Polymeric PLGA Nanoparticles Loaded with Acalypha monostachya Leaf Hexane Extract: A Novel Strategy for Antineoplastic Activity

Abstract

Background/Objectives: Acalypha monostachya is used in rural communities in Mexico as a traditional remedy for cancer, and we previously observed cytotoxic activity of its extracts against MDA-MB-231 and HeLa cells. Methods: Because lipophilic plant fractions disperse poorly in water, we encapsulated the hexane leaf extract (LHE) of A. monostachya in poly (lactic-co-glycolic acid) (PLGA) nanoparticles prepared by nanoprecipitation, characterized them physicochemically, and evaluated their in vitro cytotoxicity. Results: The selected extract/polymer ratio (5/50, w/w) produced nanoparticles with a mean diameter of 131.4 ± 0.5 nm and a PDI of 0.122 ± 0.028, with an encapsulation efficiency of 92.03% and a loading of 8.43%. We next evaluated cytotoxicity by MTT after 24 h in HeLa and MDA-MB-231 cells and compared the response with non-tumorigenic HaCaT keratinocytes. Encapsulation increased potency relative to free LHE, yielding IC₅₀ values of 30 µg/mL (HeLa), 60 µg/mL (MDA-MB-231), and 95 µg/mL (HaCaT). These values corresponded to selectivity indices of 3.2 (HaCaT/HeLa) and 1.6 (HaCaT/MDA-MB-231). Conclusions: Overall, encapsulation of LHE in PLGA nanoparticles yields an aqueous PLGA nanoparticle suspension and is associated with improved in vitro potency while maintaining measurable selectivity against cancer cells.

1. Introduction

Cancer morbidity and mortality remain high worldwide. In Latin America and the Caribbean, the highest incidence for both sexes is reported for prostate (14.6%), breast (14.2%), and colorectal cancer (9.4%) [1]. In Mexico, GLOBOCAN 2022 reported 207,154 new cancer cases and 96,210 deaths, with 577,487 prevalent cases [1]. Among newly diagnosed cases, breast cancer was the most frequent malignancy overall (15.0%), followed by prostate (12.8%), colorectal (7.8%), thyroid (5.5%), and cervical cancer (5.0%) [1]. Among women, breast and cervical cancers were the most common (27.9% and 9.3%, respectively) [1]. National data from INEGI also indicate that malignant breast tumors are the leading cancer diagnosis in women aged 30–59, followed by malignant cervical tumors [2]. Many cancers are linked to modifiable risk factors, including obesity, infections, UV radiation, and alcohol consumption.

Surgery, chemotherapy, and radiotherapy remain central to cancer treatment, but outcomes are often limited in advanced or metastatic disease. Adverse effects are common, and newer approaches such as immunotherapy benefit only a subset of patients and may also cause substantial toxicity. Furthermore, these therapies are often associated with significant adverse effects, such as damage to non-tumor cells and tumor recurrence [3]. Together, all these constraints justify exploring adjunct approaches, including plant-derived candidates and improved delivery systems.

Traditional medicine has historically represented a fundamental source for the discovery of antineoplastic agents, as a substantial proportion of currently used chemotherapeutic drugs are derived directly or indirectly from natural products. Systematic analyses indicate that more than 60% of anticancer agents approved in recent decades originate from natural compounds or their semisynthetic derivatives, many of which were initially identified through ethnomedical knowledge and the traditional use of medicinal plants [4,5]. Notable examples include vinca alkaloids derived from Catharanthus roseus, taxanes from the genus Taxus, and camptothecin from Camptotheca acuminata, all of which were originally employed in traditional medical systems and subsequently validated through pharmacological and clinical studies [6,7]. These compounds exert their antitumor effects by modulating key cellular processes, including cell cycle arrest, microtubule disruption, and the induction of apoptosis—mechanisms closely associated with tumor growth suppression [8,9]. In this context, ethnopharmacology and the systematic exploration of biodiversity remain highly relevant strategies for the identification of novel bioactive molecules with antineoplastic potential, particularly in megadiverse regions such as Mexico, where numerous medicinal plants have been traditionally used for the empirical treatment of cancer [10,11]. However, many phytochemical compounds exhibit low polarity, which limits their solubility and bioavailability.

Given these limitations, nanomedicine has emerged as a promising alternative in oncology, enabling the targeted delivery of therapeutic agents, including bioactive compounds of plant origin. Within this field, one of the most studied systems is nanoparticles (NPs) based on the polylactic-co-glycolic acid (PLGA) copolymer, approved by the FDA for medical use. One practical barrier in developing plant-derived candidates is formulation, particularly for lipophilic fractions with poor aqueous dispersibility. PLGA NPs are widely used as drug-delivery systems and have a long track record in biomedical applications. This polymer stands out for its biocompatibility and biodegradability, as it degrades in vivo through hydrolysis of its ester bonds, generating lactate and glycolate as end products [12] and has been used to encapsulate natural compounds such as curcumin to address low aqueous solubility and limited bioavailability [13]. Because plant extracts contain multiple constituents, they remain of interest as sources of anticancer candidates [14].

Previous studies reported by our research group have demonstrated that the hexane extract (HE) of Acalypha monostachya (A. monostachya), a plant from the Euphorbiaceae family traditionally used in Mexico, exhibits selective cytotoxic activity against breast cancer (MDA-MB-231) and cervical cancer (HeLa) tumor cell lines [15]. Given the limited aqueous solubility of non-polar compounds in hexane extracts, this study aimed to encapsulate the HE of A. monostachya in PLGA NPs to improve its solubility, stability, and cytotoxic efficacy. Encapsulation of this active fraction in a nanocarrier may improve its handling in aqueous media and support further biological evaluation. This study first compared hexane extracts from stems, leaves, and inflorescences to identify the most active plant part. We then prepared and characterized PLGA NPs loaded with the selected HE to improve aqueous dispersibility and evaluate selective cytotoxicity in human cancer cell lines.

2. Materials and Methods

2.1. Plant Material and Collection

The aerial parts (stems, leaves, and inflorescences) of A. monostachya were collected in July 2023 from Zona Loma Larga Oriente in San Pedro Garza García, Nuevo León, northern Mexico (25°39′21.9″ N, −100°20′07.3.14″ W). The plant material was authenticated by Dr. Marco A. Guzmán-Lucio in the Facultad de Ciencias Biológicas, UANL. A specimen was deposited at the herbarium of this Faculty with accession number 030641.

2.2. Preparation of Hexane Extracts

Following collection, the plant materials were separated into stems, leaves, and inflorescences, then shade-dried at room temperature (RT, 23 ± 2 °C) for three days. Each dried plant fraction was subsequently ground to a fine powder using a manual grain mill (Victoria, Medellín, Colombia). To obtain the extracts, the powdered material (50 g) from each fraction was macerated in 500 mL of HPLC-grade n-hexane for 24 h at RT with constant stirring. The resulting hexane extract (HE) was filtered and concentrated under reduced pressure using a rotary evaporator (Yamato Scientific Co., Ltd. RE-200, Chūō, Japan). The final extract yield was calculated as:

Yield (% w/w) = (final extract weight/initial plant weight) × 100

(1)

The hexane extracts from stems (SHE), leaves (LHE), and inflorescences (IHE) were stored in amber glass bottles at 4 °C until further use.

