https://orcid.org/0000-0002-7244-3105
Figures
Abstract
Liver cancer, especially hepatocellular carcinoma (HCC), remains a major global health challenge, causing high mortality worldwide. Conventional chemotherapy often results in severe side effects due to its systemic distribution, which limits its effectiveness in targeting cancer cells specifically. The development of targeted drug delivery systems can enhance the precision and efficacy of chemotherapeutic agents while reducing their side effects. In this context, we used lactobionic acid (LA) as a targeting moiety due to its ability to bind to the asialoglycoprotein receptor (ASGPR), which is highly expressed on the surface of hepatocytes (liver cells). Conjugating lactobionic acid to liposomes creates an efficient delivery system that specifically targets liver cancer cells, thereby ensuring a higher uptake of the drug by these cells. Ultrasound is used to further facilitate drug release by enhancing drug targeting, promoting accumulation at the tumor site, and triggering drug release, thereby making the treatment more effective and less toxic. This study successfully synthesized stable DOX-loaded liposomes conjugated with lactobionic acid (LL) on their surface, with a size range of 89.4 ± 0.9 nm and a polydispersity index of 13.06 ± 3.5. Successful LA-DSPE-PEG-NH2 conjugation to the LL was confirmed via Fourier Transform Infrared spectroscopy. The sulfuric acid colorimetric assay quantified lactobionic acid conjugation as 14 ± 1.35% (w/w), while DOX encapsulation was measured at 40.1 ± 1.6%. LL exhibited strong absorption, fluorescence properties and good stability at 37 ˚C. LL showed controlled release via ultrasound, making it suitable for precise drug delivery. In vitro studies on HepG2 cells confirmed enhanced drug uptake and therapeutic efficacy. The effects of ultrasound-enhanced drug delivery to HepG2 cells demonstrated that the combination of ultrasound and targeted liposomes significantly increased the internalization of the drug (DOX) and triggered apoptosis in HepG2 cells, leading to cell death. Morphological observations on treated cells via phase contrast microscopy supported the signs of apoptosis, indicating LL’s potential to target liver cancer cells effectively.
Citation: Radha R, Anjum S, Abusamra RH, Paul V, Pitt WG, Husseini GA, et al. (2026) Ultrasound-triggered doxorubicin targeted delivery for liver cancer treatment: Reduced toxicity and improved efficacy. PLoS One 21(3): e0345161. https://doi.org/10.1371/journal.pone.0345161
Editor: Lei Zhang, University of Waterloo, CANADA
Received: August 19, 2025; Accepted: March 3, 2026; Published: March 24, 2026
Copyright: © 2026 Radha et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Data Availability: All relevant data are within the paper and its Supporting Information files.
Funding: This study was financially supported by the American University of Sharjah (https://www.aus.edu) in the form of grants received by GAH (FRG20-L-E48 and FRG22-C-E08), MHA (FRG24-C-S08), and the OAP program award received by GAH.
Competing interests: The authors have declared that no competing interests exist.
1. Introduction
Liver cancer continues to be a significant global public health challenge, with Hepatitis B (HBV) and Hepatitis C (HCV) infections being the primary risk factors [1]. Hepatocellular carcinoma (HCC), the most common form of primary liver cancer, accounted for around 830,180 deaths worldwide in 2020, making it the fourth leading cause of cancer-related mortality globally [2]. This trend is projected to worsen, with disease and deaths from liver cancer expected to increase by over 55% by 2040, driven by factors such as chronic hepatitis infections (hepatitis B and C), excessive alcohol consumption, and rising rates of obesity, metabolic disorders and fatty liver disease [2,3].
Preventive measures, early diagnosis, and effective treatment strategies are crucial in combating this rising threat. While chemotherapeutic drugs such as Atezolizumab, Bevacizumab, Doxorubicin, Daunorubicin, Cytarabine, Vincristine, Topotecan, Irinotecan, Teniposide, Cisplatin, Paclitaxel, and Nelarabine are highly effective in treating cancer, their use is often plagued by severe side effects, which continues to be a major concern for patients [4–8]. Emerging drug delivery systems (DDS) are demonstrating significant potential in increasing the concentration of therapeutic drugs within cancerous tissues, extending their circulation time, and boosting overall treatment effectiveness [9,10]. Among these systems, highly compatible nanoformulations, such as liposomes, nanoemulsions and micelles designed to encapsulate specific therapeutic agents, have shown considerable promise in targeted tumor therapies [11,12].
Liposomes have emerged as promising nanocarriers for drug delivery, offering unique advantages due to their biocompatibility and structural similarity to cellular membranes [13–16]. Composed of phospholipid bilayers, liposomes can encapsulate both hydrophilic and lipophilic drugs, making them highly useful for drug formulation [15]. Since their FDA approval in 1995, liposomes have become a well-researched and widely adopted platform in cancer treatment [17]. Their ability to improve drug bioavailability, enhance therapeutic outcomes, and minimize adverse effects has made them particularly appealing for commercialization. Medical experts widely accept liposomal encapsulation technology as a tissue-specific drug delivery method, as it forms a protective barrier around the drug, shielding it from enzymatic, redox, or other deleterious chemical reactions in the human body [18]. These attributes position liposomes as a highly promising option for delivering anticancer drugs in cancer treatment, especially in conjunction with emerging targeted therapies [19].
Traditional drug delivery methods often rely on passive diffusion or systemic circulation, leading to suboptimal drug concentrations at tumor sites and increased toxicity to healthy tissues [10,20]. New approaches, however, aim to address these challenges by utilizing active, controlled-release mechanisms. One such technique is ultrasound-triggered drug delivery, where externally applied and focused ultrasound waves cause localized changes in temperature and pressure, triggering the release of drugs from liposomes directly at the tumor site [13,21]. The integration of nanocarriers such as liposomes, dendrimers, solid lipid nanoparticles, and micelles with ultrasound technology further enhances the potential for targeted cancer therapies [22,23]. These nanocarriers can passively accumulate in tumors due to the enhanced permeability and retention (EPR) effect, a phenomenon that allows nanoparticles to concentrate in tumor tissues due to their leaky vasculature and poor lymphatic drainage of many cancerous tissues [10,24,25]. Additionally, combining passive targeting with active receptor-mediated endocytosis using specific ligands further increases drug accumulation and active uptake at the tumor site [26–29].
