conditions, PE₂₈C nanodiscs remained structurally stable. DLS measurements showed no significant change in hydrodynamic size, and TEM imaging after 30 days revealed well-defined, intact disc-like morphologies, indicating negligible structural degradation under neutral conditions (Figure S23). The hydrophobic bottlebrush nanodisc core is composed of PEE, which undergo acid-catalyzed hydrolysis of the pendent acetal groups into water-soluble polyglycidol. This chemical conversion turns the hydrophobic core-forming segments into hydrophilic counterparts, resulting in the gradual disassembly of the nanodiscs (Figure ). This mechanism is consistent with established reports on the acid-labile nature of acetal-functionalized poly­(glycidyl ether)­s. , DLS, TEM, and cryogenic electron microscopy (cryo-EM) were used to follow structural changes under acidic conditions. DLS analysis initially showed a dominant population at ∼230 nm, suggesting these nanodiscs have remained largely intact in their sizes during the first few days of acidic incubation (Figure A,B). However, TEM imaging at day 1 already revealed irregular disc edges and partial distortion of the circular morphology, indicating that degradation begins at the periphery before any morphology changes are detected in their size distribution (Figure C). By day 5, DLS measurements began to reflect this structural degradation, with a broader distribution and the emergence of smaller particles (Figure D). At later stages (day 10 to day 15), the DLS traces have completely shifted toward smaller hydrodynamic sizes (Figure E,F). TEM imaging revealed that by day 10, the nanodiscs maintained their general framework but exhibited a loosened, porous morphology due to internal hydrolysis (Figure E). This internal erosion creates a finer textured appearance before the structure reaches a critical point of collapse. By day 15, continued degradation leads to the physical fragmentation of this weakened framework into discrete, irregular fragments (Figure F). Complementary cryo-EM imaging further confirmed these results (Figures S24 and S25). On day 0, nanodiscs appeared as well-defined, shape-anisotropic discs, whereas on day 15 only fragmented structures can be observed. Together, these results suggest that acid-triggered degradation proceeds via edge erosion, ultimately leading to disassembly into hydrophilic copolymers. While the nanodisc disassembly currently occurs over several days, the acetal-based chemistry will allow for adjusting of disassembly kinetics; albeit a gradual disassembly may be most advantageous for sustained drug release and prolonging structural integrity for drug delivery applications.

3.

PE₂₈C nanodiscs degradation studies at pH 5.0. DLS measurements of (A) intensity- and (B) number-weighted size distributions. TEM micrographs (negatively stained) of PE₂₈C nanodiscs at different degradation stages (C) day 1, (D) day 5, (E) day 10, (F) day 15.

Building on this pH-responsiveness, we next evaluated PE₂₈C nanodiscs as drug carriers using A-205804, a selective small-molecule inhibitor of ICAM-1, with no change in hydrodynamic volume (Figure S26). This inhibitor suppresses cancer cell migration by blocking ICAM-1-mediated adhesion and signaling. , PE₂₈C nanodiscs achieved a drug loading content of 3.6 wt % with an encapsulation efficiency of 15% (Figure S27). ICAM-1 is typically administered as an oral drug which might explain the lower drug loading efficiency compared to previous results using anticancer drugs like doxorubicin. Subsequent A-205804 release studies showed minimal drug leakage under neutral conditions (7.1 ± 0.1 wt %), confirming the structural stability of the nanodiscs and effective encapsulation of A-205804 within their hydrophobic core. In contrast, under acidic conditions, release was markedly accelerated, reaching 48.1 ± 1.8 wt % within 9 days (Figure S28). Note that carrier degradation and drug release are related but distinct processes: while degradation aids release, the kinetics of drug release do not necessarily mirror those of the nanodisc disassembly.