2.3. Phytochemical Screening of Extracts

A preliminary phytochemical screening was performed for the qualitative determination of secondary metabolites in the extracts according to Guillén-Meléndez et al. (2021) [15]. For each test, 2 mg of each extract (SHE, LHE, and IHE) was dissolved in 2 mL of methanol. The presence of various phytochemicals was assessed via the following colorimetric and precipitation reactions:

• Unsaturation: Addition of three drops of 2% KMnO₄.
• Carbonyl group: Addition of 1 mL of a saturated solution of 2,4 dinitrophenylhydrazine in 6N HCl.
• Phenolic compounds: Addition of three drops of 2.5% FeCl₃ in water.
• Steroids and terpenes (Salkowski test): Addition of 2 mL of H₂SO₄ to 2 mg of extract dissolved in chloroform.
• Saponins: Addition of three drops of 10% NaHCO₃ solution.
• Alkaloids (Dragendorff test): Addition of three drops of Dragendorff’s reagent. The reagent was prepared by mixing two solutions: Solution A (0.85 g of Bi(NO₃)₃ in 10 mL of CH₃COOH and 40 mL of water) and Solution B (8 g of KI in 20 mL of water). The final reagent consisted of 5 mL of solution A, 4 mL of solution B, and 100 mL of water.
• Coumarins and lactones: Addition of three drops of 10% NaOH.
• Sesquiterpene lactones (Baljet test): Addition of three drops of Baljet’s reagent, consisting of Solution A (1% C₆H₃N₃O₇ in ethanol) and Solution B (10% NaOH).
• Carbohydrates (Molisch test): Addition of 1 mL of Molisch’s reagent and 2 mL of H₂SO₄.
• Aromatic compounds: Addition of three drops of a mixture of 1 mL of concentrated H₂SO₄ with one drop of CH₂O (formaldehyde).

The results were evaluated semi-quantitatively, with the intensity of the reaction corresponding to the presence of the compound: (−) negative or not detected; (+) slightly positive; (++) positive; and (+++) strongly positive [15,16].

2.4. Cell Viability MTT Assay

The cytotoxic effects of the SHE, LHE, and IHE of A. monostachya were assessed using the MTT assay (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) [17] to determine which of the extracts exhibited the greatest cytotoxic effect on both human tumor and non-tumor cell lines. Cell viability and culture conditions were performed according to Guillén-Meléndez et al. (2021) [15].

For cytotoxicity evaluation, we employed two human cancer cell lines (HeLa and MDA-MB-231), and the non-tumor cell line HaCaT was used as a control.

(a) HeLa Cell Line: cells derived from adenocarcinoma of the human cervix (ATCC: CCL-2) from a 31-year-old Black female patient. They are adherent cells with epithelial morphology and are positive for cytokeratin. They have been reported to contain human papillomavirus 18 (HPV-18) sequences [18]; additionally, these cells express low levels of p53 and normal levels of pRB [19]. The cells were incubated in 1× Advanced DMEM supplemented with 4% v/v of inactivated FBS, 1% penicillin/streptomycin, and 1% L-glutamine.

(b) MDA-MB-231 cell line: these cells were derived from human mammary adenocarcinoma (ATCC: CRM-HTB-26) from a 51-year-old Caucasian female patient. They are adherent cells of epithelial morphology and express the WNT7B oncogene. They express epidermal growth factor (EGF) and transforming growth factor-alpha (TGF) [20].

HeLa and MDA-MB-231 cell lines were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA).

(c) Non-tumor control HaCaT cell line: immortalized human keratinocytes. This cell line was obtained from Cytion (Eppelheim, Germany).

All cell lines were cultured in 1X Advanced Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 4% v/v of inactivated fetal bovine serum (FBS), 1% v/v penicillin-streptomycin, and 1% glutamine. Cells were incubated at 37 °C in a humidified atmosphere containing 5% CO₂. Media was refreshed every two days until cells reached 80% confluence, at which point the experiment tests were initiated.

Cells (5 × 10⁴ per well; n = 7) were seeded in 96-well plates (Corning^®, Corning, NY, USA) and incubated for 24 h. Subsequently, cells were treated with increasing concentrations (0, 10, 50, 100, 300, and 500 µg/mL) of SHE, LHE, or IHE for 24 h. Advanced DMEM without any HE was used as a negative control.

After 24 h of treatment, light micrographs were taken 2 h before the end of the incubation to evaluate changes in cell morphology. Then, 15 µL of MTT reagent (3 mg/mL in 1× PBS; Sigma-Aldrich, St. Louis, MO, USA) was added to each well, and plates were incubated for 2 h at 37 °C. The supernatant was then carefully removed, and 150 µL of 2-propanol was added to each well to dissolve the formazan crystals. Absorbance was measured at 590 nm, with a reference filter at 620 nm, using a microplate reader (iMark™, Bio-Rad, Thane, India). The absorbance value is directly proportional to the number of metabolically active cells, which is an indirect measure of cell viability.

Based on this preliminary screening, LHE was selected as the most active extract for encapsulation. The cytotoxic activity of LHE-PLGA NPs was then evaluated using the same MTT protocol described for extract evaluation. Control groups included cells treated with culture medium alone (vehicle control), empty PLGA-NPs, and unencapsulated LHE.

2.5. Gas Chromatography-Mass Spectrometry (GC-MS) Analysis

The A. monostachya LHE was analyzed using a 6890N gas chromatograph coupled to a 5973 INERT mass spectrometer (Agilent Technologies, Santa Clara, CA, USA), equipped with an HP-5MS column (30 m × 0.25 mm, 0.25 µm; Agilent). The injection was performed in splitless mode at 250 °C, with an injection volume of 1 µL. Helium was used as the carrier gas at a flow rate of 0.9 mL/min. Separation was performed starting at a temperature of 70 °C, held for 2 min, then increased to 270 °C at 8 °C/min, and subsequently raised to 290 °C at 10 °C/min. The total analysis time was 34 min. Data acquisition was performed in full scan mode; the MS transfer line temperature was 290 °C, the source temperature was 230 °C, and the quadrupole temperature was 150 °C.

2.6. Ultra-High-Performance Liquid Chromatography-Mass Spectrometry (UHPLC-MS) Analysis

The phytochemical profile of the A. monostachya LHE was investigated by UHPLC-MS using a Thermo Fisher Scientific Ultimate 3000 system fitted with a UV-Vis detector and interfaced to an LCQ Fleet ion-trap mass spectrometer (Thermo Fisher Scientific, Waltham, MA, USA). Chromatographic separation was achieved on a Supelco Discovery HS F5 column (150 × 2.1 mm, 3 µm) (Supelco, Bellefonte, PA, USA) employing a binary mobile phase consisting of aqueous 0.1% formic acid (solvent A) and acetonitrile containing 0.1% formic acid (solvent B). The gradient program was as follows: initial composition 10% B maintained for 1 min, linear increase to 70% B in 19 min, held for 20 min and followed by a 20 min re-equilibration period. The flow rate was set at 0.25 mL/min, and the column oven temperature was kept at 45 °C. UV detection was monitored at 254 nm. Mass spectrometric analysis was conducted in both positive and negative electrospray ionization modes. Key source settings included a capillary temperature of 350 °C, capillary voltage of +15 V (positive mode) or −22 V (negative mode), sheath gas flow of 60 arbitrary units, and auxiliary gas flow of 20 arbitrary units. Spectra were recorded in full-scan mode within the m/z range 50–2000, and MS/MS experiments were performed at a normalized collision energy of 35%. The LHE was prepared at a concentration of 0.2 mg/mL in methanol, filtered through a 0.22 µm membrane, and an injection volume of 10 µL was used.

