Engineering Folic Acid‐Modified Nanoparticles to Enhance Letrozole's Anticancer Action

Abstract

The development of biodegradable nanoparticles (NPs) for delivering anticancer drugs, such as letrozole (LTZ), offers a targeted approach for cancer therapy. In this study, we synthesized poly(ε‐caprolactone)‐co‐poly(ethylene glycol) (PCL‐co‐PEG) and fabricated LTZ‐loaded PCL‐co‐PEG NPs (LTZ‐NPs) via emulsion‐solvent evaporation. Folic acid (FA), a folate receptor‐targeting molecule, was conjugated to the LTZ‐loaded NPs (LTZ‐FNPs) to enhance treatment efficacy against hormone receptor‐positive breast cancer cells. Both NPs and FNPs exhibited a spherical morphology (60–90 nm), with FNPs showing higher drug entrapment efficiency and controlled release. LTZ release was minimal at physiological pH but increased in acidic, cancer‐like environments, following the Korsmeyer‐Peppas model, indicating a combination of Fickian and non‐Fickian diffusion. In cytotoxicity assays, LTZ‐FNPs exhibited higher toxicity against MCF‐7 cells than LTZ‐NPs. Controlled LTZ release altered gene expression, reducing B‐cell leukemia/lymphoma 2 protein (Bcl2) and increasing caspase 8 (Casp8), promoting apoptosis. A shift to the SubG1 phase further confirmed enhanced LTZ‐FNP‐mediated cell death. Furthermore, p53 expression increased, while matrix metalloproteinase 9 (MMP‐9) decreased, inhibiting cell invasion. This study introduces a biodegradable system with FA‐functionalized, pH‐sensitive NPs for the targeted and controlled delivery of LTZ. This approach holds great potential for selective, efficient treatment while minimizing systemic toxicity in breast cancer therapy.

Folate‐functionalized, pH‐sensitive nanoparticles (NPs) are developed for the targeted delivery of letrozole (LTZ) in breast cancer therapy. LTZ‐loaded PCL‐co‐PEG NPs exhibit controlled release, enhanced cytotoxicity, and apoptosis induction in MCF‐7 cells. Functionalization with folic acid improves drug entrapment and release, reducing invasion markers. This biodegradable system offers a promising strategy for selective and efficient cancer treatment.

1. Introduction

Cancer remains one of the leading causes of mortality worldwide, presenting an ongoing challenge to global public health. In response to this critical issue, researchers have developed diverse therapeutic approaches, including small molecules, aptamers, peptides, and proteins.^{[ 1} , ^{2 ]}

Among all cancer types, breast cancer stands out as particularly significant, representing the most prevalent cancer diagnosis in women and a major contributor to cancer‐related mortality.^{[ 3} , ^{4 ]} For hormone receptor‐positive breast cancers, which constitute a predominant subtype, aromatase inhibitors have emerged as a cornerstone of treatment.^{[ 5} , ^{6 ]} These therapeutic agents function by disrupting the estrogen signaling pathway, which plays a fundamental role in cancer cell proliferation and survival.^{[ 7 ]}

LTZ, a notable member of the aromatase inhibitor class, is a standard adjuvant therapy for postmenopausal women diagnosed with early‐stage hormone receptor‐positive breast cancer.^{[ 8} , ^{9 ]} Clinical evidence demonstrates that this therapeutic strategy significantly reduces cancer recurrence risk and improves overall survival rates, underscoring the vital role of aromatase inhibition in breast cancer treatment.^{[ 10 ]} However, despite these promising therapeutic outcomes, LTZ's clinical efficacy is hampered by its poor aqueous solubility, which presents a significant barrier to optimal drug delivery and effectiveness.^{[ 11 ]}

Recent advancements in analytical techniques have greatly enhanced our understanding of LTZ pharmacology and its delivery systems.^{[ 12} , ^{13 ]} Modern methods such as mass spectrometry and dynamic light scattering enable precise characterization of drug‐nanocarrier interactions, particle size distribution, drug loading efficiency, and release kinetics. These insights are critical for optimizing nanoparticle (NP)‐based drug delivery formulations.^{[ 14} , ^{15 ]}

Sophisticated imaging technologies, including scanning electron microscopy (SEM), transmission electron microscopy, and atomic force microscopy, provide detailed structural analyses of nanocarriers.^{[ 16} , ^{17 ]} These methods reveal NP morphology, surface characteristics, and drug distribution, all of which are essential for developing systems that enhance LTZ's bioavailability, hydrophilicity, and therapeutic efficacy while minimizing toxicity.^{[ 18} , ¹⁹ , ²⁰ , ²¹ , ²² , ^{23 ]} Among these, polymeric NPs stand out due to their biocompatibility, biodegradability, and ability to provide sustained drug release. Polymers such as polycaprolactone (PCL) offer versatility in synthesis, enabling precise control over size, surface properties, biodegradation, and release profiles.^{[ 24} , ²⁵ , ^{26 ]} For example, poly(ε‐caprolactone)‐co‐poly(ethylene glycol) (PCL‐co‐PEG) exhibit amphiphilic properties, allowing encapsulation of both hydrophilic and hydrophobic drugs while ensuring controlled release and prolonged circulation.^{[ 27 ]}

Surface modifications of polymeric NPs, such as incorporating targeting ligands or stealth coatings, enhance tumor targeting and reduce immune recognition.^{[ 28 ]} Functionalizing NPs with molecules like peptides^{[ 29 ]} or folic acid (FA) enables precise targeting of overexpressed folate receptors (FRs) on cancer cells, particularly FRα, which is upregulated in breast cancer.^{[ 30 ]} Studies have shown that FA‐conjugated NPs increase cellular uptake in MCF‐7 breast cancer cells, leading to enhanced therapeutic outcomes.^{[ 31 ]} Targeting FRs minimizes off‐target effects and improves drug delivery efficiency.^{[ 32 ]} Furthermore, conjugating FA to the surface of NPs can enhance steric stability by adding a hydrophilic layer, which reduces aggregation and improves colloidal stability.^{[ 33 ]} Additionally, FA conjugation can influence drug release characteristics, enabling more controlled release in tumor environments and helping maintain the stability and effectiveness of the system.^{[ 34 ]}

The primary objective of this research was to design FA‐functionalized NPs for the efficient delivery of LTZ, a hydrophobic agent, in cancer‐like acidic environments.^{[ 35 ]} In this study, we synthesized and characterized LTZ‐loaded PCL‐co‐PEG NPs using various methods. Molecular docking was employed to investigate the interactions between the copolymers and LTZ, providing insights into the mechanisms underlying their effectiveness as drug‐delivery vehicles. The release profile and biological performance of the drug delivery system (DDS) were rigorously evaluated through biological assays. This work introduces a novel DDS that combines FA functionalization with pH‐sensitive, biodegradable NPs for targeted breast cancer therapy, enhancing LTZ specificity and therapeutic efficacy, and advancing nanomedicine for more precise cancer treatment.