HCC is characterized by the overexpression of the Asialoglycoprotein Receptor (ASGPR), a liver-specific receptor predominantly found on hepatocytes and hepatoma cells [30,31]. ASGPR facilitates the uptake of desialylated glycoproteins via receptor-mediated endocytosis, making it an ideal target for developing therapies aimed at selectively targeting malignant liver cells [32–34]. Conjugating the surface of liposomes with lactobionic acid could promote targeted binding to ASGPR-overexpressing hepatoma cells, enhancing the uptake of drug-loaded liposomes by cancer cells [35,36]. This strategy enhances the therapeutic efficacy of the drug while significantly reducing off-target effects and toxicity in healthy tissues, offering a more precise and safer treatment approach for HCC [37].
Previous studies have shown that LA conjugation enhances drug uptake in HCC through ASGPR mediated targeting in liver tumors [38]. Another study by Samaei et al. reported the liposome-mediated phototherapy and other stimuli-responsive systems suppress HCC tumorigenesis [39]. Our LFUS-LA liposomes uniquely integrate two mechanisms that are rarely combined in a single platform. First, they achieve active ASGPR targeting via lactobionic acid. While LA-modified liposomes have been reported for hepatocyte targeting, none have coupled LA-mediated receptor targeting with non-thermal, low-frequency ultrasound activation. Second, they enable mechanical, non-thermal ultrasound-triggered drug release. Unlike investigational heat-sensitive liposomal doxorubicin formulations like ThermoDox, which depend on heat-induced phase transitions triggered by radiofrequency ablation or high-intensity focused ultrasound [40], our formulation is designed for LFUS-mediated mechanical destabilization, allowing drug release without hyperthermia and providing a fundamentally distinct release mechanism.
The combination of these two strategies produces a synergistic effect. To our knowledge, no prior study has reported the dual effect of LA targeting and LFUS-mediated mechanical release for DOX delivery to HCC. This dual mechanism enhances cellular uptake, ensures selective cytotoxicity, and strongly induces apoptosis in ASGPR-positive cells, effects that cannot be achieved by thermal liposomes or non-targeted ultrasound-responsive carriers. So here we present a novel liver cancer drug delivery strategy using liposomes to encapsulate doxorubicin (DOX), with ultrasound serving as a trigger to enhance drug release. By synthesizing DOX-loaded liposomes and optimizing ultrasound-mediated release and cellular uptake, we present a promising platform for targeted and effective chemotherapy.
2. Materials and methods
2.1. Chemicals/materials
Lactobionic acid, cholesterol (≥99%), Sephadex® G-25, chloroform, HEPES sodium salt, ammonium ferrothiocyanate, phosphate-buffered saline (PBS) and 2,4,6-trichloro-1,3,5-triazine (cyanuric chloride) were purchased from Sigma-Aldrich (St. Louis, MO, USA) and supplied by Labco LLC (Dubai, UAE). EDC (1-ethyl-3-(3-dimethylaminopropyl) and Sulfo-NHS (N-hydroxysulfosuccinimide) were procured from Thermo Fisher Scientific (Waltham, USA) through Biomedical Scientific Services LLC, UAE. Doxorubicin hydrochloride was from Euro Asia (Mumbai, India). Ammonium sulfate salt ((NH4)2SO4), Triton X-100, dimethyl sulfoxide (DMSO), and trypsin were from Sigma-Aldrich Chemie GmbH (Munich, Germany) and supplied by Labco LLC (Dubai, UAE). 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[amino (polyethylene glycol)-2000] (DSPE-PEG (2000)-NH2) were purchased from Avanti Polar Lipids Inc. (Alabaster, AL, USA), supplied by Labco LLC (Dubai, UAE). 200-nm polycarbonate filters and filter supports were purchased from Nucleopore Track Etch membrane filtration products. MTT salt (3-(4,5-dimethyl thiazolyl)-2,5-diphenyl-tetrazolium bromide), Dimethyl sulfoxide (DMSO), Roswell Park Memorial Institute (RPMI-1640) and Dulbecco’s Modified Eagle Medium (DMEM), were purchased from Sigma-Aldrich Chemie GmbH and supplied by Labco LLC (Dubai, UAE).
2.2. Cell lines and culture conditions
HepG2 cells known to express the asialoglycoprotein receptor (LA-positive), were used as a positive control for cytotoxicity studies. DOX cellular uptake in HepG2 cells was compared with two other cell lines: Jurkat (immortal human T lymphocytes) and NIH-3T3 (an embryonic mouse fibroblast cell line). All cell lines were purchased from the European Collection of Authenticated Cell Cultures (ECACC, Salisbury, UK). The cells were cultured in RPMI-1640 or DMEM media, both supplemented with 10% Fetal Bovine Serum (FBS) and 1% Penicillin/Streptomycin and maintained in a 5% CO2 incubator at 37 °C.
2.3. Major instrumentations for the study
The synthesis, characterization, and cytotoxicity/uptake evaluation of DOX-loaded lactobionic acid-conjugated liposomes (LL) were conducted using the following instruments. Liposomes were synthesized using a Stuart RE300 rotary evaporator, sonicated in an Elma D-78224 bath (Melrose Park, IL, USA), and extruded with an Avanti Polar Lipids extruder (Alabaster, USA). Size and polydispersity were determined by dynamic light scattering (DynaPro Nanostar, Wyatt Technology Corp., Santa Barbara, CA, USA). Fluorescence characterization was performed with an FLSP920 spectrometer (Edinburgh Instruments Ltd., Livingston, UK). To monitor the release of the encapsulated DOX, a 20-kHz low-frequency ultrasonic probe (model VCX750, Sonics and Materials Inc., Newtown, CT) was used, with fluorescence variations being monitored via a Quanta Master QM 30 Spectrofluorometer (Photon Technology International, Edison, NJ, USA). UV–Vis absorbance was measured with an Evolution 60 S spectrophotometer (Thermo Scientific, Waltham, Massachusetts, USA). Optical densities of 96-well plates were read with an Accu Reader (Metertech, Taiwan). Cellular uptake and apoptosis were analyzed using a Beckman Coulter FC500 (Indianapolis, United States) and BD FACS Calibur flow cytometer (BD FACS Calibur; Becton Dickinson, Franklin Lakes, NJ, USA), respectively.