2.7. Hemolysis Assay of LHE

The hemolytic activity of the unencapsulated LHE was evaluated to assess its hemocompatibility, following a previously described method [21,22]. Human whole blood was obtained from one healthy adult volunteer after written informed consent prior to sample collection. Samples were used exclusively for in vitro hemocompatibility testing and were processed in a de-identified manner. Erythrocytes were isolated by centrifugation, washed four times with phosphate-buffered saline (PBS, 10 mM, pH 7.4), and resuspended in PBS 1× to a final concentration of 5% (v/v).

Aliquots of the erythrocyte suspension were incubated for 30 min at 37 °C in the dark with different concentrations of LHE (0, 10, 50, 100, 300, and 500 μg/mL). Samples treated with PBS and sterile distilled water served as negative (0% hemolysis) and positive (100% hemolysis) controls, respectively. After incubation, the tubes were centrifuged at 9085× g for 3 min at 4 °C, and 200 µL of the supernatant was transferred to a 96-well plate.

The optical density (OD) of the released hemoglobin was measured at 540 nm using a microplate reader (BioTek™ EPOCH™, Winooski, VT, USA). The percentage of hemolysis for each sample was calculated using the following formula:

% Hemolysis = [(OD₅₄₀ Treatment − OD₅₄₀ Negative control)/(OD₅₄₀ Positive control − OD₅₄₀ Negative control)] × 100

(2)

2.8. Synthesis of LHE-PLGA NPs

LHE-loaded PLGA NPs (LHE-PLGA NPs) were prepared using the nanoprecipitation method as described by Elizondo-Luevano et al. (2023) [21]. The organic phase was composed of 50 mg of acid-terminated PLGA (Resomer^® RG 503 H, lactide:glycolide 50:50, molecular weight 24,000–38,000; Sigma-Aldrich^®, St. Louis, MO, USA) and 5 mg of LHE dissolved in acetone, and sonicated for 1 min to ensure complete solubilization (Ultrasonic Branson 2510MT, Merck KGaA, Darmstadt, Germany). This solution was then added dropwise to 12 mL of distilled water (aqueous phase) under continuous magnetic stirring. The solvent was removed by evaporation under reduced pressure at RT, with agitation at 90 rpm. The nanoparticle suspension was subsequently stored at 4 °C until further use.

2.9. Characterization of NPs

2.9.1. Raman Spectroscopy

Raman spectra were acquired for empty PLGA NPs, unencapsulated LHE of A. monostachya, and LHE-PLGA NPs to assess the chemical composition of the samples. Spectra were recorded using a Cora 5500 Raman spectrometer (Anton Paar, Graz, Austria) with a 785 nm excitation laser operating at 300 mW. A deep-cooled CCD detector was used to improve signal sensitivity. Prior to measurement, all samples were dispersed to ensure optimal laser–sample interaction.

2.9.2. Fourier-Transform Infrared (FT-IR) Spectroscopy

FT-IR spectra were collected at RT using a PerkinElmer Frontier spectrophotometer (PerkinElmer, Shelton, CT, USA) equipped with a UATR module featuring a diamond/KRS-5 crystal. A spectral resolution of 8 cm⁻¹ was used for all measurements.

2.9.3. Dynamic Light Scattering (DLS)

As part of the characterization of the NPs, the particle size and polydispersity index (PDI) were determined to evaluate the uniformity of particle size distribution within the sample. PDI is a key parameter in nanoparticle characterization, as particle size distribution significantly affects cellular permeability, biodistribution, and retention at the target site [23]. Zeta potential was measured to assess the surface charge density. NPs exhibiting a ζ potential between −10 and +10 mV were classified as ‘neutral,’ whereas values outside the ±30 mV range were considered strongly cationic or anionic [24].

Nanoparticle size distribution and PDI were determined by DLS using a Zetasizer Nano ZS (Malvern Panalytical, Worcestershire, UK) with a 633 nm laser and 173° backscatter detection. Samples were dispersed in distilled water prior to analysis. The refractive index (RI), viscosity, and absorption parameters were set to 1.460, 0.887 cP, and 0.1, respectively, at 25 °C. Five measurements were performed per sample, and mean values were reported.

2.9.4. Encapsulation Efficiency (EE%)

To determine the amount of extract successfully encapsulated within the NPs, LHE-PLGA formulations were centrifuged at 25,000 rpm for 4 h at 4 °C. The supernatants containing unencapsulated (free) LHE were collected and quantified using a GENESYS 10S UV–Vis spectrophotometer (Thermo Scientific, Waltham, MA, USA). A calibration curve was generated using known concentrations of LHE to enable accurate quantification.

The amount of encapsulated extract (EXE) was calculated by subtracting the amount of free extract in the supernatant from the initial amount used in the formulation (EXI). The encapsulation efficiency (EE %) and drug load (%L) were calculated based on Azzazy et al. (2022) [25], with modifications, as follows:

EE (%) = (EXE/EXI) × 100

(3)

where EXE corresponds to the mass of the encapsulated extract and EXI corresponds to the mass of the initial extract,

% L = (ME/MN) × 100

(4)

where ME corresponds to the mass of the nanoencapsulated extract and MN corresponds to the total mass of the NPs.

2.9.5. Scanning Electron Microscopy (SEM)

The morphology and particle size of the NPs were evaluated by scanning electron microscopy (SEM) using a Zeiss EVO MA25 microscope (Zeiss, Oberkochen, German). Samples were diluted, sonicated, and deposited onto aluminum stubs. Micrographs were acquired at an accelerating voltage of 20 kV and a magnification of 20,000×.

2.9.6. Temperature and Stability Tests

NP stability was assessed over 3 months at two storage temperatures: 25 °C and 4 °C. At monthly intervals, samples were analyzed via DLS to evaluate changes in size, PDI, and zeta potential in order to determine the optimal storage conditions.

2.10. Preparation of Working Solutions

Using the LHE-PLGA NPs of A. monostachya, 10 mg/mL stock solutions were prepared in 1.5 mL Eppendorf tubes by dissolving this amount of extract in 50 µL of 100% DMSO (dimethyl sulfoxide) and 950 µL of culture medium with a final DMSO concentration of 5%. Subsequently, 1 mg/mL working solutions were prepared by diluting the stock solution 1:10 to a DMSO concentration of 0.5%. From these working solutions, dilutions were then prepared at concentrations of 0, 10, 50, 100, 300, and 500 µg/mL.

2.11. Morphological Analysis

To evaluate morphological changes in cells exposed to A. monostachya extracts, 1 × 10⁶ cells were seeded in 60 mm dishes and incubated (95% humidity; 5% CO₂) for 24 h to allow adherence. Cells were then treated with 0, 10, 50, and 100 µg/mL of empty PLGA NPs, LHE-PLGA NPs, or unencapsulated LHE, prepared in Dulbecco’s Modified Eagle Medium (DMEM, Gibco™, Grand Island, NY, USA) supplemented with 10% fetal bovine serum (FBS), and incubated for 24 h. Representative photomicrographs were acquired at 40× using an inverted microscope (Southern Precision Instrument, Navi Mumbai, India) 2 h before the end of the incubation period to document treatment-associated morphological changes.

The selectivity index (SI) was calculated as follows (5) [26]:

SI = IC₅₀ Non tumor cells/IC₅₀ Tumor cells

(5)

The IC₅₀ values were calculated for each treatment. Extracts were classified based on their cytotoxicity as highly cytotoxic (IC₅₀ < 10 μg/mL), moderately cytotoxic (IC₅₀ > 10 and <50 μg/mL), cytotoxic (IC₅₀ > 50 and <100 μg/mL), or potentially noncytotoxic (IC₅₀ > 100 and <500 μg/mL).