Inhibition of Cancer Cell Migration by A-205804-Loaded PE₂₈C Nanodiscs

ICAM-1 has been reported to be a critical regulator for cancer cell adhesion and migration, and its overexpression is linked to enhanced metastatic potential in aggressive tumors. , A-205804 is a selective small-molecule inhibitor that suppresses ICAM-1 transcription, thereby reducing cell migration and invasion. To assess whether PE₂₈C nanodiscs could serve as effective pH-responsive delivery systems for this inhibitor, A-205804-loaded PE₂₈C nanodiscs (A-205804@PE₂₈C) were tested using wound healing and Transwell migration assays. In the wound healing assay, untreated MDA-MB-231 cells displayed efficient closure of the scratched area (53.1 ± 1.0%) within 48 h. By contrast, A-205804@PE₂₈C nanodiscs produced the most pronounced inhibition, limiting wound closure to 6.5 ± 1.5%, compared with 16.6 ± 1.5% for free A-205804 and 25.4 ± 1.5% for empty PE₂₈C nanodiscs (Figure A,B).

4.

Wound healing assays with (A) representative microscopy images and (B) quantitative analysis of wound closure at 0 and 48 h (n = 3). Transwell migration assays with (C) representative microscopy images and (D) quantitative analysis of cells migrating into the lower chamber (n = 3). All scale bars are 200 μm.

A similar trend was observed in the Transwell migration assay, where A-205804@PE₂₈C nanodiscs yielded the most substantial suppression of migration, with only 49.3 ± 8.0 cells crossing the membrane, corresponding to more than 85% inhibition relative to the blank (361.7 ± 13.1). Both free A-205804 (129.7 ± 4.0) and empty PE₂₈C nanodiscs (215.3 ± 17.2) exhibited partial suppression, yet the combination of A-205804@PE₂₈C nanodiscs considerably outperformed either component alone (Figure C,D). Interestingly, empty nanodiscs retained moderate activity, which may arise from the influence of PEG chain density and length on evading the protein adsorption and cell–substrate interactions. Such processes are closely linked to adhesion and motility. , Moreover, the superior efficacy of A-205804@PE₂₈C relative to the free drug may be attributed to effects established mechanisms of polymeric nanocarriers, which can act as pericellular depots to enhance local drug concentrations, protect against premature dilution, and prolong exposure of cells to the inhibitor. , Consistent with these observations, fluorescence colocalization studies show that PE₂₈C nanodiscs localize to lysosomal compartments following cellular internalization (Figure S29 and Table S1). Combined with the accelerated release of A-205804 under acidic conditions that mimic the lysosomal environment (Figure S28), these results support a mechanism in which lysosomal delivery and pH-responsive degradation of PE₂₈C facilitate intracellular drug liberation. Collectively, these results demonstrate that PE₂₈C nanodiscs function as stimuli-responsive delivery systems and may offer synergistic effects with their cargo molecules, enabling more potent inhibition of cancer cell migration.

Conclusions

We presented a topology-assisted self-assembly strategy utilizing BBCs to engineer stimuli-responsive nanodisc in aqueous media. Adjusting the side chain length within the bottlebrush segment of the BBCs, enabled control over nanodisc amphiphilicity leading to controllable nanodisc diameters which in turn influenced cellular association. The nanodisc cores could hydrolyze and cause a gradual disassembly of nanodiscs under acidic conditions. This enabled the controlled release of encapsulated ICAM-1 inhibitors (A-205804), suppressing breast cancer cell migration in vitro. Altogether, our findings established side chain engineering of BBCs as a versatile strategy to program both their self-assembled nanostructures and biological functions. With pH-triggered degradation in acidic environments, these nanodiscs show potential as a next-generation drug delivery system for intracellular delivery and therapeutic modulation of cancer cell behaviors.

