Early diagnosis of pancreatic ductal adenocarcinoma (PDAC) is challenging because of its depth, which often leads to misdiagnosis during ultrasound examinations. The unique PDAC tumor microenvironment (TME) is characterized by significant fibrous tissue growth, and high interstitial pressure hinders drug penetration into tumors. Additionally, hypoxia and immune suppression within the tumor contribute to poor responses to radiotherapy and chemotherapy, ultimately leading to an unfavorable prognosis. This study, aPDL1-DTX/PFP@Lipid nanoparticles were synthesized and had an average diameter of 61.63 nm with 84.3% antibody modification. We demonstrated that the nanoparticles exhibited excellent PDAC-targeting capabilities both in vitro and in vivo. Upon exposure to low-intensity pulsed ultrasound (LIPUS) stimulation, the nanoparticles underwent a phase transition to form microbubbles with substantial molecular ultrasound diagnostic effects, and combined treatment resulted in a tumor growth inhibition rate of 88.91%. This treatment strategy also led to the infiltration of CD8+ T cells, the downregulation of Treg cells, the promotion of M1 macrophage polarization, the inhibition of fibrosis to reduce tumor stromal pressure, and the facilitation of perfluoropropane (PFP) gasification to release O2 and improve tumor hypoxia. In conclusion, aPD-L1-modified liquid‒vapor phase-transition nanoparticles loaded with docetaxel (DTX) were successfully combined with ultrasound for the molecular diagnosis and targeted treatment of PDAC. aPDL1-DTX/PFP@Lipid nanoparticles could reshape the PDAC TME, offering a new approach for ultrasound-mediated diagnosis and treatment with promising clinical applications.
Research Article
Targeted ultrasound-mediated molecular diagnosis and tumor microenvironment remodeling of pancreatic cancer with aPD-L1-modified docetaxel-loaded phase-transition nanoparticles
https://doi.org/10.21203/rs.3.rs-4806427/v1
This work is licensed under a CC BY 4.0 License
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Contrast-enhanced ultrasound
Molecular diagnosis
Targeted therapy
Tumor microenvironment
Pancreatic cancer
In recent years, the incidence of pancreatic cancer (PDAC) has been steadily increasing at a rate of 0.5–1% annually [1, 2]. PDAC is insidious at disease onset, challenging to diagnose in the early stages, and often is diagnosed in advanced stages, which results in a missed window for surgical resection. The tumor microenvironment (TME) is characterized by immune evasion, hypoxia, and high stromal pressure, which hinder drug delivery, resulting in poor responses to immunotherapy and chemotherapy [3]. Owing to the highly invasive nature and poor prognosis of PDAC, the 5-year survival rate of patients in China decreased from 11.7% from 2003–2005 to 7.2% from 2012–2015, which is the lowest among all cancer-related survival rates [4]. Hence, there is an urgent clinical need to address the current challenges in the clinical diagnosis and treatment of PDAC by developing a sensitive method that can detect PDAC at an early stage and achieve efficient drug delivery.
The depth of the pancreas and its location behind the peritoneum and the presence of gastrointestinal gas pose challenges during abdominal ultrasound examinations. Furthermore, other factors, such as examiner experience and skill, have historically restricted the clinical effectiveness of abdominal ultrasound. Despite the increased clinical use of contrast-enhanced ultrasound (CEUS) imaging techniques in recent years, the resolution of pancreatic imaging remains inadequate. CEUS is a dynamic, continuous process that lasts approximately three minutes, during which time gas interference can compromise the quality of contrast enhancement, leading to suboptimal diagnostic outcomes. Additionally, the average particle size of a microbubble contrast agent used in CEUS is approximately 2.5 µm [5], which hinders the ability of the contrast agent to traverse endothelial gaps, bind selectively to cancer cells, and consequently perform targeted tumor diagnosis [6].