2.12. Cell Death Analysis

For the assessment of cell death, the Apoptosis/Necrosis Detection Kit (AB176749, Abcam, Burlingame, CA, USA) was used. In this assay, green fluorescence corresponds to the binding of Apopxin Green, an indicator of apoptosis through the detection of phosphatidylserine externalization, while red fluorescence corresponds to 7-AAD, a marker of necrosis, associated with the loss of plasma membrane integrity. These images allow for a visual comparison of the cellular response under the different experimental conditions evaluated. To validate our results, we employed doxorubicin 1 ug/mL as a positive control for cell death; culture medium was employed as the negative control.

2.13. Statistical Analyses

Descriptive statistics were applied to summarize quantitative variables as mean ± standard deviation (SD). To assess the distribution of the data, the Kolmogorov–Smirnov test for normality was performed, confirming a normal distribution for all variables. An ANOVA test was then performed to evaluate the presence of differences between treatments. Subsequently, one-way analysis of variance (ANOVA) was used to evaluate statistically significant differences between treatment groups.

3. Results

3.1. Partial Characterization (Partial Phytochemical Screening)

The phytochemical profiling of the SHE, LHE, and IHE of A. monostachya was compared with a negative control consisting of the solvent used in each test, and subsequently, the corresponding reagents were added without the presence of any crude extract. Our results showed the presence of unsaturations, phenols, steroids, terpenoids, carbohydrates, coumarins, lactones, sesquiterpene lactones, flavonoids, and saponins. Then, through a semi-quantitative analysis based on colorimetric and precipitation reactions, the presence of different groups of secondary metabolites was determined. The results were represented semi-quantitatively with crosses according to the intensity observed in each reaction (

Table 1. Partial phytochemical screening of the HE of A. monostachya.

Phytoconstituents	Test	Observations	Results
			SHE	LHE	IHE
Unsaturations	Potassium permanganate test	Brown precipitate	+	++	+
Phenols	Ferric chloride test	Green colour	+	+++	+
Steroids and Terpenoids	Test of Salkowski	Formation of a Reddish-brown ring	+	+++	++
Coumarins and lactones	Sodium hydroxide test	Yellow colour	-	++	-
Sesquiterpene lactones	Test of Baljet	Orange colour	-	++	+
Flavonoids	Sulphuric acid test	Reddish colour	+	+++	+
Alkaloids	Test of Dragendorff	Orange precipitate	-	-	-
Saponins	Foam test	Presence of stable foam	+	+	+++
Carbohydrates	Test of Molisch	Formation of a purple ring	+	++	+
Aromatic compounds	Formaline test	Red colour	-	-	-
Carbonyl group	2-4 dinitrophenylhydrazine test	Orange colour	-	-	-

Abbreviations: + Slightly positive reaction; ++ positive reaction; and +++ strong positive reaction; -negative reaction. The first columns indicate the name of the secondary metabolite and the used test to perform the detection. The column of observations shows the expected color for each reaction, followed by the results obtained in the hexane extracts (HE) from stems (SHE), leaves (LHE), and flowers (IHE) of A. monostachya.

). These data provide a foundation for future identification and quantification of secondary metabolites in this plant.

3.2. The HE of A. monostachya Leaves Have a Greater Cytotoxic Effect on Human Cancer Cell Lines

First, we evaluated whether the HE from different parts of the A. monostachya plant had the same selective cytotoxic effect previously demonstrated by our study group against MDA-MB-231 and HeLa cancer cell lines [15]. We evaluated the cytotoxic effect of SHE, LHE, and IHE against these human cancer cell lines compared with the nontumorigenic cell line HaCaT.

The results obtained from the MTT assay indicated that LHE from A. monostachya exhibited the highest cytotoxic activity against the HeLa and MDA-MB-231 cancer cell lines compared with the immortalized HaCaT cell line, suggesting a potential selective effect on cancerous cells (Figure 1). In further experiments, we employed the LHE from A. monostachya.

3.3. Chromatographic Analysis of the LHE of A. monostachya

The GC-MS analysis of the A. monostachya LHE revealed the presence of several lipophilic compounds characteristic of non-polar fractions.

Table 2. Volatile and semi-volatile compounds identified by GC-MS in the LHE of A. monostachya [27].

Identified Compound	Formula	MW	Ions (m/z)
Octadecanal	C18H36O	268.5	43 (bp), 41, 57
Palmitic acid methyl ester	C17H34O2	270.5	87 (bp), 143, 83, 227, 101
Linoleic acid methyl ester	C19H34O2	294.5	81 (bp), 95, 82, 96, 79
Linolenic acid methyl ester	C19H32O2	292.5	79 (bp) 95, 80, 55, 108
Phytol	C20H40O	296.5	143 (bp), 73, 75, 81, 123
Eicos-9-ene-1,20-diacetate	C22H44O4	396.6	43 (bp), 55, 82
Vitamin E (Tocopherol)	C29H50O2	430.71	237 (bp), 236, 238, 277, 208

MW: Molecular weight, bp: base peak, m/z: mass-to-charge ratio.

summarizes the obtained data. The major identified constituents included fatty acid methyl esters such as methyl palmitate (methyl hexadecanoate), methyl linoleate (methyl octadeca-9,12-dienoate), and methyl linolenate (methyl octadeca-9,12,15-trienoate), which are common derivatives arising from transesterification during methanolic extraction or natural occurrence. Additionally, octadecanal (a long-chain aldehyde), phytol (a diterpene alcohol derived from chlorophyll degradation), eicos-9-ene-1,20-diacetate (a long-chain diacetate alkene), and vitamin E (α-tocopherol) were detected. These findings are consistent with the lipophilic nature of the extract and highlight the presence of bioactive components with known antioxidant properties, particularly vitamin E and unsaturated fatty acid esters [27].

3.4. UHPLC-MS Analysis of the LHE of A. monostachya

The UHPLC-MS/MS analysis of the LHE of A. monostachya revealed a diverse array of phenolic compounds, predominantly flavonoids and phenolic acids, consistent with the known phytochemical profile of the Euphorbiaceae family. Figure 2 shows representative UHPLC-UV-MS chromatograms of the LHE of A. monostachya, while the tentatively identified metabolites are detailed in

Table 3. UHPLC-ESI-MS/MS tentative identification of metabolites from LHE of A. monostachya [28,29,30,31,32,33,34].

Retention Time (min)	[M+H]+ (m/z)	Fragment Ions (m/z)	[M-H]− (m/z)	Fragment Ions (m/z)	Identified Compound
12.80	287	275, 269, 241, 127			Kaempferol derivative
15.70			477	409, 316, 179	Methyl ellagic glucoside
17.40			483	271	Digalloylglucose
17.84			367	179	3,5-di-O-caffeoylquinic acid
19.58			285	229, 257, 213, 241, 201,225	Kaempferol
19.66	303	275,247,234, 229,191, 175			Ellagic acid
22.17	287	269, 259, 241, 227, 222, 191, 143			Kaempferol derivative
22.60	331	316, 313, 303, 298, 285, 270, 242, 215			3,3-di-O-methyl ellagic acid
23.20			483	465, 331, 313, 271, 211, 169	digalloyl-hexose
23.81	273	255, 247, 203, 175, 153			Naringenin
24.50	291	235, 217, 219, 273			Prenyl-flavonoid
24.7			537	303, 449, 397, 277, 399, 337	Quercetin derivative
25.20			457	457, 915, 914, 458, 388, 442, 373, 427, 384, 233,	(-)-Epigallocatechin gallate
26.67	273	255, 217, 175			Flavanone
28.99			515	191, 179	1,5-di-O-caffeoylquinic acid

[M+H]⁺: protonated quasi-molecular ion, [M-H]⁻: deprotonated quasi-molecular ion, m/z: mass-to-charge ratio.