Experimental Section

Materials

Poly­(ethylene glycol) methyl ether 5000 (mPEG 5K), 4-cyano-4-[(dodecylsulfanylthiocarbonyl)­sulfanyl] pentanoic acid (CDTPA, 97%), anhydrous dichloromethane (DCM, ≥99.8%), N-ethyl-N′-(3-(dimethylamino)­propyl)­carbodiimide hydrochloride (EDC·HCl, 98%), glycidyl methacrylate (GMA, 97%), N,N,N′,N″,N″-pentamethyldiethylenetriamine (PMDETA, 99%), copper­(I) bromide (CuBr, 98%), glycidol (98%), 3,4-dihydro-2H-pyran (97%), benzyl alcohol, anhydrous (≥99.8%), p-toluenesulfonic acid monohydrate (p-TsOH, ≥98.5%), phosphazene base P₄-t-Bu Solution (t-BuP₄, 0.8 M in hexane), benzoic acid (≥99.5%), propargyl bromide solution (80 wt % in toluene) were sourced from Sigma-Aldrich. Ethyl vinyl ether (>98.0%) was purchased from Tokyo Chemical Industry. Tetra-n-butylammonium bromide (TBAB, > 98%) was obtained from Thermofisher Scientific. N,N-dimethylpyridin-4-amine (DMAP, ≥99%), ammonium chloride (NH₄Cl, ≥99.5%), diethyl ether were sourced from Merck. Magnesium sulfate (MgSO₄), sodium azide (NaN₃, pure solid) were sourced from Ajax. Dimethylformamide (DMF, 99.8% by gas chromatography (GC)) was sourced from RCI LabScan. Zinc tetraphenylporphyrin (ZnTPP) was synthesized according to a previously reported procedure. Note that toluene was stored in a solvent purification system from VAC (USA) and further dried with 3Å molecular sieve for 24 h. Dulbecco’s Modified Eagle’s Medium (DMEM), fetal bovine serum (FBS), NucBlue Live ReadyProbes Reagent (Hoechst 33342) and LysoTracker Red were purchased from and Thermofisher Scientific. Cy5 alkyne (95%), thiazolyl blue tetrazolium bromide (MTT, 98%) and Dulbecco’s phosphate buffered saline (DPBS) were purchased from Sigma-Aldrich. Antibiotic-antimycotic (100×) and trypsin-EDTA (0.25%, phenol red) were purchased from Life Technologies. Crystal Violet Hydrate was purchased from Tokyo Chemical Industry. Costar Transwell cell culture inserts are purchased from Corning. Culture-Insert 2 Well in μ-Dish 35 mm are purchased from ibidi. MDA-MB-231 cells were sourced from Prof. Elizabeth New’s research group at the School of Chemistry, University of Sydney.

Characterization

Nuclear Magnetic Resonance (NMR) Spectroscopy

NMR spectra of polyether copolymers were acquired at Yonsei University using a Bruker AVANCE III HD 400 MHz spectrometer. Additional NMR analyses, including spectra of the polymer backbone, alkyne-functionalized side-chain polyether copolymers, and their corresponding grafted bottlebrush block copolymers, were conducted at the University of Sydney on a Bruker NEO 300 MHz spectrometer. ¹H NMR experiments were performed using the standard single-pulse (zg) sequence with a 90° pulse and a recycle delay (D1) of 2–5 s. Chemical shifts were referenced to residual solvent peaks: CDCl₃ at 7.26 ppm and MeOD at 3.31 ppm, as appropriate. Deuterated solvents were purchased from Sigma-Aldrich and Cambridge Isotope Laboratories and used as received without further purification.

Size Exclusion Chromatography (SEC)

SEC measurements were carried out on a Shimadzu Prominence UFLC (ultrafast liquid chromatography) system equipped with a Shim-pack GPC-800DP guard column and two Phenogel analytical columns (5 μm particle size, 10⁴ Å and 10⁵ Å pore sizes) connected in series. The mobile phase was HPLC-grade dimethylacetamide (DMAc) containing 0.03 wt % lithium bromide (LiBr) and 0.05 wt % butylated hydroxytoluene (BHT) to minimize polymer aggregation and degradation. The eluent was delivered at a constant flow rate of 1.0 mL min⁻¹, and the column set was maintained at 50 °C. Retention times were calibrated using PMMA narrow standards from PSS.