Advances in molecular ultrasound imaging have opened new possibilities for the diagnosis and treatment of tumors [7]. The small size of liquid–vapor phase-transition nanoparticles allows them to easily penetrate tumor blood vessels and bind specifically to cancer cells. These nanoparticles (NPs) can be converted into microbubbles (MBs), which improve ultrasonography imaging quality by inducing a phase change through the application of heat or ultrasound irradiation. This noninvasive approach enables the visual diagnosis of tumor biological behaviors at the cellular and subcellular levels [8–10]. Upon triggering a phase change in the nanoparticles, the encapsulated perfluoropentane (PFP) is gasified to release O2, leading to an increase in reactive oxygen species (ROS) production in tumor cells [11]. Moreover, microbubble rupture by ultrasound-targeted microbubble destruction (UTMD) generates ultrasound-induced biological effects that increase the permeability of cancer cell membranes, aiding drug entry into cancer cells, improving drug delivery efficiency, and improving treatment outcomes [12–14].
In this study, we propose the construction of aPD-L1-modified liquid‒vapor phase-transition nanoparticles carrying docetaxel (aPDL1-DTX/PFP@Lipid) (Scheme 1). These small nanoparticles are designed to effectively target PDAC cells by penetrating the blood vessel wall. Upon exposure to low-intensity pulsed ultrasound (LIPUS), the nanoparticles undergo a phase change, enabling molecular diagnosis via ultrasonography. The physical effects induced by UTMD can directly harm cancer cells, while an anti-PD-L1 antibody (aPD-L1) and docetaxel (DTX) are simultaneously delivered for immunotherapy and chemotherapy. Moreover, the PFP transported by the nanoparticles acts as an oxygen reservoir to improve the hypoxic conditions in the PDAC TME. This comprehensive strategy alters the PDAC TME in multiple ways, resulting in integrated molecular ultrasound diagnosis and targeted therapy for PDAC.
Materials and animals
The following reagents were used in this study: 1,2-distearoyl-sn-glycero-3-phosphocholine (DSPC), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N- [methoxy(ethylene glycol)-2000] (DSPE-PEG2000) (AVT Pharmaceutical, China), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[carboxy(polyethylene glycol)-2000] (DSPE-PEG2000-COOH) (Xi'an Ruixi Biological Technology, China),
cholesterol (Sigma, USA), docetaxel (J&K Scientific, China), anti-mouse PD-L1 antibody (#HY-P99145, MedChemExpress, USA), rabbit anti-mouse CD8 antibody (#ab217344, Abcam, UK), Ki67 monoclonal antibody (SolA15) (Invitrogen, USA), rabbit anti-α-smooth muscle actin (α-SMA) antibody (#ET1607-53, HUABIO, China), pancytokeratin (Pan CK) monoclonal antibody Alexa Fluor™ 488 (#53-9003-82), donkey anti-rabbit IgG (H+L) highly cross-adsorbed secondary antibody Alexa Fluor™ 568 (#A10042), and donkey anti-mouse IgG (H+L) highly cross-adsorbed secondary antibody Alexa Fluor™ 647 (#A-31573) (Thermo Fisher, USA) were used.
Mouse PDAC cells (Pan02) were purchased from the National Experimental Cell Resource Sharing Platform (Beijing, China). C57BL/6 mice (4 weeks; 18‒22 g) were obtained from Silaike Experimental Animal Co., Limited Liability Company (Shanghai, China).
Synthesis of the targeted drug-loaded phase-transition nanoparticles
DSPC, DSPE-PEG2000, cholesterol, poloxamer, and DSPE-PEG2000-COOH were dissolved at a mass ratio of 20 mg:3 mg:1 mg:1.8 mg:2.5 mg in 2 mL of trichloromethane.
Five milligrams of docetaxel was dissolved in 1 mL of methanol and added to the above mixture. The solvent was then evaporated via vacuum rotary evaporation in a water bath at 50 °C for 30 min to form a phospholipid mixture film. The film was subsequently hydrated with 4 mL of MES buffer (0.1 M, pH 6) by ultrasonic dispersion at 300 W in a water bath at 50 °C to create a phospholipid suspension.
To obtain DTX-loaded lipid phase-transition nanoparticles (DTX/PFP@Lipid), 1 mL of the phospholipid suspension was mixed with 15 μL of PFP and emulsified via noncontact ultrasonication (XM08-II, Xiaomei Ultrasonic Instruments) for 10 min (1000 W, 30 s on/off) in a water bath at 3 °C.