2. Experimental Section

2.1. Materials

ε‐caprolactone and polyvinyl alcohol (PVA) were obtained from Sigma‐Aldrich (St. Louis, MO). N‐hydroxy‐succinimide (NHS), poly(ethylene glycol) amine (HO‐PEG‐NH₂, 3 kDa), chloroform, deuterated chloroform (CDCl₃), dimethylsulfoxide (DMSO), diethyl ether, dichloromethane (DCM), N,N′‐dicyclo‐hexylcarbodiimide (DCC), ethanol (EtOH), stannous octoate (Sn(Oct)₂), 3‐(4,5‐dimethylthiazol‐2‐yl)‐2,5‐diphenyltetrazolium bromide (MTT), propidium iodide (PI), and FA were purchased from Merck (Munich, Germany). Human breast cancer cells (MCF‐7) were supplied by the Pasteur Institute of Iran. Fetal bovine serum (FBS), streptomycin, trypsin, Dulbecco's Modified Eagle Medium (DMEM), and penicillin were purchased from Kalazist (Tehran, Iran). LTZ was generously gifted by Aburaihan Pharmaceutical company (Tehran, Iran).

2.2. Synthesis of LTZ‐FNPs

2.2.1. Synthesis of PCL‐co‐PEG

The synthesis of amphiphilic PCL‐co‐PEG‐NH₂ followed a previously established method.^{[ 36 ]} The copolymers were obtained through the ring‐opening polymerization of ε‐caprolactone, using HO‐PEG‐NH₂ as the starting material and Sn(Oct)₂ as the catalyst. Initially, a three‐neck round‐bottom flask equipped with a magnetic stirrer and nitrogen inlet was charged with HO‐PEG‐NH₂ (20 mg). The flask was connected to a vacuum line and heated in an oil bath at 130 ± 1 °C for 2 h to remove residual moisture and any volatile impurities. Subsequently, ε‐caprolactone (40 g) and Sn(Oct)₂ solution in toluene were introduced into the flask under positive nitrogen pressure (≈0.1 Torr) using degassed syringes. The polymerization reaction was conducted with continuous stirring at 400 rpm at 130 ± 1 °C for 20 h. The progress of the reaction was monitored by tracking the disappearance of the ε‐caprolactone monomer peak using ¹H NMR spectroscopy. After completion, the reaction mixture was cooled to room temperature, and the crude product was dissolved in chloroform (40 mL). The polymer solution was then precipitated dropwise into cold diethyl ether at 4 °C under vigorous stirring. The precipitated copolymer was collected by filtration using a Buchner funnel and dried under vacuum (0.1 mmHg) at room temperature for 24 h. Further purification was achieved by centrifugation at 10 000 rpm for 30 min in deionized water to remove any unreacted monomer and low molecular weight species. The centrifugation and washing steps were repeated three times using fresh deionized water. Finally, the purified copolymer was lyophilized for 48 h to obtain a white powder. The final product was characterized by ¹H NMR and FTIR to confirm its chemical structure.

2.2.2. FA Conjugation

FA was conjugated to PCL‐co‐PEG‐NH₂ through a DCC‐mediated reaction. In a beaker, 2 mg of FA was dissolved in 10 ml of DMSO, followed by the addition of 0.1 mg of NHS and 0.2 mg of DCC to the solution. The mixture was stirred for 3 h in the dark at room temperature to ensure full activation. Next, 15 mg of the PCL‐co‐PEG‐NH₂ solution was added to the activated FA solution. Additionally, 20 mL of DMSO was introduced to the solution to facilitate the continuation of the reaction. The conjugation reaction was allowed to proceed overnight under stirring in the dark at 30 °C. This process resulted in the successful conjugation of FA to PCL‐co‐PEG‐NH₂ via the formation of amide bonds (Figure 1 ).^{[ 37} , ^{38 ]} FA‐functionalized PCL‐co‐PEG was then purified by centrifugation to remove unreacted FA, following the same purification steps as described earlier. Proton nuclear magnetic resonance (¹H NMR; Bruker Ac 500, Germany, 500 MHz in CDCl₃) and Fourier transform infrared spectroscopy (FTIR; Nicolet 550 A, USA) were used to characterize FA‐functionalized PCL‐co‐PEG block copolymer.

Figure 1.

A streamlined process for the synthesis of PCL‐co‐PEG‐NH₂ and PCL‐co‐PEG‐FA. The synthesis of PCL‐co‐PEG‐NH₂ and its functionalization with FA require several chemical reactions (Step 1: HO‐PEG‐NH₂ initiates the ring‐opening polymerization of ε‐caprolactone, forming PCL‐co‐PEG‐NH₂. Step 2: FA reacts with NHS/DCC to form FA‐NHS. Step 3: FA‐NHS reacts with PCL‐co‐PEG‐NH₂ to form PCL‐co‐PEG‐FA through an NHS‐mediated coupling reaction).

2.2.3. NP Preparation

LTZ‐loaded NPs were prepared using an emulsion method. Initially, 10 mg of LTZ was dissolved in 4 mL of DCM, and this solution was added drop‐wise into a mixture of 10 mg of PCL‐co‐PEG or FA‐functionalized PCL‐co‐PEG dissolved in 15 mL of DCM. The two‐phase mixture was sonicated for 50 s in an ice bath to facilitate emulsification. A 3% (w/v) solution of PVA was prepared by dissolving the appropriate amount of PVA in distilled water under continuous stirring, and 9 mL of this solution was then added to the emulsion. The mixture was sonicated for an additional 50 s to form a water‐in‐oil (w/o) single emulsion. The resulting emulsion was stirred for 20 min to ensure uniform distribution. To remove the organic solvent, the emulsion was diluted with 50 mL of a 0.3% (w/v) aqueous solution of PVA under constant stirring for 10 min. Next, the organic solvent was removed using a rotary evaporator. The PCL‐co‐PEG NPs (NPs) or FA‐functionalized PCL‐co‐PEG (FNPs) formed during the process were then subjected to centrifugation at 11 000 rpm for 30 min to isolate them. Subsequently, they were washed twice with deionized water to remove any remaining impurities and lyophilized to obtain a dry powder for further analysis and use (Figure 2 ). The surface charge and particle size of the NPs were determined by dynamic light scattering (DLS, Malvern Zeta sizer 3000HS, Malvern, UK). Additionally, the morphology of the sample was examined using SEM (FEI, California, USA). Contact angle measurements (SDC100, Minder‐Hightech, China) were conducted to evaluate the amphiphilic properties of LTZ‐NPs and LTZ‐FNPs. Borosilicate glass slides served as the substrate, with a sample concentration of 0.6% w/v. The droplet size was standardized across all measurements to ensure reproducibility.