Fourier-Transform Infrared Spectroscopy (FTIR)

FTIR spectra were collected using a Bruker α II Platinum ATR spectrometer equipped with a monolithic diamond crystal. Each spectrum was acquired with 16 scans at a resolution of 4 cm^–1, with automatic atmospheric background subtraction.

Dynamic Light Scattering (DLS)

DLS measurements were carried out using a Malvern Zetasizer Ultra instrument equipped with a 633 nm He–Ne laser. The hydrodynamic diameters of polymer self-assemblies were measured directly without further sample treatment.

Atomic Force microscopy (AFM)

AFM imaging was performed in air using a Bruker Multimode 8 system equipped with a NanoScope V controller, operating in standard tapping mode with Tap300Al-G cantilevers (Budget Sensors; nominal resonance frequency ∼300 kHz, spring constant ∼30 N/m). Samples were prepared by depositing 20 μL of an aqueous dispersion of polymer self-assembled aggregates onto a clean silicon wafer. After air drying for 10 min, any remaining solution was removed by gently touching the edge of the droplet with filter paper, followed by drying under a gentle stream of nitrogen. Image analysis and processing were carried out using Bruker NanoScope software.

Transmission Electron Microscopy (TEM)

TEM imaging was conducted using a JEM-fi2100CR transmission electron microscope (JEOL) equipped with a 5k × 4k CMOS camera (EMSIS). Images were acquired in bright-field mode with a spot size of 1, and diffraction contrast was enhanced using a 20 μm objective aperture. The accelerating voltage was set to 200 kV. Samples were prepared by depositing 4 μL of the polymer self-assembly solution onto a carbon-coated copper grid, followed by air drying for 10 min. Residual solution was removed by gently touching the edge of the grid with filter paper. Negative staining was performed using a 2 wt % uranyl acetate (UA) solution. Briefly, a 10 μL droplet of UA solution was placed on a piece of parafilm, and the dried grid was placed face-down onto the droplet for 30 s. After staining, excess UA was removed with filter paper, and the grid was allowed to air-dry before imaging.

Cryogenic Electron Microscopy (Cryo-EM)

The structural characteristics of the nanodiscs were analyzed using Cryo-EM electron tomograph. A 3 μL aliquot of the polymer self-assembly solution was carefully applied onto glow-discharged Lacey carbon-coated grids. Following sample deposition, the grids were rapidly vitrified by plunge-freezing into liquid ethane under controlled temperature and humidity conditions using a Vitrobot Mark IV (Thermo Fisher Scientific). The frozen grids were then transferred into a Thermo Fisher Scientific Glacios Cryo-EM operating at 200 kV with a field-emission gun. Image data were recorded using a Falcon 3 direct electron detector, and the automated tilt-series acquisition was handled by the Thermo Fisher Tomography software. Particle identification and reconstruction across multiple tilt angles were subsequently performed using CryoSPARC (v4).

Synthetic Procedures

Synthesis of the Polymer Backbone

The polymer backbone was synthesized via a three step procedure following the detailed protocol from our previous work. In brief, mPEG 5K (1.0 g, 0.2 mmol, 1.0 equiv) and CDTPA (105 mg, 0.26 mmol, 1.3 equiv) were dissolved in 5 mL of anhydrous dichloromethane (DCM) in a dry round-bottom flask. The flask was cooled in an ice bath and stirred vigorously. Meanwhile, a mixture of EDC·HCl (50 mg, 0.26 mmol, 1.3 equiv) and DMAP (3 mg, 0.02 mmol, 0.12 equiv) in 2 mL anhydrous DCM was slowly added dropwise. The reaction was then allowed to proceed overnight at ambient temperature. The macro-RAFT agent was subsequently isolated via precipitation and extraction. Next, the purified macro-RAFT (100 mg, 0.02 mmol, 1.0 equiv) was chain-extended by ZnTPP-catalyzed PET-RAFT polymerization with GMA monomers (53 mg, 0.38 mmol, 20 equiv) for 24 h, reaching over 95% conversion. The resultant polymers were purified, dried, and characterized by ¹H NMR (300 MHz, CDCl₃, δ, ppm), SEC (DMAc/LiBr, 50 °C, PMMA). Finally, azide-functionalized block copolymers, PEG₁₁₄-b-(PGMA-N₃)₂₀, were synthesized through ring-opening of the epoxide groups on PEG₁₁₄-b-PGMA₂₀ using sodium azide at 50 °C for 48 h. The intermediates and final polymers were analyzed by ¹H NMR (300 MHz, CDCl₃, δ, ppm), SEC (DMAc/LiBr, 50 °C, PMMA), and FTIR.