The carbodiimide method was used to modify the nanoparticles. Briefly, a solution of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide/N-hydroxysuccinimide (EDC/NHS) in MES buffer (0.1 M, pH 6) was prepared and added to DTX/PFP@Lipid (-COOH:EDC:NHS= 1:10:30, molar ratio) for 2 h of reaction at 4 °C to activate the carboxyl groups, followed by ultrafiltration and centrifugation for 2 h (4 °C, 6000 rpm) to remove the buffer and excess EDC/NHS.
Finally, the activated DTX/PFP@Lipid mixture was resuspended in MES buffer (0.1 M, pH 8), and aPD-L1 was introduced at a DTX:aPD-L1 mass ratio of 6:1. After thorough mixing, the mixture was incubated at 4 °C with slow shaking for 2 h to obtain targeted aPD-L1-modified docetaxel-loaded liquid–vapor phase-transition nanoparticles (aPD-L1-DTX/PFP@Lipid).
Characterization of the NPs
A copper mesh with Formvar® film was coated by the dropwise addition of 20 μL of aPDL1-DTX/PFP@Lipid (diluted 1:500 in ultrapure water), and after 15 min of adsorption, the sample was negatively stained with 2% (W/V) phosphotungstic acid (pH 6.5) for 30 s. The internal structure and morphology of the nanoparticles were observed via transmission electron microscopy (TEM) (Tecnai G2, FEI). The particle size distribution, zeta potential, and polydispersity index (PDI) of the NPs were determined with a particle size analyzer (LitesizerTM 500, Anton Paar), and these measurements were repeated on days 1, 5, and 10 following preparation to assess the in vitro stability of the nanoparticles.
Encapsulation efficiency (EE) and drug loading capacity (LC) of the NPs
Liquid chromatography‒tandem mass spectrometry (LC‒MS/MS) (Triple Quad™ 4500, Applied Biosystems & LC‒30AC, Shimadzu) was used to determine the EE and LC of DTX in the drug-loaded phase-transition nanoparticles. The chromatographic column used was a Shim-pack GSP-HPLC C18 column (3 μm, 2.1 mm × 50 mm). Mobile phase A (aqueous phase) was an aqueous solution of 0.1% formic acid, while mobile phase B (organic phase) consisted of 0.1% formic acid in acetonitrile. Gradient elution was performed with a column temperature of 40 °C, an injector temperature of 8 °C, an injection volume of 4 μL, and a flow rate of 0.6 mL/min. Additionally, the electrospray ionization source was operated in positive ion mode via multiple reaction monitoring (MRM). The ion source settings were as follows: voltage, 5.5 kV; temperature, 500 °C; air curtain gas, 35 psi; spray gas, 50 psi; and auxiliary heating gas, 50 psi. The mass transition of DTX from 830.5→549.3 m/s was quantified with a declustering potential (DP) of 140 V and a collision energy (CE) of 60 V.
EE (%) = (DTX content in the NPs/total amount of DTX delivered) × 100%
LC (%) = (DTX content in the NPs/total mass of the NPs) × 100%
Determination of aPD-L1 modification efficiency
DiI-labeled aPDL1-DTX/PFP@Lipid was mixed with 1 mL of PBS and 2 μL of donkey anti-mouse IgG (H+L) Alexa Fluor Plus 488 (#A32766, Thermo Fisher) and incubated for 2 h at 4 °C. The mixture was then centrifuged at high speed (14,000 rpm, 4 °C) to eliminate any unbound secondary antibody. The sample was subsequently washed three times with PBS, after which the dual fluorescently labeled nanoparticles were suspended in ultrapure water. Fluorescence images were captured with a Leica DMi8 fluorescence microscope, and image colocalization was analyzed with Leica application suite X 3.5.7 (LAS X) software. The rate of aPD-L1 conjugation to DTX/PFP@Lipid was assessed by flow cytometry (BD Accuri C6 Plus), and quantitative analysis was performed with FlowJo 10.8.1 software.