Figure 2.

NP preparation using the water‐in‐oil (W/O) emulsion method. The schematic describes the process, which involves several steps: A) dissolution of the polymer in the organic phase, B) addition of the aqueous phase containing the active compound, C) emulsification through high‐speed homogenization, D) solvent evaporation leading to NP formation, and E) collection of the NPs after centrifugation and washing.

2.3. Docking Methodology

In this study, docking procedures were carried out to investigate the interaction patterns between LTZ and the copolymers.^{[ 20} , ³⁹ , ^{40 ]} Prior to the docking process, nanosystems were designed using ChemDraw (v19.0) and prepared using Autodock tools (v1.5.6). The complexes were assigned charges using the Gasteiger algorithm. Docking of NPs and FNPs with LTZ was performed using AutoDock Vina, an open‐source tool. Polar hydrogens were assigned to the complexes before docking.^{[ 41} , ^{42 ]} The desired grid box for the systems was created with dimensions of 30 × 30 × 30 Å and a spacing value of 0.258 Å. The genetic algorithm was employed to explore conformations, with GA runs and a population size set at 15 and 200, respectively.^{[ 39} , ^{41 ]} Interactions were visualized using the Discovery Studio Visualizer (v2021).

2.4. Evaluation of Encapsulation Efficiency (EE)

To evaluate the EE, formulation parameters were systematically optimized. Solvent ratios of EtOH/PBS were tested at 5:95, 10:90, 15:85, and 20:80 v/v, with a 10:90 v/v ratio identified as optimal for drug extraction. Centrifugation speed (8000—15 000 rpm) and duration (10–30 min) were adjusted to achieve complete separation, with ideal conditions established. Additionally, UV/Vis (Taitec, BR‐42FL, Japan) was performed using multiple wavelength scans ranging from 200 to 400 nm, and the results identified 239 nm as the optimal wavelength for detecting LTZ. These optimizations ensured accurate and reproducible results for EE assessment. The drug loading (DL) and EE were calculated using Equations (1) and (2):^{[ 43} , ^{44 ]}

$$\mathrm{DL}(\%)=\frac{\mathrm{Drug}\mathrm{weight}}{\mathrm{Weight}\mathrm{of}\mathrm{drug}\mathrm{-}\mathrm{loaded}\mathrm{nanocarriers}}\times 100$$ (1)

$$\mathrm{EE}(\%)=\frac{\mathrm{Weight}\mathrm{of}\mathrm{entrapped}\mathrm{drug}\mathrm{in}\mathrm{nanocarriers}}{\mathrm{Weight}\mathrm{of}\mathrm{feeding}\mathrm{drug}}\times 100$$ (2)

2.5. Drug Release Study

The release of LTZ was assessed using a dialysis method. LTZ‐NPs and FNPs, with a 10:90 w/w ratio, were prepared by dispersing 2 mg of each formulation in 2 mL of an EtOH/PBS solution (10:90 v/v). For the dialysis process, NP samples were placed in 12 kDa dialysis membranes, which were immersed in 50 mL of EtOH/PBS solution (10:90 v/v) at pH 5.4 and 7.4, with continuous stirring at 37 °C. Each formulation was placed in a separate dialysis bag and incubated in its respective EtOH/PBS solution. Drug release was then monitored over time. This method enabled the assessment of drug release profiles under various pH conditions, simulating both the physiological environment and the acidic tumor microenvironment. Additionally, the concentration of LTZ was determined using UV/Vis at 239 nm. LTZ release profiles were obtained for the various samples and evaluated using the Korsmeyer–Peppas model,^{[ 44 ]} which was widely recognized as a comprehensive semi‐empirical equation for describing drug release from polymeric systems. The release kinetics of LTZ released can be described by Equation (3), where various parameters influence the release dynamics:

$$f_1=\frac{M_i}{M_{\infty }}=K{t}^n$$ (3)

f₁ represents the percentage of released LTZ. M _i stands for the quantity of released LTZ. M _∞ denotes the quantity of LTZ at equilibrium. K is the constant that encapsulates the integration of structural alterations and geometric attributes of the system. n represents the release exponent as a function of time (t), with values typically ranging from 0.5 to 1. When n = 0.5, the release follows a Fickian diffusion model, where drug release was primarily controlled by diffusion. In this scenario, the rate at which the solvent diffused through the polymer was much faster than the relaxation of the polymer chains. As a result, the polymer surface quickly reached absorption equilibrium, and the drug release was governed by time‐dependent diffusion. The kinetics of this process were characterized by the diffusivity of the drug through the polymer matrix. When n = 1, a non‐Fickian release occurs, indicating that other mechanisms, such as polymer relaxation, dominate. For values of n between 0.5 and 1, an anomalous release (non‐Fickian) pattern was observed, suggesting that the drug release mechanism was influenced by both diffusion and polymer swelling dynamics.^{[ 45 ]}

2.6. Cell Culture

MCF‐7 cells were cultured at 37 °C in a humidified atmosphere containing 95% air and 5% CO₂. The culture medium consisted of DMEM supplemented with 10% (v/v) FBS, 100 µg mL⁻¹ streptomycin, and 100 U mL⁻¹ penicillin.^{[ 46 ]} All cell culture processes were conducted under sterile conditions using a class II biological safety cabinet.