Synthesis of Hydrophobic Acetal Glycidyl Ethers

Ethoxy ethyl glycidyl ether (EE) was prepared following the method reported by Song et al. In brief, ethoxy ethyl vinyl ether (21 mL, 225 mmol) was dissolved in 150 mL of chilled dichloromethane (DCM) and stirred for 10 min in an ice bath. Subsequently, p-toluenesulfonic acid (p-TsOH, 0.258 g, 1.5 mmol) was added dropwise. After completion of the addition, the ice bath was removed, and the reaction mixture was allowed to proceed at room temperature for an additional 2 h. The reaction was then quenched by the slow addition of saturated sodium bicarbonate solution. The organic phase was separated, washed thoroughly with water, and dried over anhydrous sodium sulfate. The crude product was initially concentrated using a rotary evaporator and further purified via vacuum distillation. The purified EE was then purged with nitrogen gas and stored in an argon-filled glovebox over 3 Å molecular sieves to maintain dryness.

Preparation of Polyether Copolymers

A series of hydrophobic PEE_n polymers were synthesized by varying the feed ratios of EE monomers inside an argon-filled glovebox. In a flame-dried Schlenk flask, benzyl alcohol (20.8 μL, 0.20 mmol), t-BuP₄ (0.08 mL, 0.07 mmol, 0.8 M in hexane), and 1 mL of dry toluene were combined and stirred at 25 °C for 10 min. Subsequently, EE monomers (0.41, 0.82, and 1.23 mL; 2.8, 5.6, and 8.4 mmol, respectively) were added to obtain PEE₁₄, PEE₂₈, and PEE₄₂. Polymerization was allowed to proceed for 24 h. Reaction aliquots were analyzed to confirm complete monomer conversion and a narrow molecular weight distribution by SEC. The resulting crude polymers were directly used for the subsequent postpolymerization modification reactions without further purification. The aliquots were characterized by ¹H NMR (400 MHz, CDCl₃, δ, ppm).

Preparation of Alkyne Functionalized Polyether Copolymer Side Chains for Grafting-To

Alkyne-functionalized polyether copolymer side chains (EE) were synthesized via propargylation of the corresponding EE_n polymers in one-pot reaction. For example, to prepare EE₁₄, a 50 wt % NaOH aqueous solution (0.2 mL) and tetrabutylammonium bromide (5 mg, 0.016 mmol) were added to a crude solution of EE₁₄, and the mixture was stirred vigorously for 10 min. Propargyl bromide (0.11 mL, 1.0 mmol, 80 wt % in toluene) was then slowly added dropwise, and the reaction temperature was raised to 50 °C. After 24 h, the reaction mixture was transferred to a separatory funnel, diluted with diethyl ether, and washed multiple times with water to remove residual NaOH and NaBr byproducts. The organic phase was dried over anhydrous magnesium sulfate and concentrated under reduced pressure. To eliminate any remaining propargyl bromide, the crude product was redissolved in toluene and dried again under vacuum, yielding the final EE₁₄ product. EE₂₈ and EE₄₂ were modified using the same procedure. All purified products were characterized by ¹H NMR (300 MHz, CDCl₃, δ, ppm), SEC (DMAc/LiBr, 50 °C, PMMA).