2.4. Preparation of non-targeted liposomes and DOX loading
Liposomes were prepared via a modified film hydration method, using a rotary evaporator [10,41,42]. A lipid mixture of DPPC, DSPE-PEG (2000)-NH2, and cholesterol in a molar ratio of 13:1:6 was dissolved in 4 mL of chloroform in a round-bottom flask. The chloroform was evaporated under vacuum in a rotary evaporator, operated at 120 rpm and 50 °C for 15 minutes, resulting in the formation of a lipid film. The lipid film was then hydrated with a 0.11 M (NH4)2SO4 solution at pH 5.5, under continuous agitation at 100 rpm, 60 °C for 50 min. Following hydration, the liposomal suspension was sonicated in a 35-kHz ultrasonic bath for 2 min to result in multi lamellar liposomes. These liposomes were extruded at 60 °C using a mini extruder with 0.2-µm polycarbonate filters to produce unilamellar liposomes.
DOX encapsulation was performed via the remote (NH4)2SO4 transmembrane gradient method [43], where a pH gradient was established by passing the liposomes through a HEPES column at pH 7.4. DOX, at a concentration of 16 mg/mL HEPES buffer, was added to the liposomal solution in a 1:6 (w/w) ratio. The mixture was stirred for 45 min at 60 °C to enhance DOX loading. The final DOX-loaded liposomes were purified using size-exclusion chromatography on a Sephadex G-25 gel column, which was equilibrated with phosphate-buffered saline (PBS) at pH 7.4. The purified liposomes, referred to as DOX control liposomes (CL), were then collected for further analysis.
2.5. Preparation of targeted liposomes (LL)
The proposed mechanism for functionalization involved the EDC/NHS-mediated coupling of LA with DSPE, representing an effective strategy for the creation of functionalized lipids suitable for drug delivery applications [42]. In this reaction, the control liposome is conjugated with LA, as shown in the reaction scheme in Fig 1. The reaction used LA (0.01 mmol), NHS (25-fold excess), and EDC (5-fold excess). Initially, EDC, NHS, and LA were mixed in 400 µL of MES buffer and allowed to react at room temperature for 15 minutes. The resulting conjugate was then combined with DSPE/control liposomes at a 1:10 molar ratio of ligand to lipid. The reaction was allowed to continue at room temperature for 2 hours. The resulting conjugated liposomes were purified using Sephadex G-25, centrifuged, and stored, following the same protocol as the control liposomes (section 2.4).
Illustration showing the functionalization or conjugation reaction steps for DSPE-PEG-NH2 conjugation with Lactobionic acid (LA).
2.6. Characterization of liposomes (CL and LL)
2.6.1. Size and polydispersity analysis.
Dynamic light scattering (DLS) was performed to measure the hydrodynamic radius, polydispersity index, and size distribution of the liposomes at ambient temperature. The analysis was carried out using the DynaPro® NanoStar™ DLS system (Wyatt Technology Corp., CA, USA). For sample preparation, 10–15 µL of the liposome solution was diluted in 1 mL of PBS buffer. Triplicate samples were measured, averaged and reported in the results.
2.6.2. Fourier transform infrared spectroscopy (FTIR).
Fourier-transform infrared (FT-IR) spectroscopy (PerkinElmer FT-IR spectrometer, MA, USA) was performed to confirm the successful synthesis of the Lactate-DSPE conjugate. Samples of conjugate were prepared by drying the sample onto potassium bromide (KBr) pellets, followed by spectral acquisition. The resulting spectra were compared to those of the individual starting materials DSPE and lactobionic acid to confirm the occurrence of chemical conjugation based on characteristic functional group shifts and the appearance/disappearance of specific absorption bands.
2.6.3. Confirmation test for lactate conjugate formation- Phenol H2SO4 assay.
The conjugation of LA to the lipid was further verified through the phenol-sulfuric acid colorimetric assay. This widely used method for carbohydrate quantification is based on the acid-catalyzed hydrolysis of saccharides into monosaccharides, which subsequently undergo dehydration in the presence of concentrated sulfuric acid to form furfural or 5-hydroxymethylfurfural [44]. These intermediates react with phenol to form a stable, chromogenic complex, which is measurable at 490 nm.
Assay calibration was done using a stock solution of LA (2.1 mM). Aliquots of LA at various concentrations were prepared and mixed with 500 µL of 5% phenol solution, followed by vortexing for 10 seconds. Subsequently, 2.5 mL of concentrated sulfuric acid was added to each mixture, vortexed again for 10 seconds, and then incubated at room temperature for 30 min. Absorbance was recorded at 490 nm using a UV-Vis spectrophotometer. The intensity of the developed color correlated with the product concentration.
For sample analysis, 1 mg of LA-conjugated DSPE and 1 mg of unconjugated DSPE (serving as the negative control) were each dissolved in 500 µL of distilled water and subjected to the same assay protocol. A baseline (blank) solution was prepared by mixing 500 µL of distilled water with 2.5 mL of concentrated sulfuric acid, omitting both phenol and LA.
2.6.4. Quantification of phospholipid content in nanoliposomes.
The phospholipid content of liposome solutions was quantified using the Stewart assay [42]. Approximately 50 µL of the liposome solution was dried in a rotary evaporator under vacuum at 45 °C for 15 minutes. After drying, 1 mL of chloroform was added, and the mixture was sonicated for 10–15 min in a 35-kHz sonicating bath until a clear solution was obtained. Aliquots of varying volumes from this solution were then transferred to centrifuge tubes, and the total volume in each tube was adjusted to 2 mL with chloroform. Subsequently, 2 mL of ammonium ferrothiocyanate reagent was added to each tube. The mixture was thoroughly vortexed and centrifuged at 1000 rpm for 10 minutes. After centrifugation, the dark upper aqueous layer was carefully removed, ensuring no contact with the lower chloroform layer. The transparent chloroform layer, containing the extracted phospholipids, was transferred to a quartz cuvette for absorbance measurement. The optical density of the chloroform layer was measured at 485 nm using a UV-Vis spectrophotometer with a blank sample consisting of chloroform and assay solution.