2.7. Cytotoxicity Assay

The cytotoxicity of the NPs was assessed using the MTT assay. Initially, cells (7.5 × 10³ cells well⁻¹) were cultured under standard physiological conditions. Cells were then exposed to various concentrations (30—210 nM) of LTZ‐free NPs, LTZ‐NPs, LTZ‐free FNPs, and LTZ‐FNPs for 48 h. Subsequently, an MTT solution (5 mg mL⁻¹ MTT in sterile PBS) was added to each well (20 µL well⁻¹) and incubated for 4 h at 37 °C. After incubation, the medium was aspirated, and 60 µL of DMSO was added to each well. The samples were then further incubated for 15 min with gentle shaking.^{[ 21 ]} The absorbance values were measured at 590 nm using a plate reader (⁺MR4, Hyperion, Germany), with background absorbance at 620 nm subtracted from the readings at 590 nm for all groups. PBS‐treated cells were considered as the control group. Cytotoxicity was calculated as depicted in Equation (4).

$$\begin{array}{c}\hfill \mathrm{Cytotoxicity}\left(\%\right)=\left[1-\left(\frac{\mathrm{Absorbance}\mathrm{of}\mathrm{treated}\mathrm{cells}}{\mathrm{Absorbance}\mathrm{of}\mathrm{control}\mathrm{cells}}\right)\right]\times 100\end{array}$$ (4)

2.8. Analysis of Gene Expression by RT‐qPCR

Gene expression analysis was conducted to investigate the impact of NPs on apoptotic and tumor suppressor genes, following established protocols.^{[ 47 ]} Reverse Transcription quantitative polymerase chain reaction (RT‐qPCR) was used to measure the expression levels of key genes: Bcl2, Casp8, MMP‐9, and p53 in MCF‐7 cells treated with free LTZ, LTZ‐NPs, and LTZ‐FNPs. Total RNA was extracted from the cells and converted to complementary DNA using a Qiagen kit (Germantown, MD, USA). The specific primers used for the gene expression analysis are detailed in Table 1 .^{[ 47 ]} Glyceraldehyde‐3‐phosphate dehydrogenase (GAPDH) was utilized as the reference gene to normalize the gene expression data. The RT‐qPCR reaction conditions were as follows: 95 °C for 10 min (pre‐incubation step), followed by 40 cycles of 95 °C for 30 s, 60 °C for 30 s, and 72 °C for 30 s (amplification step). The 2^−ΔΔCt method was employed to evaluate and compare changes in gene expression between the different treatment groups.^{[ 48 ]}

Table 1.

Designed primers for target gene amplification. Primers were designed using the NCBI primer design tool.

Gene	Primer sequence	Strand	Product size
GAPDH	TGGAAGGACTCATGACCACA	Plus	119
	AGAGGCAGGGATGATGTTCT	Minus
Bcl2	TGGGATCGTTGCCTTATGCA	Plus	101
	GTCTACTTCCTCTGTGATGTTGT	Minus
MMP‐9	GATGCCTGCAACGTGAACAT	Plus	88
	AGAATCGCCAGTACTTCCCATC	Minus
Casp8	AAGGAGCTGCTCTTCCGAAT	Plus	134
	CGGCAGAAGTGGAACCTGTA	Minus
P53	TGTGACTTGCACGTACTCCC	Plus	84
	GAATCAACCCACAGCTGCAC	Minus

2.9. Cell Cycle Analysis

MCF‐7 cells (1 × 10⁶ per well) were seeded into 6‐well plates and incubated in DMEM for 24 h to allow cell attachment and growth. After incubation, cells were treated with different samples at the half‐maximal inhibitory concentration (IC₅₀) of LTZ for 48 h. Following treatment, the cells were detached from the plates, fixed with 70% EtOH at −20 °C overnight to preserve cell morphology and prevent further cellular processes, and then stained with 500 µL of a PI solution containing RNase. The staining procedure was performed in the dark for 30 min at 25 °C to minimize photobleaching of the PI. The stained samples were subsequently analyzed using flow cytometry to assess cell cycle analysis.

2.10. Cell Invasion Assay

The Transwell system (SPL Life Sciences Co., Ltd., Pochon, Kyonggi‐do, South Korea) was set up by placing porous membrane polycarbonate (PC) inserts with an 8 µm pore size into a 24‐well plate. Each PC membrane was coated with 50 µl of Matrigel (Corning, Life Sciences Corp., NY, USA) to create a basement membrane‐like barrier. Cells (5 × 10⁴) were then seeded into the upper chamber of the Transwell insert, suspended in 180 µl of FBS‐free culture medium to induce a starvation condition that promotes invasive behavior. The lower chamber was filled with 500 µl of DMEM containing 10% FBS, with no added penicillin or streptomycin. During the assay, PBS‐treated cells served as the control group. In contrast, other experimental groups were treated with free LTZ, LTZ‐NPs, and LTZ‐FNPs, each at their respective IC₅₀ values.

2.11. Statistical Analysis

Statistical analyses were performed using Statistical Package for the Social Sciences (SPSS) Inc. (version 21, Chicago, IL). For analyzing differences among multiple groups, used one‐way analysis of variance (ANOVA) was used. Following the ANOVA, post hoc analyses were conducted with the Bonferroni correction to adjust for multiple comparisons, ensuring that the significance level was set at p ≤ 0.05.^{[ 49 ]}

3. Results

3.1. Physical and Structural Characterizations

The ¹H NMR spectrum of PCL‐co‐PEG‐NH₂ displayed several distinct peaks, corresponding to various chemical groups. Peaks at 2.29 and 1.64 ppm confirmed the presence of methylene protons in the PCL block., while a peak at 3.36 ppm was attributed to methoxy in the PEG block. Additionally, a peak at 1.21 ppm indicated the protons of the terminal amine group. Peaks observed at 6.75, 7.65, and 8.75 ppm were assigned to the folate ring protons, supporting evidence of successful caprolactone ring‐opening polymerization and synthesis of the functionalized block copolymer (Figure 3A,B).

Figure 3.

Physical and structural characterization. ¹H NMR spectra of A) PCL‐co‐PEG‐NH₂ and B) FA‐functionalized PCL‐co‐PEG. C) FTIR of FA, PCL‐co‐PEG, and FA‐functionalized PCL‐co‐PEG. SEM image of D) PCL‐co‐PEG NPs (i.e., NPs), and E) PCL‐co‐PEG‐FA NPs (i.e., FNPs). Contact angles of F) LTZ‐NPs and G) LTZ‐FNPs, conducted by depositing aqueous droplets containing the polymers onto the surfaces. Contact angle measurements were conducted in triplicate and reported as mean ± standard deviation (SD).