. As can be observed, key identified metabolites included kaempferol and its derivatives, naringenin and other flavanones, a quercetin derivative, and a prenyl-flavonoid, highlighting the richness in flavonols and flavanones with recognized antioxidant, anti-inflammatory, and potential cytotoxic properties. Additionally, several hydroxycinnamic acid derivatives were detected, such as 3,5-di-O-caffeoylquinic acid and 1,5-di-O-caffeoylquinic acid, which are known for their strong radical-scavenging activity. Ellagic acid-related compounds, including ellagic acid itself, methyl ellagic glucoside, and 3,3′-di-O-methyl ellagic acid, along with galloyl esters like digalloylglucose, digalloyl-hexose, and (-)-epigallocatechin gallate, further underscore the presence of hydrolyzable tannins and ellagitannin precursors, compounds associated with antimicrobial and antiproliferative effects [28,29,30,31,32,33,34]. Overall, the identified metabolites support the medicinal use of A. monostachya and suggest significant potential for antioxidant and anti-inflammatory activities [15], warranting further bioassay-guided isolation and pharmacological evaluation.

3.5. The Unencapsulated LHE of A. monostachya Shows Dose-Dependent Hemolytic Activity

The hemolytic activity of the LHE of A. monostachya was evaluated by incubating it with human erythrocytes at different concentrations (0–500 µg/mL) to assess its hemocompatibility and suitability for pharmaceutical applications.

As shown in Figure 3, LHE exhibited a concentration-dependent hemolytic response. Hemolysis remained below 10% at concentrations up to 50 µg/mL. While the ASTM F756 standard classifies materials as non-hemolytic only below 2%, this threshold is primarily applied to implantable medical devices. In the context of plant extracts and nanoparticle formulations, several studies consider hemolysis values under 10% to reflect acceptable hemocompatibility [35,36]. At 100 µg/mL, a moderate but statistically significant increase (~15%, p < 0.05) was observed.

At higher concentrations, a pronounced increase in hemolysis was detected, reaching approximately 50% at 300 µg/mL and exceeding 80% at 500 µg/mL. Statistical analysis showed highly significant differences between the highest concentrations and the negative control (p ≤ 0.05 to p ≤ 0.0001), confirming the hemolytic toxicity of the extract at high doses. According to commonly accepted hemocompatibility criteria, materials inducing less than 10% hemolysis are considered non-hemolytic, whereas values above this threshold indicate increasing membrane damage and reduced blood compatibility [35,37]. Therefore, the present results indicate that LHE is hemocompatible at low concentrations but induces substantial erythrocyte membrane disruption at higher doses. Overall, these findings define a clear safety window for LHE and emphasize the importance of dose optimization when considering its incorporation into pharmaceutical formulations or nanocarrier-based delivery systems [38,39].

3.6. Generation of NPs with LHE of A. monostachya Using Different Formulations of PLGA Polymer

Table 4. Physicochemical characterization of different formulations of PLGA-NPs loaded with the LHE of A. monostachya (LHE-PLGA-NPs) and unloaded NPs (Empty-PLGA-NPs).

Ratio w/w		Particle Size (nm)	PDI	EE%	%L
5/50	Empty PLGA-NPs	103.1 ± 1.504	0.138 ± 0.016	92.03%	8.43%
	LHE-PLGA-NPs	131.4 ± 0.5033	0.122 ± 0.028
10/60	Empty PLGA-NPs	101.0 ± 0.750	0.112 ± 0.035	95.02%	13.33%
	LHE-PLGA-NPs	143.5 ± 0.611	0.134 ± 0.010
18/60	Empty PLGA-NPs	101.0 ± 0.750	0.112 ± 0.035	92.39%	21.37%
	LHE-PLGA-NPs	152.0 ± 1.474	0.152 ± 0.029
21/60	Empty PLGA-NPs	101.0 ± 0.750	0.112 ± 0.035	92.70%	24.49%
	LHE-PLGA-NPs	181.5 ± 2.335	0.277 ± 0.029

The ratio is indicated as extract/polymer (mg/mg). Particle size (nm), polydispersity index (PDI), encapsulation efficiency (EE%), and percentage extract loading (%L) were evaluated. Values correspond to the mean ± standard deviation (n = 3).

shows the physicochemical parameters evaluated for different formulations of PLGA NPs loaded with the LHE of A. monostachya, varying the extract/polymer ratio. Formulations with higher extract loadings (18/60, 21/60) showed a considerable increase in particle size (greater than 150 nm) and polydispersity index (PDI > 0.15), suggesting a heterogeneous distribution and potential colloidal instability. Furthermore, the encapsulation efficiency (EE%) decreased as the extract amount increased, likely due to supersaturation of the polymer matrix and the expulsion of some of the contents during the NP formation process.

In contrast, formulation 5/50 presented the best overall profile, with an average particle size of 131.4 ± 0.5033 nm, a PDI of 0.122 ± 0.028 (indicative of a monodisperse population), and an encapsulation efficiency (EE) of 92.03%, along with an acceptable loading percentage (8.43%). This profile is consistent with NPs considered optimal for controlled-release systems, as a size smaller than 200 nm favors cellular permeability, prolonged circulation, and prevents uptake by the reticuloendothelial system [40,41].

Furthermore, a PDI of less than 0.2 indicates a uniform particle size distribution, which is crucial for ensuring predictable pharmacokinetic behavior [42]. The high encapsulation efficiency observed in this formulation indicates good compatibility between the lipophilic extract and the PLGA matrix [43]. Therefore, the 5/50 formulation was selected as the most suitable for subsequent biological assays due to its balance of loading efficiency, colloidal stability, and optimal size.

3.7. The 5/50 Formulation Shows the Best Cytotoxic Effect Against Tumor Cells

In addition to quantifying viable cells by establishing the non-hemolytic effect of the generated NPs with different formulations of encapsulated LHE (compared with empty PLGA NPs), we performed an MTT assay to correlate mitochondrial activity with cell viability and evaluate energy metabolism in response to the extracts. Viable cells were able to metabolize the tetrazolium salt, reducing it to formazan, a purple-blue compound, whose absorbance values provided the following results. After 24 h, a cytotoxic effect was observed. In HeLa cells, significant reductions in viability were detected at values ≥ 50 µg/mL with the 5/50 formulation, and similar results were observed in MDA-MB-231 cells. In contrast, HaCaT cells began to show cytotoxic effects only at concentrations ≥ 100 µg/mL (Figure 4).

3.8. Raman Spectra of A. monostachya LHE-PLGA NPs

In our study, Raman spectroscopy was employed to investigate the molecular characteristics of empty PLGA polymer (Resomer^® RG 503 H) NPs, the hydroalcoholic extract of A. monostachya (LHE), and the LHE-loaded PLGA NPs to confirm the incorporation of the extract into the polymer matrix (Figure 5).

The Raman spectrum of LHE exhibited multiple intense and well-defined bands in the 200–1500 cm⁻¹ region, reflecting the chemical complexity of the extract and the presence of various vibrational modes associated with phenolic and aromatic compounds. In particular, bands in the 700–900 cm⁻¹ region are commonly attributed to aromatic ring breathing modes and C–H bending vibrations of aromatic structures present in secondary metabolites, which are characteristic of polyphenolic, flavonoids or terpenoids constituents [44,45,46].

The Raman spectrum of the loaded LHE–PLGA NPs displayed several characteristic bands of LHE, although with reduced intensity compared to free LHE, together with PLGA-related signals. The attenuation and partial overlap of LHE bands in the nanoparticle spectrum suggest successful encapsulation of the extract within the PLGA matrix. Importantly, no new Raman bands or significant peak shifts were observed after encapsulation, indicating the absence of strong chemical interactions or structural modifications. This behavior suggests that LHE is physically entrapped within the PLGA NPs, preserving its chemical integrity [47].