Thermotropic capability of the liquid–vapor phase-transition nanoparticles
A nanoparticle solution was diluted to a concentration of 0.5 mg/mL with double-distilled water and placed in transparent flat-bottom glass vials. These vials were then heated in a water bath at 37 °C, 40 °C, 45 °C, 50 °C, 55 °C and 60 °C for 5 minutes at each temperature. The number and morphology of the phase-transition microbubbles were determined under an optical microscope (CKX41, Olympus). Three random fields of view were chosen from each group for image capture, and the numbers and diameters of the MBs were assessed by ImageJ for statistical analysis.
Acoustic droplet vaporization (ADV) and ultrasound imaging of the NPs in vitro
The aim of this study was to investigate the effects of thermal and LIPUS in vitro-triggered ADV on ultrasound imaging signal enhancement. First, 1 mL of aPDL1-DTX/PFP@Lipid diluted in double-distilled water was added to centrifuge tubes and heated in a water bath at temperatures ranging from 37 °C to 60 °C for 5 minutes at each temperature. B-mode and CEUS images of the nanoemulsions in the centrifuge tubes were acquired at various temperatures with a Canon i800 diagnostic ultrasound instrument equipped with a line array probe (model: i18LX5, center frequency: 12 MHz).
Subsequently, 1 mL of NPs (50 μg/mL) was added to a centrifuge tube to determine the effects of duration (1–5 min) and acoustic intensity (0.5, 1.0, 1.5, 2, 2.0, 2.5 W/cm2) on the ADV induced by LIPUS stimulation of the NPs in vitro. Ultrasound images of the nanoparticle emulsions in the centrifuge tubes were captured after each irradiation session with a Canon i800 diagnostic ultrasound device equipped with an i18LX5 line array probe. The CEUS signal intensity values and B-mode image grayscale values were quantified with the device’s integrated TCA software and ImageJ.
Confirmation of PD-L1 expression in Pan02 cells
Pan02 cells were seeded at a density of 1×105 cells per well in 96-well plates and cultured at 37 °C in a humidified incubator with 5% CO2. Upon reaching 60% confluence, the culture medium was aspirated, and the cells were fixed with 4% paraformaldehyde at room temperature. After being rinsed three times with phosphate-buffered saline (PBS), the cells were blocked with bovine serum albumin (BSA) for 30 minutes. A rabbit anti-mouse PD-L1 antibody (#ab213480, Abcam, 1:500) was subsequently added, and the samples were incubated for 1 h at 37 °C, followed by three additional washes with PBS. Furthermore, goat anti-rabbit IgG-FITC (ab6717, Abcam, 1:500) was applied, and the samples were incubated for 30 min at 37 °C in the dark prior to washing three times with PBS. The cells were then stained with DAPI staining solution, incubated for 10 min at room temperature in darkness, and washed with PBS, after which the fluorescence of FITC on the surface of the Pan02 cells was visualized with a fluorescence microscope.
Pan02 cells in the logarithmic growth stage were harvested, fixed with 4% paraformaldehyde for 15 minutes, and washed three times with PBS. After the cell density was adjusted to 2×106/mL, 500 μL of the cell suspension was incubated with 2 μL of anti-mouse PD-L1 antibody at 37 °C for 1 h in the dark, followed by centrifugation at 1000 rpm for 5 min and resuspension of the cells in 500 μL of PBS. Next, goat anti-rabbit IgG-FITC was added, and the cells were incubated on a shaker at 37 °C for 30 min. After an additional centrifugation step and three washes, the Pan02 cells were resuspended in 500 μL of PBS and analyzed by flow cytometry.
Targeting efficiency of aPDL1-DTX/PFP@Lipid in vitro
Pan02 cells were placed in 96-well plates at predetermined concentrations and incubated for 24 h, after which the medium was discarded. Serum-free DMEM was then added for an additional 4 h of culture. Next, the cells were divided into three groups: nontargeting, targeting, and antagonist. In the antagonist group, an excess of anti-mouse PD-L1 antibody was introduced 30 min before the end of the starvation culture. Next, 10 μL of DiI-labeled DTX/PFP@Lipid or aPDL1-DTX/PFP@Lipid (2 mg/mL) was added to each group after the starvation culture was complete, and the cells were incubated for an additional 2 h. The culture medium was then withdrawn, any unbound nanoparticles were removed by thorough washing with PBS, and the cells were fixed in 4% paraformaldehyde and stained with DAPI solution. Finally, the cells were washed with PBS three times and observed under a fluorescence microscope.