FTIR spectroscopy revealed a distinctive carbonyl band at 1722 cm⁻¹, indicating the formation of copolymers, specifically PCL‐co‐PEG. Changes in the FTIR spectrum included an increased intensity of the aliphatic CH stretching band of ε‐caprolactone at 2943 cm⁻¹ and a decreased absorption band associated with the CH stretching vibration in PEG at 2882 cm⁻¹, confirming the formation of copolymers (PCL‐co‐PEG) and successful incorporation of both ε‐caprolactone and PEG into the polymer structure. The absorption bands at 3427 cm⁻¹ were associated with terminal hydroxyl groups in the PCL‐co‐PEG block copolymer. Furthermore, the presence of the peak at 1604.3 cm⁻¹ (aromatic) in the copolymer structure reaffirmed the successful synthesis of FA‐PCL‐co‐PEG, indicating the incorporation of folate (Figure 3C).

Comprehensive size distribution analyses confirmed that all synthesized NPs were within the nanoscale range. The LTZ‐free NPs had an average size of 64.7 ± 1.6 nm, while LTZ‐containing NPs, specifically LTZ‐NPs and LTZ‐FNPs, showed slightly larger hydrodynamic diameters (Table 2 ). Additionally, LTZ‐free NPs displayed a zeta potential of ≈−23.7 ± 0.9 mV, whereas LTZ‐containing NPs, especially those functionalized with FA, showed reduced zeta potential values (Table 2). A statistical analysis using an independent t‐test revealed a significant difference in zeta potential between LTZ‐NPs and LTZ‐FNPs (p‐value < 0.001). This suggests that FA conjugation significantly alters the surface charge of the NPs. Consequently, these findings indicate that LTZ‐functionalized NPs (LTZ‐NPs and LTZ‐FNPs) may have enhanced cellular uptake potential compared to LTZ‐free NPs, due to increased electrostatic attraction to the negatively charged cell membrane.

Table 2.

Characterization by DLS of the hydrodynamic diameter (Z‐average), polydispersity (PDI), and surface charge (zeta potential) of NPs.

Sample	Z‐average ± SD [nm]	PDI ± SD	Zeta potential ± SD [mV]
NPs	64.7 ± 1.6	0.17 ± 0.02	−23.7 ± 0.9
LTZ‐NPs	70.7 ± 1.3	0.34 ± 0.01	−11.9 ± 1.2
LTZ‐FNPs	84.9 ± 1.5	0.24 ± 0.03	−19.9 ± 1.1

Additionally, the morphology of these NPs and FNPs was analyzed using SEM, revealing a predominantly spherical shape (Figure 3D,E). Hydrophilicity emerged as a key characteristic for targeted delivery, potentially influencing the efficiency and effectiveness of the NPs. To assess this, contact angle measurements were conducted, yielding values of 24.2° ± 1.7° and 37.3° ± 2.1° for LTZ‐NPs and LTZ‐FNPs, respectively (Figure 3F,G). These results suggest that FA functionalization did not significantly reduce the hydrophilicity of the NPs, even with LTZ encapsulation, which is beneficial for prolonged circulation time in the bloodstream.

3.2. Docking Outputs

In the molecular docking analysis, our primary objective was to examine the binding interactions between LTZ and both NPs and FNPs. The results showed that LTZ had a low affinity for binding to NPs alone. However, in the presence of FA, the binding affinity between LTZ and FNPs increased significantly, with a binding energy of −15.9 kcal mol⁻¹). This analysis highlighted the crucial role of the triazole ring of LTZ in stabilizing the interaction with FNPs. Furthermore, in the presence of FA, FNPs predominantly engaged in π–π interactions, providing detailed insights into the interaction mode of LTZ with the designed structures. These findings strongly suggest LTZ's inclination toward entrapment within FNPs, accentuating the potential for FNPs to effectively encapsulate and deliver LTZ (Figure 4 ).

Figure 4.

Molecular docking to assess binding mode and molecular interactions. Various types of interactions were evaluated between LTZ and FNPs, providing valuable insights into the molecular mechanisms governing drug encapsulation.

3.3. Encapsulation Efficiency and Drug Loading

The analysis of LTZ encapsulation was conducted through visible spectroscopy, with the results quantified in terms of encapsulation effectiveness (expressed as a percentage). In the case of NPs, the maximum encapsulation effectiveness of LTZ was determined to be 39.59 ± 0.67%. Notably, an even higher encapsulation efficiency of 67.63 ± 1.1% was achieved when utilizing the FNPs. Furthermore, LTZ loading was observed to be 2.1 ± 0.09% in NPs, whereas in FNPs, it reached 7.9 ± 0.81%. These findings underscore the enhanced encapsulation capacity of FNPs compared to NPs, indicating their potential for efficient drug delivery applications.

3.4. Release Profiles

This study examined the release behavior of LTZ from various nanoformulations in both physiological and acidic environments, as depicted in Figure 5 . LTZ solutions served as control samples and were tested for release at pH 7.4 in a solvent (EtOH: PBS = 10:90 v/v). Additionally, the release profiles of LTZ from NPs and FNPs were investigated at pH 5.4 and 7.4, representing cancerous and normal environments, respectively. One‐way analysis revealed significant differences in the release rates between free LTZ and the other groups (p‐value < 0.0001), indicating that the formulation affects drug release. Additionally, statistical analysis showed significant differences in the release profiles of LTZ‐NPs at pH 5.4 and 7.4 (p‐value < 0.006), as well as LTZ‐FNPs at these pH levels (p‐value < 0.0001). Figure 5 illustrates that LTZ solutions displayed an initial burst of release in the solvent during the first 10 h, as the drug rapidly diffused along the concentration gradient. The release pattern shows initial diffusion, followed by a 40‐hour plateau due to drug exhaustion, equilibrium, and matrix degradation. This is typical in release systems when the concentration gradient or degradation limits further release.

Figure 5.

Drug release profiles. Cumulative release of LTZ from LTZ‐NPs and LTZ‐FNPs ([LTZ]: [NPs or FNPs] = 10:90 w/w) and free LTZ in PBS at pH 5.4 and 7.4 at 37 °C. Values indicate the mean ± SD (n = 3).

However, this phenomenon was alleviated when LTZ was encapsulated within the nanoformulations. These nanoformulations displayed prolonged and controlled release of LTZ, with ≈70% of the drug being released over ≈50 h. Therefore, the encapsulated LTZ within NPs and FNPs showed significantly improved release profiles compared to free LTZ, suggesting their potential for controlled and sustained drug delivery applications. Given the importance of the Korsmeyer–Peppas model in predicting release profile behavior, this study specifically examined its applicability. The results demonstrate that the Korsmeyer–Peppas model exhibits excellent curve fitting, with high R‐square values for various nanoformulations (Table 3 ). As per the findings, the model's n values range from 0.65 to 0.71 for LTZ‐FNPs, suggesting non‐Fickian or anomalous transport behavior. This result correlates with changes observed in the NP surface and shape transition from spherical, initiated by the addition of FA and modification of the polymeric chain.