On the other hand, the empty PLGA NPs spectrum shows the typical bands of the copolymer, particularly at 875 cm⁻¹ (C–COO stretching), 1450 cm⁻¹ (CH₂/CH₃ deformation), and 1760 cm⁻¹ (C=O stretching), which are also observed in the loaded NPs, indicating the conservation of the polymer structure after loading [48]. These results confirm the effective incorporation of LHE into the PLGA NPs.

3.9. FT-IR Spectra of A. monostachya LHE-PLGA NPs

Figure 6 shows the infrared spectra obtained by FT-IR for the empty PLGA NPs, the LHE of A. monostachya, and the LHE-PLGA NPs, intended to identify chemical interactions and confirm the incorporation of the extract into the polymeric system. The PLGA spectrum exhibits characteristic bands around 1730 cm⁻¹ (C=O stretching of the ester groups) and ~1240 cm⁻¹ (C–O vibrations of the ester backbone), in agreement with what has been reported for lactic and glycolic acid copolymers [31,34]. For its part, the LHE shows multiple bands, with a marked absorption in the ~1750 cm⁻¹ region, attributed to aromatic C=C vibrations or stretching of conjugated carbonyls present in phenolic compounds and flavonoids [44,49]. In the LHE-PLGA sample, combined signals from both components are observed, where the carbonyl band of the polymer persists at ~1730 cm⁻¹ and an absorption is identified around ~1750, indicating the possible incorporation of the extract. There are also other signals of the extract at approximately 1330, 1200, and 108 cm⁻¹, associated mainly with secondary metabolites with C=C, C–O, and C–N functional groups. The slight decrease in the intensity of the LHE bands suggests encapsulation, which could reduce the free vibration of its functional groups when confined within the polymer matrix [50]. These results complement the previous Raman analysis and support the effective loading of the extract into the PLGA NPs.

The FTIR spectrum of empty PLGA NPs exhibited characteristic absorption bands at approximately 1750 cm⁻¹, corresponding to the ester carbonyl (C=O) stretching vibration, and bands in the range of 1080–1180 cm⁻¹ attributed to C–O–C stretching of the polymer backbone. The LHE spectrum displayed a broad absorption band around 3200–3500 cm⁻¹, associated with O–H stretching vibrations, as well as bands near 1600 cm⁻¹ corresponding to aromatic C=C stretching. In the spectrum of LHE–PLGA NPs, the main characteristic bands of both PLGA and LHE were present, with slight shifts and changes in band intensity. These observations indicate the successful encapsulation of LHE within the PLGA NPs and suggest that no significant chemical interaction or structural modification of the PLGA polymer occurred during nanoparticle formulation.

3.10. Morphological Assessment of A. monostachya LHE-PLGA-NPs

Morphological characterization by SEM allowed the evaluation of the surface structure of the PLGA NPs obtained by nanoprecipitation, both unloaded and loaded with LHE of A. monostachya (LHE-PLGA-NPs). Figure 7a, corresponding to the empty PLGA-NPs, shows a well-defined spherical morphology with a homogeneous distribution and uniform size, typical characteristics of the nanoprecipitation method, which favors the formation of smooth, small particles due to the rapid diffusion of the organic solvent into the aqueous phase [51].

Regarding the LHE-PLGA-NPs, Figure 7b shows the spherical morphology, although slightly rougher surfaces and areas with higher material density are evident, possibly associated with surface deposition of the extract or a redistribution of the polymer during the loading process. These changes are consistent with previous reports in which the inclusion of lipophilic compounds alters the surface tension of the system during nanoparticle formation by nanoprecipitation, generating slight irregularities without compromising structural integrity [52].

This ultrastructural analysis supports the results obtained using DLS, where a slight increase in size and dispersion was observed after extract loading. Furthermore, maintaining the spherical shape is key to ensuring adequate colloidal stability and sustained release, since particles with a high surface-to-volume ratio, such as spheres, allow for better interaction with the biological medium [40].

All these results confirm that the nanoprecipitation process was effective in generating NPs with good morphological definition and that the incorporation of the extract does not significantly alter their structure, validating this technique as a useful strategy for encapsulating lipophilic natural compounds with therapeutic potential.

3.11. Temperature and Stability Tests of Generated NPs

The stability of PLGA NP formulations, both empty and loaded with LHE of A. monostachya, was evaluated over three months at two storage temperatures (4 °C and 25 °C), considering particle size, PDI, and zeta potential as critical parameters.

As shown in Figure 8, NPs stored at 4 °C exhibited greater stability in all evaluated parameters. Regarding particle size, both the empty PLGA NPs and the LHE-PLGA NPs maintained values close to 150–180 nm until month 3. In contrast, the empty PLGA NPs stored at 25 °C exhibited a progressive increase in size, reaching approximately 350 nm by month 3, suggesting particle aggregation or fusion at RT. This phenomenon was accompanied by an increase in the PDI (>0.2), reflecting a loss of homogeneity and monodisperse distribution, especially in the extract-free formulations.

The LHE-PLGA NPs, at both 4 °C and 25 °C, showed better stability than the unloaded NPs at RT, which could be due to a stabilizing interaction between the extract compounds and the polymer matrix, as has been reported for other phytocomposites [53]. However, we observed that even these formulations showed a slight tendency for the PDI to increase over time if not kept refrigerated.

Regarding the zeta potential, all formulations remained between −18 and −25 mV, with no statistically significant changes throughout the study period. This negative value is within the range reported for NPs with good colloidal stability, due to the electrostatic repulsion between particles [54].

In summary, these results indicate that storage at 4 °C maintains the physicochemical integrity of the NPs for up to three months, which is the recommended condition for maintaining the functionality of the developed systems.

3.12. LHE-PLGA NPs Induce Morphological Changes in Cultured Human Cancer Cells

To determine the cytotoxic effect of the LHE-PLGA NPs on HeLa and MDA-MB-231 cancer cell lines or HaCaT, a nontumorigenic cell line, light microscopy observations were performed after treatments at concentrations of 0, 50, 100, 300, and 500 μg/mL. Figure 9, Figure 10 and Figure 11 show bright-field micrographs of HeLa, MDA-MB-231, and HaCaT cell lines, respectively, after exposure to concentrations of 10, 50, and 100 μg/mL for 24 h.

Concentration-dependent changes were observed upon exposure to the LHE-PLGA NPs, mainly in cancer cell lines. HeLa cells were the most affected compared with MDA-MB-231 cells. These changes were accentuated in LHE-PLGA NP-treated cells, suggesting a cytotoxic and antiproliferative effect, followed by those treated with unencapsulated LHE and finally empty PLGA NPs.

Morphological changes were also observed, such as cellular contraction, acquiring a rounded shape caused by the loss of adhesion. These results were compared with those of the control cells, which were treated with vehicle; no such changes occurred in the control cells. In nontumorigenic HaCaT cells, a slight decrease in confluency was observed at a concentration of 100 μg/mL of LHE-PLGA NPs after 24 h.

3.13. Encapsulated LHE of A. monostachya in PLGA NPs Decreases Viability in Human Cancer Cell Lines

MTT analysis correlated with the morphological changes observed in the three evaluated cell lines. HeLa cells were the most sensitive to the LHE-PLGA NP treatment from 10 µg/mL onwards. These results demonstrated a selective concentration-dependent cytotoxic effect. In contrast, MDA-MB-231 cells showed resistance and, in some cases, higher relative viability than the HaCaT cell line; however, it decreased at 100 µg/mL (Figure 12).