2.6.5 Absorbance and fluorescence profiles by UV-Vis and fluorescence spectroscopy.
The encapsulation of DOX within liposomes was quantified using both UV-Vis and fluorescence spectroscopy. UV-visible absorption spectra of the liposomal formulation (LL) were recorded over a wavelength range of 300–700 nm. Fluorescence measurements were conducted with an excitation wavelength of 480 nm, and emission spectra were collected between 500–800 nm. To confirm successful DOX encapsulation, complete release was induced by adding 20 µL of Triton-X to the LL suspension, after which absorbance and fluorescence spectra were re-measured.
2.7. Low-frequency ultrasound (LFUS) triggered release of DOX
DOX release from liposomes was triggered using a low-frequency ultrasonic device (VCX750 model, Sonics & Materials Inc., Newtown, CT, USA), operating at 20 kHz. Acoustic parameters were carefully calibrated to ensure reproducibility and accurate interpretation of results. The 20-kHz probe system was calibrated using a Bruel & Kjaer 8103 hydrophone (Bruel & Kjaer, Nærum, Denmark). Acoustic pressures and spatial-average intensities at 20% and 30% amplitude (as displayed on the monitor) are reported. The corresponding power densities are 6.2 mW/cm² (20%) and 10 mW/cm² (30%), while the root-mean-square acoustic pressures are 9.6 kPa (20%) and 12.2 kPa (30%). Peak-to-peak pressures were 13.6 kPa (20%) and 17.3 kPa (30%). For the 35-kHz bath, the manufacturer-rated power was verified using the same hydrophone placed at the bath center, and the measured intensity of 1 W/cm². LFUS-triggered release experiments employed pulsed ultrasound with a duty cycle of 20 seconds on and 10 seconds off. Temperature was monitored in real time using a thermocouple during all sonication experiments, and the increase did not exceed 1.5 °C. This minimal rise, attributed to pulsing, confirms that the observed release behavior is primarily mechanical rather than thermal.
Fluorescence changes were monitored using a Quanta Master QM 30 Phosphorescence spectrofluorometer (Photon Technology International, Edison, NJ, USA), with excitation and emission wavelengths set at 470 nm and 560 nm, respectively. Baseline fluorescence (I0) was obtained by recording for 30 seconds before the start of sonication pulses. Subsequently, pulsed ultrasound was applied in cycles of 20 seconds on and 10 seconds off, continuing until a steady release was observed (total ultrasound exposure time of 4 min). Following the sonication sequence, Triton X-100 was added to lyse all the remaining liposomes and ensure complete DOX release. Experiments were conducted at two power densities: 6.2 mW/cm² (20% on the machine display) and 10 mW/cm² (30% on the machine display).
The percentage fraction release of DOX is calculated using the formula:
Where Iₜ represents the fluorescence intensity at time t, I₀ is the baseline intensity, and ITX is the intensity after Triton X-100 addition.
2.8. Stability and release studies on liposomes at 37 °C
Stability and the in vitro release studies on liposomes are essential to confirm their safe use in clinical applications. Aliquots of liposomes were mixed with 3 mL of PBS (pH 7.4) in a glass cuvette and incubated at 37 °C. At regular intervals, the samples were analyzed using a fluorometer, and the fluorescence spectra were recorded to calculate the percentage of DOX released over time. The release percentage at each time point was calculated based on the fluorescence readings using the equation (1).
2.9. In vitro experiments
2.9.1. Ultrasound conditions for cell studies.
Cells were exposed to ultrasound in a 35-kHz sonication bath (Branson 3510-DTH Ultrasonic Cleaner, Ohio, USA) at a power density of 1 W/cm², measured using a low-frequency hydrophone (Bruel & Kjaer 8103, Denmark).
2.9.2. Cytotoxicity assays.
The cytotoxicity of CL and LL on HepG2 cells, both with and without ultrasound treatment, was assessed using the MTT assay, to evaluate the treatment efficacy [10,45]. HepG2 cells were seeded in 12-well plates at a density of 1 × 10⁵ cells per well and incubated for 24 hours. Following this, cells were treated with varying concentrations of CL or LL in fresh media and incubated for 3 hours. One set of plates was subjected to ultrasound treatment for 1 min. The cells of treated and untreated plates were washed with PBS, treated media was replaced with fresh media and incubated for an additional 24 hours in a 5% CO2 incubator. After incubation, the cells were washed twice with phosphate-buffered saline (PBS, pH 7.4), followed by the addition of MTT (0.5 mg/mL). The MTT assay relies on the reduction of MTT to formazan by metabolically active cells [46,47]. Subsequently, the plate was incubated for 4 hours at 37 °C, followed by the addition of 1 mL of dimethyl sulfoxide (DMSO) to solubilize the formazan crystals. The optical density was measured at 570 nm using a microplate spectrophotometer, with untreated wells serving as controls. Cell viability was calculated by comparing the mean optical density of treated cells to that of control cells. Optimization studies were initially conducted to determine the DOX IC50, which was then used for subsequent experiments.
Culture viability was calculated by the following relationship:
2.9.3. DOX Uptake studies by Flow cytometry analysis.
For the DOX uptake studies, three different cell lines, HepG2, Jurkat, and 3T3 were used. The cells were seeded at a density of 1 × 105 cells per well in 12-well plates and incubated for 24 hours under standard culture conditions (37°C, 5% CO2, and 95% humidity). After incubation, the cells were treated with CL or LL (at a final DOX concentration of 8 µM) and further incubated under the same conditions for 3 hours. Additionally, one plate from each cell line was subjected to LFUS at 35 kHz for 1 min using a sonicating bath. After treatment, the cells were washed, harvested, pelleted, and resuspended in phosphate-buffered saline (PBS). Fluorescence analysis was then conducted using flow cytometry (Beckman Coulter FC500), with an excitation wavelength of 488 nm and emission fluorescence of DOX at 585 nm.
2.9.4. DAPI staining and phase-contrast microscopy.
The study used DAPI (4′,6-diamidino-2-phenylindole) staining, an effective method for visualizing nuclear morphology and evaluating DNA integrity in cells treated with DOX-loaded liposomes. The experiment began by culturing cell lines on a coverslip (3 × 105 cells per well in a 6-well plate), as detailed in Section 2.9.3, which were then treated with DOX-encapsulated liposomes (CL or LL) with or without ultrasound treatment. Cells on the coverslip were then fixed with 4% paraformaldehyde for 10–15 min to preserve their structures. The fixed cells were permeabilized to allow DAPI to enter the cells and incubated with DAPI at a concentration of 0.1–1 µg/mL for 5–15 min, enabling the dye to bind to DNA. Excess DAPI was washed away with phosphate-buffered saline (PBS, pH 7.0). The samples were mounted and examined under a fluorescence microscope using UV excitation, resulting in bright blue fluorescence of the nuclei.