Synthesis of Bottlebrush Block Copolymers

The polymers were synthesized through copper-catalyzed azide–alkyne cycloaddition (CuAAC) click reactions. Typically, PEG₁₁₄-b-(PGMA-N₃)₂₀ (10 mg, 0.023 mmol, 1 equiv), EE₁₄ (61 mg, 0.028 mmol, 1.2 equiv), Cy5 alkyne (0.990 mg, 0.001 mmol, 0.05 equiv), PMDETA (8 mg, 0.046 mmol, 2 equiv), and 1 mL of DMF were combined in a Schlenk flask. The reaction mixture underwent three freeze–pump–thaw cycles to remove dissolved gases, after which CuBr (7 mg, 0.046 mmol, 2 equiv) was added to initiate the click reaction. The mixture was stirred at room temperature for 48 h and subsequently quenched by exposure to air. The reaction solution was diluted with DCM and passed through a neutral alumina column to remove copper residues. The filtrate was concentrated under reduced pressure and precipitated into a diethyl ether/petroleum benzene mixture (2:8 v/v) to eliminate unreacted EE₁₄. This dissolution–precipitation cycle was repeated twice for further purification. The resulting product, PE₁₄C, was dried under high vacuum, yielding an off-white viscous solid. The degree of grafting of EE chains was determined as described below.

The PEG₁₁₄ segment served as an internal reference. For example, the number of grafted chains (N) is calculated to be 15 in the case of PE₁₄C as follows

$$[14(\mathrm{red}\mathrm{labeled}\mathrm{area}\mathrm{in}\mathrm{Figure}\mathrm{S}12)\times \mathrm{number}\mathrm{of}\mathrm{grafted}\mathrm{chains}(N)]/[(100(\mathrm{blue}\mathrm{labeled}\mathrm{area}\mathrm{in}\mathrm{Figure}\mathrm{S}12)\times N)+114\times 4(\mathrm{internal}\mathrm{standard})]=\frac{14.00(\mathrm{red}\mathrm{labeled}\mathrm{area}\mathrm{in}\mathrm{Figure}\mathrm{S}15)}{130.77(\mathrm{blue}\mathrm{labeled}\mathrm{area\; in}\mathrm{Figure}\mathrm{S}15)}$$

The grafting number N for PE₁₄C is calculated to be 15, corresponding to a grafting efficiency of 75% (Figure S13). Similar calculations for PE₂₈C and PE₄₂C (Figures S14 and S15), also indicate grafting efficiencies of approximately 75%.

The preparation of PE₂₈C and PE₄₂C followed the same procedure as PE₁₄C, with adjustments to the weight of EE₂₈ and EE₄₂ to maintain comparable grafting efficiency. Characterization can be found in Supporting Information (Figures S12–S14).

Preparation of PE_nC Self-Assemblies in Water

In a typical procedure, 10 mg of PE₁₄C polymer was dissolved in 5 mL of DMF and stirred at room temperature for 2 h. The solution was then transferred into a dialysis bag (MWCO ∼ 3000 Da) and dialyzed against deionized water for 3 days, with the dialysis medium replaced twice daily. The final concentration of the PE₁₄C self-assemblies was adjusted to 1 mg/mL by dilution with deionized water. The size and morphology of the self-assemblies were characterized by DLS, AFM, and TEM.

Degradation Studies of PE₂₈C Nanodiscs in Acidic and Neutral Environments

Self-assembled PE₂₈C nanodisc solutions at 1 mg mL⁻¹ were transferred into dialysis bags (MWCO ∼ 3000 Da) and dialyzed against acetate buffer (pH 5.0). The dialysis medium was refreshed twice daily. At predetermined time points, samples inside the dialysis bags were collected for DLS and TEM analyses to observe changes in particle size and morphology over time. For comparison, nanodisc stability under neutral pH conditions was evaluated by dialyzing the samples against deionized water under otherwise identical conditions.