Table 3.

The release rate constants (k), coefficients of determination (R ²), and mean ± SD (n = 3) calculated by fitting the release profiles to the Korsmeyer–Peppas models.

Samples		R 2	K	N
LTZ‐NPs	(pH 5.4) (pH 7.4)	0.9812 ± 0.0038	0.1615 ± 0.004	0.5003 ± 0.0660
		0.9867 ± 0.0049	0.2773 ± 0.003	0.5011 ± 0.0540
LTZ‐FNPs	(pH 5.4) (pH 7.4)	0.9742 ± 0.0109	0.4461 ± 0.001	0.6510 ± 0.0320
		0.9833 ± 0.0089	0.3823 ± 0.011	0.7110 ± 0.0660

3.5. Cytotoxicity Assessment

In this study, the cytotoxic effects of LTZ‐NPs and LTZ‐FNPs were evaluated on MCF‐7 cells, a hormone receptor‐positive human breast cancer cell line (Figure 6 ). Both LTZ‐NPs and LTZ‐FNPs were tested at different LTZ concentrations ranging from 30 to 210 nM. Interestingly, LTZ‐FNPs demonstrated superior efficacy compared to LTZ alone, resulting in the highest percentage of cancer cell death (up to 70% at 210 nM). The IC₅₀ values against MCF‐7 cells were determined to be 184.1 ± 2.43 nM, 149.6 ± 2.21 nM, and 121.8 ± 1.95 nM for free LTZ, LTZ‐NPs, and LTZ‐FNPs, respectively. LTZ‐FNPs exhibited increased cytotoxicity toward MCF‐7 cells due to their ability to utilize FR‐mediated endocytosis for targeted delivery, leading to enhanced drug uptake and intracellular LTZ accumulation.

Figure 6.

Evaluation of the cytotoxic effects of LTZ‐NPs and LTZ‐FNPs on breast cancer cells. The cytotoxicity assay examined the dose‐dependent effects of free LTZ, LTZ‐NPs, and LTZ‐FNPs at various concentrations on MCF‐7 cells over 48 h. Control groups included LTZ‐free NPs and LTZ‐free FNPs. Values indicate the mean ± SD and statistical analysis was performed using one‐way ANOVA (n = 3). p* < 0.05, p** < 0.01, and p*** < 0.001.

3.6. Gene Expression Analysis

The inhibitory properties of different formulations influenced the expression levels of various genes within the breast cancer cells. Real‐time PCR analysis was employed to assess the transcriptional expression of Bcl2, Casp8, MMP‐9, and p53, in treated cancer cells (Figure 7 ). A comparison between gene expressions in MCF‐7 cells and those treated with LTZ‐NPs and LTZ‐FNPs revealed a significant increase in the expression levels of pro‐apoptotic and tumor suppressor genes such as Casp8 and p53. Additionally, there was a notable decrease in the expression of the anti‐apoptotic gene Bcl2 after 48 h of exposure to LTZ‐NPs and LTZ‐FNP formulations. Moreover, the effects of LTZ‐NPs on the expression levels of Casp8, Bcl2, and p53 were less pronounced compared to LTZ‐FNPs. Furthermore, the engineered carriers demonstrated decreased MMP‐9 expression levels compared to control cells, with LTZ‐FNPs showing the most significant reduction.

Figure 7.

Gene expression analysis. The changes in gene expression levels of Bcl2, Casp8, MMP‐9, and p53 were quantified by qPCR after treatment with free LTZ, LTZ‐NPs, and LTZ‐FNPs over 48 h. Values indicate the mean ± SD and statistical analysis was performed using one‐way ANOVA (n = 3). p* < 0.05, p** < 0.01, and p*** < 0.001. Samples were compared to free LTZ.

3.7. Cell Cycle Analysis

The impact of the NPs on the MCF‐7 cell cycle was evaluated using flow cytometry, with the results illustrated in Figure 8 . The findings revealed that nanoencapsulation of LTZ resulted in a notable shift toward the SubG1 phase in MCF‐7 breast cancer cells, indicating a significant increase in apoptotic cell populations. Notably, LTZ‐FNPs induced a more significant shift, suggesting enhanced therapeutic efficacy of LTZ when delivered in this formulation. Specifically, the distribution of cells in the SubG1 phase was 35.8% for LTZ‐NPs and 42.1% for LTZ‐FNPs, compared to 18.1% for free LTZ. This increase in apoptotic cells confirms the results obtained from the cytotoxicity assay and indicates that the designed DDS effectively inhibited cell proliferation by promoting apoptosis.

Figure 8.

Cell cycle distribution. Flow cytometry analysis showing the distribution of MCF‐7 cells in each cell cycle phase following treatment with free LTZ, LTZ‐NPs, and LTZ‐FNPs at IC₅₀ concentration for 48 h. The phases are defined as follows: SubG1 represents cells with fragmented DNA, typically indicative of apoptosis; G1 is the first growth phase, characterized by high protein synthesis and cell growth; S phase is when DNA replication occurs; and G2 is the second growth phase following DNA replication. Results are presented as means from three independent experiments (n = 3).

3.8. Cell Invasion Assay

The invasion analysis was conducted to assess the effects of free LTZ, LTZ‐NPs, and LTZ‐FNPs on the invasion of MCF‐7 cells (Figure 9 ). For each treatment condition, eight images were captured per insert to determine the average invasion count across all experimental groups. The results showed that free LTZ, LTZ‐NPs, and LTZ‐FNPs each inhibited tumor cell invasion, aligning with our findings from gene expression analysis. Notably, the LTZ‐FNPs group demonstrated the most substantial anti‐invasion effect among the treatments. In the analyzed fields, the average number of invading cells was 190.3 ± 6.41 for the control group (untreated), 167.4 ± 5.32 for free LTZ, 151.6 ± 4.78 for LTZ‐NPs, and 135.6 ± 4.03 for LTZ‐FNPs. These results underscore the superior efficacy of LTZ‐FNPs in significantly reducing MCF‐7 cell invasion, highlighting the promising potential of this formulation as an anticancer therapy for hormone receptor‐positive breast cancer.

Figure 9.