2.9.5. Phase-contrast microscopy.
Phase-contrast microscopy was used to analyze the morphological alterations in HepG2 cells, such as cellular shrinkage, membrane blebbing, and cell rounding, in response to various treatments. A comparative analysis was conducted between ultrasound-treated and untreated cells, with and without the influence of targeted ligand-modified liposomes (LL).
2.9.6. FITC Annexin-V/propidium iodide (PI) apoptosis assay.
To assess apoptosis under various treatment conditions, HepG2 cells were cultured in 6-well plates at a density of 3 × 105 cells per well. After overnight incubation, cells were exposed to liposomal formulations (CL or LL) with and without ultrasound treatment. After 3 hours, the cells were washed in PBS and incubated for 24-hour in fresh media in 5% CO2 incubator.The cells were washed with PBS, harvested, and resuspended in 0.5 mL of binding buffer at a density of 1 × 105 cells per sample. Annexin V (5 µL) and PI (10 µL) were added to each sample under light-protected conditions. After incubation for 20 min in the dark, apoptotic cell populations were quantified using flow cytometry (BD FACS Calibur; Becton Dickinson, Franklin Lakes, NJ, USA). Cells stained positive for Annexin V-FITC but negative for PI were classified as early apoptotic, whereas cells positive for both Annexin V-FITC and PI were considered late apoptotic.
2.10. Statistical analysis
All experiments and cytotoxicity tests were performed using at least 3 different batches of freshly prepared liposomes, with each experiment conducted in triplicate. Statistical analysis was done on results using GraphPad Prism (version 8.4.3), and data comparisons between different treatment conditions were analyzed using the ANOVA test.
3. Results
The combination of drug delivery systems with the use of external physical triggers, such as ultrasound, holds great promise for enhancing the specificity and therapeutic efficacy of chemotherapy. In this study, we focus on the design of LA-conjugated liposomes (LL) for the targeted delivery of DOX to HCC cells, a major form of liver cancer, in combination with ultrasound to mediate controlled and targeted drug release.
3.1. Characterization of liposome
Characterization studies confirmed the successful formation of LA-functionalized DOX-loaded liposomes (LL). Dynamic light scattering analysis revealed that both control liposomes (CL) and LL formulations exhibited nanoscale dimensions, with mean particle sizes of 86.1 ± 2.1 nm and 89.4 ± 0.9 nm, respectively (Fig 2). The polydispersity indices (PDI) of CL and LL were measured to be 11.2 ± 2.5 and 13.06 ± 3.5, respectively, indicating a uniform particle size distribution. Long-term stability studies conducted over a 10-week period at 4 °C showed a minimal variation in particle size and PDI, confirming the colloidal stability of the liposomal formulations.
Left Y-axis represents the hydrodynamic radius (nm) and right Y-axis indicates the polydispersity index.
The Stewart assay quantified the lipid concentration in the liposomes. NH2 liposomes (CL) had an average DPPC content of 7.3 ± 1.3 mg per mL of LL suspension, while LA liposomes contained an average of 6.9 ± 1.1 mg of DPPC, the primary lipid component of the liposomes.
The successful conjugation of lactobionic acid (LA) to DSPE-PEG-NH2 was confirmed using Fourier transform infrared (FTIR) spectroscopy (Fig 3). The FTIR spectra of the unmodified DSPE-PEG-NH2 (negative control: represented as DSPE) and the DSPE-PEG-LA conjugate were compared to assess functional group transformations. In the DSPE-PEG-LA spectrum, a distinct peak at 1659 cm ⁻ ¹ was observed, corresponding to C = O stretching vibrations of an amide bond, provides clear evidence of covalent bond formation between the carboxyl group of LA and the amine group of DSPE-PEG-NH2 [48], [49]. This is a critical indicator of successful amide bond synthesis and is consistent with previously reported conjugation strategies. Additionally, peak at 721 cm ⁻ 1, corresponding to C–O–C ether stretching, was noted and showed a slight shift compared to the control, suggesting structural changes in the PEG chain upon conjugation [38,50]. A broad O–H stretching band between 3200 and 3400 cm ⁻ 1 in the conjugate, indicating the presence of hydroxyl groups from LA. Additional peaks formation in the 1100–1140 cm ⁻ 1 region (visible between marked lines at 1091 and 1240), corresponding to C–O stretching vibrations typical of polysaccharides, were observed.
FTIR spectra comparing DSPE-PEG-NH2 (unreacted lipid), LA, a physical mixture of DSPE and LA (DSPE-LA mix), and the chemically conjugated product (DSPE-LA conjugate). The appearance of new peaks and characteristic shifts is highlighted by dashed lines, indicating successful covalent conjugation.
To further confirm the successful conjugation of LA to DSPE-PEG-NH2, phenol-sulfuric acid colorimetric assay was performed. The assay confirmed the presence of LA in the conjugated lipid samples, with minimal interference from other formulation components. Quantitative analysis revealed that approximately 14 ± 1.35% (w/w. of DSPE-LA) of lactobionic acid was successfully conjugated to DSPE-PEG-NH2. This result provided further evidence supporting the effectiveness of the LA conjugation and complements the FTIR findings.
3.2. Absorbance, emission profiles and encapsulation efficiency
The successful encapsulation of DOX inside LL was confirmed using UV-visible spectroscopy. As shown in Fig 4A (left Y-axis), the LL formulation exhibited a characteristic absorbance peak at 490 nm, corresponding to DOX, confirming its presence within the liposomal core. The encapsulation efficiency of DOX was determined to be 40.1 ± 1.6%, calculated relative to the initial amount of DOX used for liposome loading.
A) Absorbance spectrum (Left Y-axis) and fluorescence emission spectra (Right Y-axis) of LL. B) The emission spectra of LL before and after the addition of Triton X-100 (Ex: 490 nm, Em: 500–900 nm) confirm significant DOX loading.