Loading Capacity and A-205804 Release Profiles of Nanodiscs in Acidic Environments

A-205804 was selected as the model drug to evaluate the loading capacity of the PE₂₈C nanodiscs. Drug loading was performed via a solvent displacement method, analogous to the PE₂₈C nanodisc preparation described above. Briefly, 5 mg of A-205804 and 10 mg of PE₂₈C polymer were dissolved in 5 mL of DMF and stirred for 2 h. The resulting solution was transferred into a dialysis bag (MWCO ∼ 3000 Da) and dialyzed against deionized water in the dark for 3 days to facilitate nanodisc self-assembly, encapsulation of A-205804 molecule, and removal of free A-205804 and DMF. The dialysis medium was refreshed twice daily. The final concentration of A-205804@PE₂₈C self-assemblies was adjusted to 1 mg/mL by dilution with deionized water and characterized by DLS.

To determine drug loading and encapsulation efficiency, a portion of the solution was freeze-dried and weighed. The dried product was redissolved in DMF, and the absorbance was measured using a UV–vis spectrophotometer. The concentration of A-205804 was calculated based on a previously established calibration curve. Loading efficiency (%) and drug content (%) were calculated using the following equations: Loading efficiency = (mass of loaded drug/mass of actual drug used)×100%. Drug content = (mass of loaded drug/mass of nanoparticles with drug)×100%.

The release profile of A-205804 from A-205804@PE₂₈C was evaluated under different conditions. A portion of the sample was transferred into a dialysis bag and dialyzed against acetate buffer (pH = 5.0) to mimic acidic environments, while the remaining portion was dialyzed against deionized water in the dark. The external dialysis medium was replaced twice daily to maintain sink conditions. At predetermined time points, aliquots were withdrawn from inside the dialysis bag, and the absorbance was measured to determine the cumulative release of A-205804.

In Vitro Experiments

Cell Culture

MDA-MB-231 cells and HeLa cells were maintained in complete DMEM supplemented with 2% (v/v) FBS and 1% (v/v) antibiotic–antimycotic. Cells were incubated at 37 °C in a humidified atmosphere containing 5% CO₂.

MTT Assay

MDA-MB-231 cells and HeLa cells were seeded in 96-well plates at a density of 1.0 × 10⁴ cells per well and incubated for 24 h to allow for cell attachment. The culture medium was then replaced with fresh medium containing PE_nC nanodiscs at varying concentrations. Cells were treated for either 24 or 48 h. At each time point, 10 μL of MTT solution (5 mg mL^–1) was added to each well, followed by incubation in the dark at 37 °C for 4 h. Formazan crystals formed during the reaction were dissolved by adding DMSO. Absorbance was measured at 595 nm using a plate reader, and cell viability was calculated based on the optical density (OD₅₉₅) values.

Confocal Laser Scanning Microscope (CLSM)

MDA-MB-231 cells were seeded at a density of 5.0 × 10⁴ cells per dish in 3.5 cm glass-bottom confocal dishes and incubated for 24 h to allow for attachment. The cells were then treated with PE_nC nanodiscs at a concentration of 2 nmol mL^–1 for 24 h at 37 °C. Following incubation, the cells were washed three times with DPBS and maintained in fresh culture medium. For nuclear staining, one drop of NucBlue Live ReadyProbes reagent was added per dish, followed by incubation in the dark at 37 °C for 20 min. Fluorescence imaging was performed using an Olympus FluoView FV3000 confocal laser scanning microscope equipped with a 60× oil-immersion objective. Cy5 fluorescence signals were monitored to visualize the intracellular distribution of the nanodiscs.

Flow Cytometry

MDA-MB-231 cells and HeLa cells were seeded in 6-well plates at a density of 5 × 10⁵ cells per well and incubated for 24 h to allow cell attachment. Cells were then treated with PE_nC nanodiscs at a concentration of 2 nmol mL^–1 for 24 h. Untreated cells were used as a control. After treatment, the culture medium was removed, and the cells were washed three times with DPBS. Cells were detached using trypsin, and 1 mL of DPBS was added to each well. The suspensions were centrifuged at 1000 rpm for 5 min, washed twice with DPBS, and finally resuspended in 0.5 mL of DPBS. Data from 1 × 10⁴ gated events were acquired and analyzed using a BD FACSCalibur flow cytometer.