Transwell invasion assay. A) The total number of invading cells per field for each experimental group was assessed after 48 h using an inverted microscope. B–E) Representative images of invaded cells from the inserts. Multiple images were captured from each transwell to quantify the number of cells that migrated from the upper to the lower chamber. Experimental groups: B) Control (CNT), C) Free LTZ, D) LTZ‐NPs, and E) LTZ‐FNPs. Values represent the mean ± SD, and statistical analysis was performed using one‐way ANOVA (n = 3). *p < 0.05, **p < 0.01.

By effectively inhibiting cell invasion, LTZ‐FNPs demonstrate not only their ability to enhance LTZ delivery but also their promise in reducing metastatic potential, a critical factor in improving therapeutic outcomes. This enhanced performance suggests that LTZ‐FNPs could offer a targeted and efficient approach for treating aggressive breast cancer phenotypes, positioning them as a valuable candidate for further development in anticancer therapeutics.

4. Discussion

We successfully designed LTZ‐loaded polymeric NPs using PCL‐co‐PEG and PCL‐co‐PEG‐FA, synthesized through a ring‐opening polymerization method. A single emulsion technique was employed to create the NPs. Structural validation via ¹H NMR and FTIR confirmed the integrity of the synthesized copolymers. Prior research indicates that PCL‐co‐PEG systems typically use PEG with molecular weights ranging from 2 to 5 kDa, achieving an optimal hydrophilic–hydrophobic balance essential for effective drug delivery.^{[ 50} , ⁵¹ , ^{52 ]} SEM characterization confirmed the spherical morphology and uniformity of the NPs, demonstrating their suitability for drug delivery applications. The FNPs also displayed a spherical shape, but they were slightly larger than the FA‐free NPs. This size increase is attributed to the attachment of FA to the NP surface.^{[ 53 ]} It is important to note that, in addition to NP morphology, factors such as polymer composition, drug–polymer interactions, drug loading, environmental conditions, and particle size all play a crucial role in influencing drug release behavior.^{[ 14} , ²⁷ , ^{38 ]} In addition, DLS analysis was conducted to assess the size of the NPs, showing a nano size range of ≈64.7 ± 1.6 nm. Interestingly, the conjugation of FA to NPs increased the size to 84.9 ± 1.5 nm, suggesting that FA conjugation caused the NPs to slightly expand. Additionally, the PDI of NPs (0.17 ± 0.02) indicated high uniformity, reflecting excellent monodispersity, which is ideal for consistent performance. Upon loading LTZ, the PDI increased, suggesting a broader size distribution, likely due to interactions between the hydrophobic drug and the polymer matrix. Interestingly, FA‐functionalized NPs exhibited better uniformity than LTZ‐NPs, probably due to the stabilizing effect of FA. These results suggest that FA conjugation enhances particle homogeneity, improving the potential for reliable and targeted drug delivery. This helps ensure more consistent and targeted therapeutic outcomes while maintaining stability within the acceptable range for biomedical applications (PDI < 0.5). Furthermore, all formulated NPs exhibited a negative zeta potential value, as detailed in Table 2. These negative values can be attributed to the presence of free carboxyl end groups of PCL‐co‐PEG on the surface of the NPs.^{[ 54 ]}

Interestingly, the incorporation of LTZ into the NPs resulted in an increase in the zeta potential (−11.9 ± 1.2 mV). Zeta potential is closely related to electrophoretic mobility, which represents the ratio of particle migration velocity to the potential gradient. When the average size of the NPs increases, the migration velocity of charged particles under an applied electric potential slows down, which leads to a decrease in zeta potential compared to smaller NPs that exhibit higher migration velocity and more negative zeta potential values.^{[ 55 ]} This effect also applies to LTZ‐FNPs. The observed increase in zeta potential can be attributed to the larger average size of the NPs, which is influenced by the incorporation of both LTZ and FA. Furthermore, the investigation into the hydrophilicity of LTZ‐NPs and LTZ‐FNPs revealed that both structures possess a hydrophilic nature.^{[ 56 ]} This characteristic is crucial as it facilitates their efficient movement through the body's circulatory system. The study highlighted the significant hydrophilicity of both LTZ‐NPs and LTZ‐FNPs, reinforcing their suitability for circulation within the body.^{[ 57 ]} This hydrophilic characteristic of the DDS is particularly important for improving the tissue distribution of LTZ, a hydrophobic compound.

In addition to the structural studies, molecular docking simulations were used to investigate the interactions between LTZ and NPs, as well as FNPs. When NPs and LTZ were docked, minimal interaction was observed, indicating no significant binding affinity between them.^{[ 58 ]} In contrast, the interaction between LTZ and FNPs revealed strong Pi‐Pi stacking and hydrogen bond interactions. The presence of FA on the surface of the FNPs enhanced these interactions, suggesting a stronger tendency for LTZ to be encapsulated within the core of the FNPs.^{[ 59 ]} The presence of FA on the surface of the FNPs enhanced these interactions, suggesting a stronger tendency for LTZ to be encapsulated within the core of the FNPs.^{[ 67 ]} This finding emphasizes the potential of FA to improve the binding and entrapment of LTZ within the NPs, highlighting its promising role in enhancing the effectiveness of polymeric DDs.

LTZ‐FNPs exhibited higher encapsulation efficiency, which aligns with the results of the molecular docking analysis. This suggests that hydrophobic drugs like LTZ were effectively integrated into the hydrophobic block copolymer and loaded efficiently through proper conjugation.^{[ 60 ]} Additionally, the lower encapsulation efficiencies observed in smaller particles can be attributed to their larger surface area relative to the volume of the organic phase. The increased surface area facilitates more direct contact between the internal and external phases during the emulsification process, leading to greater diffusion of LTZ into the external medium and subsequent drug loss. In contrast, larger droplets, with a smaller surface area relative to the volume of the organic phase, experience less LTZ loss to the aqueous phase, resulting in higher drug entrapment efficiencies.^{[ 61 ]} Moreover, higher LTZ entrapment allows for a more uniform dispersion of LTZ both near the surface and within the core of the NPs. This leads to enhanced LTZ release from the NPs.