Fig 4A (right Y-axis) displays the emission profile of LL recorded at an excitation wavelength of 490 nm, over the emission range of 500−900 nm. The emission spectrum confirmed the characteristic fluorescence of encapsulated DOX. To evaluate liposomal integrity and confirm encapsulation stability, Triton X-100, a non-ionic surfactant known to disrupt lipid bilayers [42], was added to the LL formulation. As shown in Fig 4B, the addition of Triton X-100 resulted in a > 5-fold increase in fluorescence intensity, indicating the release and dilution of encapsulated DOX into the surrounding medium. This significant fluorescence enhancement demonstrates that DOX was primarily retained within the liposomal core under intact conditions, with minimal leakage prior to membrane disruption. The mechanism of DOX release upon Triton X-100 treatment through surfactant-induced membrane solubilization is illustrated in S1 File.
3.3. Stability and LFUS release studies
The liposomes demonstrated favorable stability and release kinetics when exposed to ultrasound, indicating potential for clinical application. The stability of both CL and LL incubated in PBS (pH 7.0) at 37 °C was assessed by comparing the rate of DOX release after 3 and 24 hours. Triton X-100 was used to lyse the liposomes, releasing 100% of the encapsulated DOX.
As shown in Fig 5, there was no significant difference in DOX release between CL (3 h: 2.69% ± 0.264; 24 h: 13.29% ± 0.303) and LL (3 h: 4.5% ± 2.2; 24 h: 17.3% ± 1.3). This enhanced stability and minimal release at 37 °C will help minimize toxicity to normal tissues during clinical applications. No significant difference in DOX release was observed between CL and LL throughout the incubation period.
A) Stability studies at 37 °C. Liposomes (CL and LL) were incubated at 37 °C, with fluorescence measurements taken for each sample over 60 s after a specified incubation period (3 and 24 hours). The results represent the average of three liposome batches. B) Normalized-averaged DOX release profiles of CL and LL at different US-intensities. Experiments were conducted at two power densities [6.2 mW/cm2 (20%) and 10 mW/cm2 (30%)].
Low-frequency sonication was used to trigger DOX release from both CL and LL (3 batches with three replicates each) using a 20-kHz probe at two power densities [6.2 mW/cm2 (20%) and 10 mW/cm2 (30%)]. DOX release was tracked by changes in fluorescence intensity. Fig 5B shows the normalized average release profiles (calculated using equation 1), where a sharp increase in fluorescence occurred during the “on” phase of sonication. No increase was observed during the “pulse off” phase. Additionally, DOX release increased with higher power densities (30%− 10 mW/cm2).
3.4. Cytotoxicity and DOX uptake studies
To determine the inhibitory concentration (IC50) of DOX on HepG2 cancer cell lines, an MTT assay was performed using both free DOX and DOX-loaded liposomes. As shown in Fig 6A, increasing concentrations of DOX led to greater toxicity and cell death, with the IC50 estimated as approximately 7.8 ± 0.5 µM.
B) MTT assay on HepG2 cell line with conjugated (LL) and non-conjugated liposomes (CL) with and without LFUS. Liposomes with DOX concentration of 8 µM (LL and CL) were used for the assays. The data illustrates the enhanced cytotoxicity in ASGPR-positive HepG2 cells, particularly with LL and US treatment. The results represent the mean value of 3 different experiment data and error bar indicates ±SD.
The cytotoxic effects of the liposomal formulations with DOX concentration of 8 µM (LL and CL) were assessed using the MTT assay after a 30-second treatment with low-frequency ultrasound (LFUS), as shown in Fig 6B. Ultrasound significantly enhanced the cytotoxicity of the LL-liposomes on HepG2 cells, likely due to increased permeability of both cellular and liposomal membranes, which facilitated higher uptake of the therapeutic agents (DOX). Ultrasound treatment combined with LL resulted in a statistically significant enhancement in DOX uptake (****p < 0.0001) (Cell viability, 36.7 ± 5.9%), compared to both control liposomes (CL) (Cell viability, 68 ± 1.4%), and CL subjected to ultrasound exposure (Cell viability, 62.5 ± 0.75%).
3.5. Cellular DOX uptake studies
Flow cytometry was performed to quantify the intracellular uptake of DOX in three different cell lines: HepG2 (ASGPR-positive), Jurkat, and 3T3 (ASGPR-negative), with the treatment of CL and LL. The results, presented in Fig 7, show that HepG2 cells treated with LL exhibited a significantly increased mean fluorescence intensity (MFI) of 2043, which represents a 105% increase compared to the MFI of 992 observed in cells treated with CL. This enhanced uptake is attributed to receptor-mediated endocytosis via the asialoglycoprotein receptor (ASGPR), which is expressed in HepG2 cells. In contrast, only modest increases in DOX uptake were observed in ASGPR-negative Jurkat and 3T3 cells, with LL treatment yielding 34% and 69% increases in MFI, respectively.
Flow cytometry analyses on different cell lines (HepG2, Jurkat, and 3T3) to evaluate the effects of CL and LL, both with and without ultrasound treatment.
Additionally, the effect of LFUS exposure on DOX uptake was evaluated. In HepG2 cells, LL treatment combined with US resulted in a 136% increase in MFI compared to non-US-treated LL cells (Fig 7A), indicating a synergistic effect between ligand-mediated targeting and ultrasound-enhanced permeability. In Jurkat and 3T3 cells, US exposure led to moderate increases in uptake of 33% and 13%, respectively (Fig 7B and 7C).
3.6. Phase contrast microscopy and DAPI staining
The morphological consequences of 3-hour Doxorubicin/CL/LL with or without ultrasound exposures on the HepG2 cells are shown in Fig 8. Notable morphological alterations included cell shrinkage, membrane blebbing, and cellular rounding, changes indicative of apoptotic progression. Cells exposed to ultrasound alone (without drug or liposome treatment) retained normal morphology, confirming that ultrasound by itself did not cause cytotoxicity. Following treatment with liposomes and ultrasound exposure, a noticeable change in cellular morphology was observed. Cells exposed to LFUS exhibited pronounced shrinkage and extensive blebbing, which are characteristic features of apoptosis [51]. The cells treated with free DOX showed fewer adherent cells and blebbing-like protrusions as compared to control cells. A similar type of change was found in the case of treatment with targeted liposome (LL) with ultrasound exposure; a portion of HepG2 cells were impaired and dead, the cell membrane formed bubble-like protrusions, and they lost adhesion properties.