3.14. Calculation of IC₅₀ and SI

Half-maximal inhibitory concentration (IC₅₀, µg/mL) of LHE–PLGA NPs, free LHE, and empty PLGA NPs was evaluated in human non-tumor keratinocytes (HaCaT) and human cancer cell lines (HeLa and MDA-MB-231). The selectivity index (SI) was calculated as the ratio between the IC₅₀ in HaCaT cells and the IC₅₀ in each cancer cell line (HaCaT/HeLa and HaCaT/MDA-MB-231). IC₅₀ values >100 µg/mL indicate absence of relevant cytotoxicity within the tested concentration range. ND, not determined; NA, not applicable (

Table 5. IC₅₀ values and selectivity index of LHE-loaded PLGA NPs in human normal and cancer cell lines.

Treatment	IC50 HaCaT (µg/mL)	IC50 HeLa (µg/mL)	IC50 MDA-MB-231 (µg/mL)	SI (HaCaT/HeLa)	SI (HaCaT/MDA)
LHE–PLGA NPs	95	30	60	3.2	1.6
LHE (free extract)	>100	~85–100	>100	>1.0–1.2	ND
Empty PLGA NPs	>100	>100	>100	NA	NA

Abbreviations: IC₅₀, half-maximal inhibitory concentration; SI, selectivity index; ND, not determinable within 0–100 µg/mL; NA, not applicable.

).

3.15. Cell Death Evaluation

Cell death induced by the different treatments was evaluated at 24 h of exposure by fluorescence microscopy, distinguishing apoptosis (Apopxin Green, green fluorescence) and necrosis (7-AAD, red fluorescence) in HaCaT, HeLa, and MDA-MB-231 cells (Figure 13).

In the control group, all three cell lines exhibited minimal green and red fluorescence, indicating high cell viability and the absence of significant cell death.

Treatment with doxorubicin induced a pronounced cytotoxic effect in all cell lines, characterized by an intense red fluorescence, accompanied by a lower proportion of green fluorescence. This pattern suggests the predominance of necrosis, together with the presence of apoptotic cells, confirming the strong cytotoxic activity of the reference drug.

Treatment with unencapsulated HE (100 µg) produced a moderate effect on cell viability. In HeLa and MDA-MB-231 cells, a discrete increase in green fluorescence was observed, indicative of early apoptosis, while red fluorescence remained limited, suggesting less severe cellular damage compared with doxorubicin. In HaCaT cells, the effect was minimal, indicating lower cytotoxicity toward non-tumor cells.

In contrast, treatment with HE-PLGA NPs (100 µg) induced a marked increase in green fluorescence, particularly in HeLa and MDA-MB-231 cells, accompanied by low red fluorescence. This pattern indicates that the nanoformulated HE predominantly promotes apoptotic cell death, with minimal induction of necrosis. In HaCaT cells, fluorescence levels remained low, suggesting selectivity toward cancer cells and reduced toxicity in normal cells.

Overall, these results demonstrate that encapsulation of HE into PLGA NPs enhances the induction of apoptosis in cancer cells while reducing necrosis-associated damage, resulting in a more controlled cytotoxic profile compared with doxorubicin.

4. Discussion

This study investigated whether hexane extracts from different parts of A. monostachya retained the differential cytotoxic profile previously reported by our group [15]. LHE and its nanoformulation showed higher potency in HeLa cells than in HaCaT cells; however, HaCaT cytotoxicity was also observed at higher concentrations, indicating a limited in vitro therapeutic window. Therefore, selectivity is reported quantitatively using estimated IC₅₀ values and selectivity indices.

The qualitative phytochemical profile is consistent with classes of compounds previously associated with anticancer activity. For example, β-sitosterol and terpenoids such as betulin and betulinic acid have been reported to induce apoptosis in HeLa cells through mechanisms that involve caspases, mitochondrial dysfunction, and oxidative stress [55]. The presence of these compound classes may contribute to the activity observed in HeLa cells. Flavonoids have also been widely reported to influence redox balance and pathways related to cell-cycle control and cell death [56,57].

Beyond phenolic compounds, other phytochemical classes reported in plant extracts (e.g., alkaloids and saponins) have been linked to antiproliferative activity through diverse mechanisms, including effects on microtubule dynamics, cell-cycle regulation, and apoptosis-related pathways [58,59]. Sesquiterpene lactones, for instance, can interact with protein thiol groups and thereby modulate signaling and survival pathways, providing an additional mechanistic rationale reported in the literature [60,61]. Although our screening was qualitative, these reports provide a plausible context for the biological activity observed and support further chemical characterization of the active fraction.

Among the formulations evaluated, 5:50 provided the most balanced physicochemical profile in terms of particle size, PDI, EE%, and loading. NPs below ~200 nm are often considered suitable for cellular uptake and prolonged circulation in controlled-release settings [40,41]. An in vitro release study in our laboratory confirmed that LHE release from LHE–PLGA NPs was pH-dependent (pH 6.5: ~60–65% at 4–5 h and ~78–80% at 24 h; pH 7.0: ~12–15% at 5 h and ≤~18–19% at 24 h) [62], supporting a time- and pH-responsive behavior rather than an indiscriminate burst release; therefore, the system is described conservatively as pH-responsive rather than broadly controlled-release.

The unencapsulated LHE exhibited a concentration-dependent hemolytic profile, with hemolysis < 10% at ≤50 μg/mL, which is considered acceptable in preliminary in vitro hemocompatibility assessments [26]. In contrast, a pronounced increase in hemolysis was observed at higher concentrations, reaching ~80% at 500 μg/mL. This effect is consistent with the highly lipophilic nature of the extract and its interaction with erythrocyte membranes [35]. These results highlight the importance of dose control and further support nanoencapsulation as a strategy to improve hemocompatibility and reduce hemotoxic risk [21,26]. If systemic exposure or blood contact is anticipated, hemocompatibility becomes a critical parameter. Encapsulation in PLGA can reduce direct contact between lipophilic constituents and erythrocyte membranes, which may help lower hemolysis [42,50]. Together, our results support PLGA encapsulation as a practical strategy to modulate the blood-contact profile of this extract while maintaining a formulation compatible with delivery.

Several methods are available to prepare polymeric NPs; here we used nanoprecipitation because it is simple, rapid, and reproducible for hydrophobic compounds [63]. This method—also referred to as solvent displacement or interfacial deposition—relies on two miscible phases: an organic phase (acetone) containing dissolved PLGA and an aqueous phase (distilled water). Upon mixing, solvent diffusion promotes polymer precipitation and nanoparticle formation under conditions of polymer solubility in the organic phase, polymer insolubility in the aqueous phase, and solvent–water miscibility [64].

When testing different polymer/extract ratios, increasing the extract content was associated with larger NPs. Nanoparticle formation and size depend on multiple factors, including polymer concentration, solvent/non-solvent ratio, and stirring conditions [65]. Solvents with higher diffusion coefficients (e.g., acetone, acetonitrile) tend to favor smaller particles, and decreasing the solvent-to-non-solvent ratio can also reduce size [65]. In addition, higher polymer concentration can increase particle size during nucleation and growth, which is consistent with the increase observed when PLGA was increased from 50 to 60 mg [64,65].