The release profiles revealed that LTZ was released more rapidly from the NPs in acidic environments compared to normal conditions. This accelerated release is due to the increased swelling and disintegration of both NPs and FNPs, as well as the faster hydrolysis of surfactants under acidic conditions. Moreover, under equivalent pH conditions, FNPs exhibited a slower LTZ release rate than NPs. This difference can be attributed to the enhanced stability of the FA‐modified NPs, which led to more controlled and sustained LTZ release. The increased stability of FNPs is likely due to the entanglement of FA molecular chains on the NP surface, which helps protect them from rapid hydrolysis. As a result, FNPs demonstrated a more controlled LTZ release at both acidic (pH 5.4) and physiological (pH 7.4) conditions compared to NPs.^{[ 62 ]} The release rate constant (K) and release exponent (n) provided further insights into the mechanism and kinetics of LTZ release from the NPs. In the Korsmeyer–Peppas model, the release exponent “n” was crucial for describing the kinetics of LTZ release, serving as an important parameter in understanding the release behavior of these nanosystems.^{[ 63 ]}

For LTZ‐NPs, Fickian diffusion (n ≈ 0.5) indicated that the solvent (or the LTZ) diffused through the polymer matrix at a notably faster rate than the relaxation or conformational changes of the polymeric chains. However, for LTZ‐FNPs (0.5 < n < 1), non‐Fickian diffusion or anomalous diffusion was observed, showing non‐linear relationships between the diffusion rate and the concentration gradient.^{[ 64} , ^{65 ]} In LTZ‐FNPs, anomalous transport was driven by the combined effects of non‐Fickian diffusion through the hydrated layers of the matrix and the relaxation of polymer chains. The drug release from the FNP matrix was governed by several mechanisms, including surface and bulk erosion, disintegration, diffusion, and desorption. Initially, drug release primarily occurred through diffusion from the polymer matrix. Over time, the release was influenced not only by the diffusion of LTZ but also by the degradation of the polymer matrix. The formation of acidic monomers facilitated further polymer degradation. Typically, bulk‐eroding polymers are expected to show an initial burst release followed by a controlled release.^{[ 66 ]} However, this study did not observe an initial burst release in any of the formulations, except for free LTZ, suggesting that LTZ was uniformly encapsulated within the NPs. Interestingly, the release mechanism of LTZ from both LTZ‐NPs and LTZ‐FNPs was consistent in both acidic (pH 5.4) and neutral (pH 7.4) environments

The cytotoxicity study demonstrated that LTZ‐FNPs were more effective at eliminating cancer cells compared to both free LTZ and FA‐free LTZ‐NPs, likely due to the overexpression of FRs in cancer cells, which enhances targeted uptake. Additionally, the higher cytotoxicity of LTZ‐NPs compared to free LTZ can be attributed to their slower, sustained drug release, enhanced stability, prolonged effective concentration, and improved drug penetration into cells. Competition studies with free FA, as reported by others, confirmed that the primary uptake mechanism for these FA‐functionalized NPs is FRα‐mediated endocytosis.^{[ 67} , ^{68 ]} The incorporation of FA onto NPs facilitated improved targeted delivery, thereby enhancing the therapeutic efficacy. RT‐qPCR analysis also showed that both LTZ‐NPs and LTZ‐FNPs significantly affected the expression of target genes. Key genes involved in apoptosis and tumor suppression, such as Casp8 and p53, were significantly upregulated, while the expression of the anti‐apoptotic gene Bcl2 decreased. Upregulation of apoptosis‐related genes accelerates apoptosis in cancer cells. Reduced expression of Bcl‐2 promotes apoptosis by facilitating the release of cytochrome c from mitochondria and the activation of caspases.^{[ 69 ]} In addition, the designed DDS significantly reduced the expression of MMP‐9, an enzyme vital for the degradation of the extracellular matrix (ECM). MMP‐9 contributes to the breakdown of collagen and other ECM components, processes that facilitate cell motility and are critical for cancer cell invasion and metastasis. By inhibiting MMP‐9 expression, the DDS not only reduces the invasive behavior of cancer cells but also prevents their spread to distant organs. This decrease in MMP‐9 expression highlights the potential of the DDS to both deliver therapeutic agents effectively and modulate key molecular pathways in cancer progression, offering a promising strategy to limit tumor growth and metastasis.^{[ 47} , ^{70 ]} These engineered DDS, with targeted and controlled drug release mechanisms, could effectively prolong their presence in the environment surrounding cancer cells. This extended exposure enhances the drug's efficacy, leading to improved therapeutic outcomes.^{[ 71 ]} Moreover, these systems may remain in the bloodstream for a longer period before being eliminated from the body. LTZ‐FNPs exhibited notable features, including significant controlled release of LTZ at pH 5.4. This release notably increased the sub‐G1 phase in MCF‐7 breast cancer cells and reduced invasion cells compared to LTZ‐NPs and free LTZ.

While this study presents promising results, several limitations could be addressed in future research. Investigating the molecular weight of the synthesized and FA‐functionalized copolymers may offer valuable insights into particle distribution and help optimize the formulation. Additionally, using confocal laser scanning microscopy and flow cytometry to study NP uptake by cells could provide a deeper understanding into the mechanisms of cellular internalization. Future studies focusing on FR‐mediated endocytosis as the primary uptake pathway will enhance our understanding of the DDs' behavior and its potential for targeted therapy.

5. Conclusion

The objective of this study was to enhance the delivery of LTZ for breast cancer treatment by developing polymeric NPs conjugated FA as a targeted nanocarrier. Our results demonstrated successful encapsulation of LTZ in both NPs and FNPs, leading to sustained drug release and high drug entrapment efficiency. The NPs were synthesized using the emulsion‐solvent evaporation method, resulting in small particle sizes and a hydrophilic surface that contributed to an extended half‐life in the MCF‐7 breast cancer cell line. Moreover, the nanosystems exhibited pH‐dependent controlled drug release, with LTZ‐FNPs showing more controlled release under acidic conditions typical of the tumor microenvironment. In vitro studies further revealed that FA‐functionalized NPs significantly increased cytotoxicity against hormone‐positive breast cancer cells, effectively inducing apoptosis and inhibiting cell invasion.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

Conceptualization was done by N.R. and S.B. Methodology was done by N.R. and H.N. Software was done by N.R. and A.N. Validation was done by S.S. and A.G. Formal analysis was done by N.R. and H.N. Investigation was done by S.S. and A.G. Data curation was done by N.R., A.G., and H.N. N.R., A.G., H.N., and S.A.B. wrote the original draft. A.N., S.S., M.A., H.A., and S.A.B. wrote the review and editing. Visualization was done by A.G. Supervision was done by N.R. and S.A.B. All authors have read and agreed to the published version of the manuscript.

Rostami N., Nikzad A., Shaybani S., Noei H., Ghebleh A., Alidadi M., Abbasi H., Bencherif S. A., Engineering Folic Acid‐Modified Nanoparticles to Enhance Letrozole's Anticancer Action. Macromol. Biosci. 2025, 25, 2400558. 10.1002/mabi.202400558

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.