Phase contrast microscopy images of HepG2 cells treated with CL and LL, both with and without LFUS. Images were captured to assess morphological changes upon treatments.
The DAPI staining of HepG2 cells treated with either CL or LL for 3 hours, both with and without US, provided valuable insights into the internalization of DOX. DAPI, a fluorescent stain that binds specifically to DNA [52], stained the nuclei of the cells blue, allowing for clear visualization of cellular morphology and nuclear characteristics (S1 File). Upon treatment with liposomal formulations, it was noted that DOX was effectively internalized by the HepG2 cells. The presence of a more intense red fluorescence in the nuclei of cells exposed to ultrasound suggests a significantly higher concentration of DOX within these nuclei compared to untreated cells or those not exposed to ultrasound. This enhanced uptake can be attributed to the ultrasound’s ability to increase cell membrane permeability, facilitating the entry of the drug-loaded liposomes into the cells.
The overlay images (S1 File), which combine the blue DAPI staining with the red fluorescence of DOX, illustrate co-localization within the nuclei. This observation strongly suggests that DOX not only penetrates cells but also intercalates with DNA in the cell nucleus. The intercalation of DOX with DNA is a critical aspect of its mechanism of action, as it disrupts the normal function of DNA, leading to cytotoxic effects [53]. These findings highlight the effectiveness of liposomal formulations, particularly in combination with ultrasound treatment, in enhancing drug delivery and accumulation within the target cells, thereby improving the potential therapeutic efficacy against cancer cells such as HepG2.
3.7. Apoptosis induced by LL with US
Flow cytometry analysis was conducted to investigate cell death in hepatocellular carcinoma (HepG2) cells treated with LL and US exposure. The HepG2 cells were treated with free DOX, CL, and LL, both with and without US treatment (Fig 9). To eliminate interference from DOX fluorescence (which has an excitation and emission range of 470–560 nm), we normalized the fluorescence readings of DOX-treated cells to those of unstained control cells prior to the analysis of the stained samples. The results indicated the following percentages of apoptotic cells (annexin V positive): 15.19%, 25.28%, 65.45%, 21.38%, 34.83%, 64.7%, and 76.56% in control cells, control cells exposed to US, free DOX, CL, CL with US exposure, LL, and LL with US treatment, respectively (Fig 9). Notably, LL treatment in conjunction with US exposure significantly increased the percentage of apoptotic cells in HepG2 cells. A marked increase in annexin V-positive cells was observed, indicating the induction of both early and late stages of apoptosis. Furthermore, HepG2 cells treated with liposomes (both CL and LL) combined with US exposure exhibited a significant (p < 0.05) increase in the percentage of apoptotic cells (34.83% for CL and 76.56% for LL), as compared to control cells.
Annexin V-FITC/PI double staining analysis revealing apoptosis in HepG2 cells. Each quadrant description is as follows: Upper right quadrant (Q2)-dead cells in late stage of apoptosis; lower right quadrant (Q3)-cells undergoing apoptosis (early-stage apoptotic cells); lower left quadrant (Q4) – healthy viable cells and upper left (Q1) represents non-viable necrotic stage. The cells treated with free DOX and targeted liposome (LL) with US exposure showed (LL + US) significantly (p < 0.0001) high late apoptotic cells as compared to control cells (insert). The data show mean ± SD percentages of late apoptosis in HepG2 cells, treated with different conditions in terms of liposome or US treatment.
4. Discussion
Chemotherapy is essential in cancer treatment, but its side effects, such as cardiac toxicity, adversely affect patient quality of life and challenge healthcare systems [24,54]. Current drug delivery methods often fail to target cancer cells efficiently, worsening these challenges and widening healthcare disparities. This study presents a novel therapeutic approach that integrates ligand-targeted liposomal drug delivery with external ultrasound stimulation to enhance chemotherapeutic efficacy in HCC. This system addresses major limitations of conventional chemotherapy, such as poor tumor selectivity, systemic toxicity, and lack of controlled drug release. DOX-loaded liposomes, surface-functionalized with lactobionic acid (LA), a galactose-rich ligand targeting the ASGPR [55], enabled active targeting and controlled drug release by LFUS.
Liposomes, because of their amphiphilic bilayer structure, can encapsulate both hydrophilic and hydrophobic drugs, offering a favorable platform for controlled delivery [10,42]. The liposomes prepared in this study exhibited a mean hydrodynamic diameter of approximately 89.4 ± 0.9 nm (Fig 2), an optimal size for passive accumulation in tumor tissues via the enhanced permeability and retention (EPR) effect, which ensures improved drug delivery efficacy in clinical applications [56]. The polydispersity indices (%PDI) of CL and LL were measured to be 11.2 ± 2.5 and 13.06 ± 3.5 (Fig 2), respectively, indicating a uniform particle size distribution and conforming to the generally accepted threshold of <20% for DLS-based assessments [57].
The observed band shifts in FTIR data (Fig 3) strongly supports the successful conjugation of lactobionic acid to DSPE-PEG-NH2.The stable LA conjugates minimize off-target binding and thereby enhance targeting accuracy and overall therapeutic efficiency. The liposomal formulation also demonstrated excellent physicochemical stability, retaining structural integrity and drug encapsulation (Fig 5A). This level of stability is important for clinical applications, ensuring extended shelf-life and consistent therapeutic performance [10]. A key feature of the present study is the use of LFUS to trigger controlled drug release from the liposomes (Fig 5B). Ultrasound-induced cavitation and localized heating cause the destabilization of liposomal membranes, facilitating the release of encapsulated DOX. The precision with which ultrasound can be applied allows for the release of the drug specifically at the site of the tumor, minimizing exposure to surrounding healthy tissues [25,58]. This controlled release mechanism was demonstrated in vitro, where DOX release was significantly enhanced under LFUS exposure, with higher power densities leading to increased drug release. The release profiles exhibited sharp increases during the “on” phase of sonication and no release during the “pulse off” phase, confirming the ultrasound-triggered drug release mechanism (Fig 5B). The DOX release from LL under US exhibited a similar pattern to previously reported studies of other conjugated liposomes (Herceptin, cRGD) [14,59].