Lysosome Colocalization Assay

MDA-MB-231 cells were seeded in 3.5 cm glass-bottom confocal dishes at a density of 5.0 × 10⁴ cells per dish and incubated for 24 h to allow cell attachment. Cells were then incubated with Cy5-labeled PE₂₈C nanodiscs at a concentration of 2 nmol mL^–1 for 24 h. After incubation, the culture medium was removed and the cells were washed three times with DPBS. Cells were subsequently stained with LysoTracker Red at a final concentration of 100 nM in serum-free medium for 30 min at 37 °C. Cells were then washed with DPBS to remove excess dye. For nuclear staining, one drop of NucBlue Live ReadyProbes reagent was added per dish, followed by incubation in the dark at 37 °C for 20 min. Fluorescence imaging was performed using an Olympus FluoView FV3000 confocal laser scanning microscope equipped with a 60× oil-immersion objective. Cy5 and LysoTracker Red fluorescence signals were collected in separate channels to visualize the intracellular distribution of the nanodiscs and lysosomes, respectively. Colocalization analysis was performed using ImageJ with the Coloc 2 plugin. Pearson’s correlation coefficient and Manders’ overlap coefficients were calculated to quantify the degree of colocalization between the PE₂₈C (Cy5) and LysoTracker (green) channels (Table S1).

Wound Healing Assay

Cell migration was evaluated using a wound healing assay with ibidi Culture-Inserts. MDA-MB-231 cells were seeded at 3 × 10⁴ cells per chamber and incubated for 24 h to allow cell attachment. After incubation, the culture inserts were carefully removed to create a defined cell-free gap, and treatment conditions were applied. Cell migration into the wound area was monitored after 48 h using confocal laser scanning microscopy (CLSM). The gap closure was quantified by measuring the wound area at 0 and 48 h using ImageJ software.

Transwell Migration Assays

Cell migration was assessed using Transwell chambers. Twenty-four h-starved MDA-MB-231 cells were digested and resuspended in serum-free medium with PE_nC nanodiscs. A total of 1 × 10⁵ cells were seeded into the upper chamber, while the lower chamber was filled with DMEM supplemented with 10% FBS as a chemoattractant. After 24 h of incubation, nonmigrated cells remaining on the upper side of the membrane were carefully removed using a cotton swab. Migrated cells on the lower compartment were fixed and stained with 0.1% crystal violet solution for 20 min, followed by washing and air drying. The number of migrated cells in the lower compartment was observed under Leica THUNDER Imager Live Cell.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/jacsau.6c00206.

• Results and data, including a detailed characterization of polymers via NMR, SEC, and IR; further analysis of the self-assembled nanomaterials via DLS, AFM, TEM, and cryo-TEM; and biological assessments via cell studies, flow cytometry, and confocal microscopy (PDF)

P.Z. and M.M. codesigned the project. P.Z. performed all syntheses and experiments, except for polyether side chains, which were synthesized by J.B. and B.Y. under supervision from B.-S.K. H.Z. aided in project design, syntheses, and analysis. P.Z., H.Z., and M.M. wrote the manuscript with input from all authors. CRediT: Ping Zeng conceptualization, data curation, formal analysis, investigation, methodology, visualization, writing - original draft; Haoxiang Zeng data curation, investigation, methodology, writing - original draft; Jinsu Baek data curation, methodology, resources, writing - review & editing; Byungwoo Yoo resources; Byeong-Su Kim funding acquisition, resources, supervision, validation, writing - review & editing; Markus Müllner conceptualization, funding acquisition, investigation, methodology, project administration, resources, supervision, writing - original draft.

Australian Research Council for a Future Fellowship (FT200100185) and Discovery Project (DP220100452), National Research Foundation of Korea (RS-2025-00558644), and Global Science and Technology Diplomacy Fund grant (GSTDS000001 – 6).

The authors declare no competing financial interest.