Alshamsan (2013) encapsulated cucurbitacin I in PLGA NPs and compared two preparation methods, reporting a higher encapsulation efficiency (48.79%) with nanoprecipitation [66]. This is consistent with our choice of nanoprecipitation for a HE rich in hydrophobic constituents. Differences in encapsulation and physicochemical behavior may also depend on PLGA composition, molecular weight, and terminal groups; in our case, the PLGA used (Resomer^® RG 503 H) is acid-terminated, meaning it has free carboxyl groups at the chain ends, which confer a slightly more hydrophilic character compared to ester-terminated PLGA. Nevertheless, the high encapsulation efficiency (~92%) observed suggests that the core hydrophobic nature of the PLGA backbone still promotes favorable interactions with the lipophilic constituents of the extract.

SEM showed predominantly spherical NPs with a smooth surface. In the dry state, LHE-loaded NPs appeared larger and less aggregated than the empty formulation (Figure 6). These SEM observations align with the DLS measurements, which showed a mean hydrodynamic diameter of ~100 nm and a negative zeta potential (−26 mV), with no obvious self-aggregation under the tested conditions [67,68]. Raman spectroscopy provided complementary chemical evidence supporting incorporation of the extract into the PLGA matrix.

Akram et al. (2021) reported Raman and FT-IR spectra of PLGA NPs that were consistent with the main features observed in our spectra, including a band around ~1300 cm⁻¹ attributable to lactic acid units [69]. In FT-IR, characteristic PLGA bands have been described in the 1100–1250 cm⁻¹ and 1750–1760 cm⁻¹ regions, corresponding to ester and carbonyl groups [69]. Sechi et al. (2012) also reported PLGA-associated bands at 1453 cm⁻¹ (C–H) and at 1130 and 1044 cm⁻¹ (C–O stretching) [70].

Raman and FTIR analyses were used as complementary techniques to support the incorporation of the LHE within the polymeric matrix. The observed attenuation and partial overlap of characteristic extract bands in the nanoparticle spectra, compared with the free extract and blank polymer, are consistent with previous reports describing homogeneous distribution of bioactive compounds in polymeric systems. However, these techniques do not allow definitive discrimination of molecular amorphization, and therefore no claims regarding the amorphous state of the encapsulated extract are made.

A concentration-dependent effect was observed, and cytotoxicity was higher for the encapsulated formulation than for the crude extract. Previous work has suggested that increasing particle size may be associated with higher IC₅₀ values, consistent with reduced potency [56]. In line with this, the most active formulations in our experiments were among the smaller-size NPs. Syahputri et al. (2025) reported that Moringa oleifera-based NPs reduced HeLa viability by ~50% at 200 µg/mL, while Vero cells maintained >80% viability [71].

The encapsulation efficiency (EE%) and drug loading (%L) were calculated by considering the HE as a multicomponent bioactive fraction rather than individual compounds. Although representative metabolites were tentatively identified by GC–MS and UHPLC–MS, these analyses were qualitative in nature. Compound-specific quantification by HPLC would require certified analytical standards and fully validated methods, which were beyond the scope of this study. Nevertheless, the high overall encapsulation efficiency (~92%) and the observed biological activity suggest an effective incorporation of bioactive constituents into the polymeric matrix.

In the case of the methanolic extract of Vernonia greggii encapsulated in PLGA NPs, nuclei with pyknosis (condensation), rounding, and increased fluorescent intensity were observed from 300 µg/mL, along with a reduction in the percentage of adhered cell area, which is a pattern compatible with apoptosis [26].

The apoptosis and necrosis patterns observed by fluorescence microscopy agree with the cytotoxicity profiles obtained from IC₅₀ analyses. Compounds and formulations exhibiting lower IC₅₀ values generally correlated with a higher proportion of apoptotic cells, whereas treatments with higher IC₅₀ values induced limited cell death. This relationship supports apoptosis as a more efficient and controlled mechanism of anticancer activity compared with necrosis [72,73].

In contrast, HE-PLGA NPs exhibited lower IC₅₀ values in HeLa and MDA-MB-231 cells compared with free HE, which correlated with a marked increase in apoptotic signaling and minimal necrotic damage. This apoptotic-dominant profile suggests that nanoencapsulation enhances intracellular uptake and provides sustained drug release, promoting activation of programmed cell death pathways rather than acute membrane disruption [40,42,74].

Selective induction of apoptosis is a highly desirable feature in anticancer therapy, as it limits inflammation and collateral damage to surrounding healthy tissues [73,75]. Notably, the reduced apoptotic and necrotic response observed in HaCaT cells is consistent with the higher IC₅₀ values obtained for this non-tumor cell line, indicating preferential cytotoxicity toward cancer cells. Similar selectivity profiles have been reported for PLGA-based nanoparticle systems delivering bioactive compounds [42,76].

Despite the rapid development of alternative nanocarrier systems for tumor therapy, including gold nanoparticles, exosomes, and liposomes, PLGA-based NPs remain one of the most extensively investigated and clinically relevant platforms for drug delivery [77,78,79]. PLGA is a biodegradable and biocompatible polymer approved by regulatory agencies, and it degrades into lactic and glycolic acids that are naturally metabolized, thereby minimizing long-term toxicity concerns. While gold NPs exhibit promising photothermal and imaging properties, their clinical translation is limited by uncertainties related to biodegradation and long-term accumulation [77]. Exosome-based systems offer intrinsic biological compatibility but face significant challenges in large-scale production, standardization, and regulatory approval [78]. Liposomes have achieved clinical success; however, issues related to stability, premature drug leakage, and limited control over release kinetics persist [79]. In contrast, PLGA NPs provide a versatile and tunable delivery system, allowing controlled drug release, surface functionalization, and improved delivery of hydrophobic antineoplastic agents, which justifies their selection in the present study.

Overall, these findings indicate that the enhanced cytotoxic efficacy of HE-PLGA NPs is mechanistically linked to their ability to preferentially induce apoptosis while minimizing necrosis. This controlled mode of cell death supports a safer and more effective anticancer strategy compared with conventional chemotherapy and highlights the therapeutic potential of HE-loaded PLGA NPs.

The predominance of apoptotic (green) fluorescence and the limited necrotic (red) signal observed in HeLa and MDA-MB-231 cells support the activation of a controlled apoptotic mechanism rather than acute membrane damage. The reduced apoptotic and necrotic response observed in HaCaT cells may be attributed to lower nanoparticle uptake and/or enhanced antioxidant and DNA repair capacities in non-tumor cells, contributing to the observed selectivity of HE-PLGA NPs toward cancer cells.

These findings suggest that HE-PLGA NPs promote selective apoptosis in cancer cells, providing a mechanistic basis for their enhanced cytotoxicity and improved safety profile. However, further studies evaluating ROS levels, mitochondrial membrane potential, caspase activation, and Bax/Bcl-2 expression would be required to confirm this proposed mechanism.

5. Conclusions

To our knowledge, this is the first study to compare HE obtained from leaves, stems, and inflorescences of A. monostachya across tumor and non-tumorigenic cell lines, and to report the preparation and physicochemical characterization of LHE-loaded PLGA NPs.

LHE from leaves showed the highest selective cytotoxicity, with IC₅₀ values of ~70 µg/mL for HeLa and ~85 µg/mL for MDA-MB-231 cells, while no IC₅₀ was reached in HaCaT cells. The 5/50 LHE–PLGA formulation achieved a particle size of 131.4 nm, PDI of 0.122, and encapsulation efficiency of 92%, indicating a uniform and stable nanoparticle system. Hemolysis remained under 10% at concentrations up to 50 µg/mL. Cytotoxicity was enhanced in the encapsulated formulation compared to the crude extract and was associated with a predominantly apoptotic profile. These results support continued investigation of PLGA-based delivery strategies for lipophilic plant extracts. Future studies should expand in vitro testing and include in vivo evaluation to further assess efficacy and safety.