Surface conjugation with lactobionic acid facilitated active targeting toward HepG2 cells, which were known to overexpress ASGPR. This dual-targeting strategy, passive (EPR) and active (ASGPR-mediated endocytosis), helped to enhance drug (DOX) accumulation in HepG2 cells while reducing overall cell toxicity (Fig 6). Importantly, the system was engineered to be responsive to LFUS, which induces acoustic cavitation effects, thereby destabilizing the liposomal bilayer and triggering rapid drug release.
The in vitro cell culture MTT experiments on HepG2 cell lines produced encouraging results. MTT assays on HepG2 hepatocellular carcinoma cell line revealed a significantly higher cytotoxic effect in cells treated with LL compared to CL, especially when ultrasound was applied (Fig 6B). The enhanced cytotoxicity in the ultrasound-treated group can be attributed to ultrasound-mediated sonoporation, which increases membrane permeability and facilitates greater intracellular uptake of DOX. Flow cytometry further supported this result, with a 136% increase in mean fluorescence intensity (MFI) of internalized DOX observed in ultrasound-treated HepG2 cells exposed to LL, relative to untreated controls (Fig 7A). Cellular uptake specificity was further confirmed by comparing the internalization of LL in ASGPR-expressing HepG2 cells with that in ASGPR-negative Jurkat (Fig 7B) and 3T3 cells (Fig 7C), which was further supported by DAPI images (S1 File). The significantly higher uptake in HepG2 cells underscores the efficacy of LA-mediated targeting and supports the role of receptor-ligand interactions in the selective delivery of drugs [60].
Furthermore, apoptosis assays using annexin V-FITC/PI staining demonstrated a markedly elevated proportion of apoptotic cells in the LL + US group, with 76.56% undergoing programmed cell death (Fig 9). The rate of apoptosis in cells treated with liposomes and ultrasound was significantly higher compared to control groups receiving either treatment alone. An interesting finding of this study was that free DOX-treated cells showed increased early apoptotic and necrotic cells compared to control and liposome-treated groups (Fig 9). However, the liposome and ultrasound exposed group (LL + US) showed an increase in early and late apoptotic cells compared to the control (CL) and free DOX-treated cells. The results were in alignment with reported ultrasound-mediated drug delivery studies [24]. Morphological observations via phase contrast microscopy supported the signs of apoptosis (Fig 8), including membrane blebbing, cytoplasmic condensation, and loss of adherence, particularly in cells exposed to both LL and ultrasound. The observed changes provide visual evidence of the cytotoxic effects of the liposomal formulations, emphasizing the role of ultrasound in enhancing the delivery and effectiveness of the treatments [25]. These results are consistent with previous reported findings that US induces apoptosis in cancer cell lines [10,55]. Our study findings suggest that the combined effects of targeted delivery and ultrasound-triggered release not only improve intracellular DOX accumulation but also amplify its pro-apoptotic activity.
Although LFUS-triggered DOX release exhibited promising outcomes under in vitro conditions, the limited tissue penetration of ultrasound may constrain its therapeutic efficacy in deep-seated malignancies, such as hepatic tumors. Focused or image-guided ultrasound systems as well as minimally invasive delivery techniques could be employed to enhance acoustic penetration and spatial precision, thereby reducing off-target effects on normal liver tissue [61,62]. The integration of such strategies may further improve the translational potential of LFUS-mediated drug delivery for the treatment of tumors located in deeper anatomical regions.
5. Conclusions
The study confirmed the successful preparation of stable DOX-loaded liposomes and their effective conjugation with lactobionic acid. The LL exhibited solid absorption and fluorescence characteristics, which are important for monitoring drug distribution and effectiveness. The excellent stability and ultrasound-mediated controlled-release kinetics of LL indicate its potential for precise drug delivery in a clinical setting.
The promising outcomes observed in the in vitro cell culture studies on HepG2 cell lines align well with the research objectives, supporting the hypothesis that our liposomal formulations can enhance drug uptake and therapeutic efficacy. These promising results suggest that the liposomes can effectively deliver DOX to ASGPR-positive cancer cells, maximizing their cytotoxic effects while minimizing potential side effects on healthy tissues. The lactobionic acid-targeted liposomes effectively accumulated in HepG2, as confirmed by drug uptake and microscopy, highlighting the specificity of drug delivery to HepG2 cells through receptor-mediated endocytosis. This targeted approach ensured a higher concentration of the therapeutic agent within the tumor microenvironment, leading to enhanced cytotoxic effects. Ultrasound exposure further improved drug release from the liposomes, inducing permeabilization of the cell membrane and facilitating deeper penetration into cancerous tissues. The findings suggest that the combination of lactobionic acid-targeted liposomes with ultrasound can serve as an effective strategy for enhancing cytotoxicity in HCC.
While this study demonstrates the strong potential of lactobionic acid–functionalized liposomes for ultrasound-responsive, receptor-targeted drug delivery to hepatocellular carcinoma cells, several limitations remain. The current work was confined to two-dimensional in vitro models, which do not fully replicate the complex architecture, perfusion dynamics, and extracellular matrix interactions present in the liver tumor microenvironment. Future studies should employ three-dimensional spheroid or organoid models to better simulate in vivo conditions and validate penetration efficiency under ultrasound exposure. Moreover, hemocompatibility and serum stability assessments are required to evaluate the safety of the formulation for systemic administration. Additional in vivo pharmacokinetic and biodistribution investigations will be necessary to determine drug accumulation, clearance profiles, and potential off-target effects. Addressing these translational barriers will be crucial for advancing the lactobionic acid-liposome platform toward preclinical and clinical applications in targeted liver cancer therapy.
Supporting information
S1 File. Supporting figures and minimal datasets underlying the analyses.
This file contains additional supporting figures and tables, including the minimal datasets used to generate the quantitative analyses for Figs 2–7 and Fig 9.
https://doi.org/10.1371/journal.pone.0345161.s001
(PDF)
Acknowledgments
The authors acknowledge the technical support of the Chemical and Biological Engineering Department, Office of Research, Department of Biology, Chemistry and Environmental Sciences at the American University of Sharjah, UAE. We acknowledge Mr. Manju Nidagodu Jayakumar, Research Institute for Medical and Health Sciences, University of Sharjah, for his assistance with the flow cytometry analyses for the apoptosis assay and support for phase contrast microscopy.
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