NP uptake by Pan02 cells in vitro
Pan02 cells in the logarithmic growth phase were seeded at a density of 1×105 cells per well in 96-well plates and allowed to adhere. Subsequently, 10 μL of DiI-labeled aPDL1-DTX/PFP@Lipid was added for coculture in the dark for 3, 6, 9, or 12 h. The cells were then rinsed with PBS, fixed with 4% paraformaldehyde, stained with DAPI solution, and observed under a fluorescence microscope.
Similarly, Pan02 cells were seeded in 12-well plates and cultured until they reached confluence. The cells were then cocultured with 100 μL of DiI-labeled aPDL1-DTX/PFP@Lipid for various durations (0, 3, 6, 9, or 12 h). The culture medium containing the nanoparticles was removed, the cells were washed three times with PBS, and cell pellets were obtained following trypsin digestion and centrifugation. After three additional washes with PBS, the cell concentration was adjusted, and the cells were analyzed by flow cytometry.
Detection of ROS production and NPs cytotoxicity
Pan02 cells were seeded in 48-well plates and divided into four groups: control, NPs, LIPUS, and NPs+LIPUS. The NPs and NPs+LIPUS groups were treated with 10 μL of aPDL1-DTX/PFP@Lipid for 6 h after cell attachment and ultrasonic irradiation (2.5 W/cm2, 3 min), followed by the LIPUS and NPs+LIPUS groups. The cells were subsequently cultured for an additional 24 h. The intracellular ROS levels were detected with an ROS fluorescence assay kit (#E-BC-K138-F, Elabscience), and the cell nuclei were labeled with Hoechst 33342 (#62249, Thermo Scientific).
The cytotoxicity of the NPs was assessed via CCK-8 assays. The cells were seeded at a density of 1 × 105 cells per well in 96-well plates and divided into the following groups: free DTX, PFP@Lipid, DTX/PFP@Lipid, and aPDL1-DTX/PFP@Lipid (n=3). Different concentrations of DTX (1.25 μg/mL, 2.5 μg/mL, 6.25 μg/mL, 12.5 μg/mL, and 25 μg/mL) were evaluated, with PFP@Lipid without DTX or aPD-L1 serving as control nanoparticles. Each group was further divided into ultrasonication-irradiation and no ultrasonication subgroups. Following 6 h of incubation after drug addition, the ultrasonication-irradiation subgroups were subjected to LIPUS irradiation (2.5 W/cm2, 3 min) and incubated for an additional 24 h. The culture medium was subsequently aspirated, the cells were washed with PBS, and fresh medium containing 10% CCK-8 was added to the wells for an additional 0.5–1 h of incubation. The OD at 450 nm was measured with a multifunctional enzyme reader (SpectraMax i3X, Molecular Devices).
Targeted NPs biodistribution in vivo
A suspension of Pan02 cells in the logarithmic growth phase was combined with an equal proportion of cell matrix gel (#354234, BD BioCoat) under cold conditions. Then, 0.2 mL (1 × 107 cells) of the suspension was injected into the right inguinal subcutis of each C57BL/6 mouse. The tumor was considered ready for the experiment once it reached a diameter of 1 cm. Prior to the experiment, the tumor-bearing mice were shaved such that the abdomen and tumor site were completely exposed. The mice were subsequently randomly assigned to two groups: the nontargeting group and the aPD-L1-targeting group (n=5). DiR-labeled NPs (200 μL) were injected into the mice in both groups via the tail vein, and images were captured from the mice under continuous isoflurane anesthesia at specific time points (2 h, 6 h, 12 h, 24 h, 48 h, 96 h, and 192 h postinjection) with a small animal in vivo fluorescence imaging system (IVIS® Spectrum, Caliper Life Sciences). The excitation and emission wavelengths used were 740 nm and 780 nm, respectively. The fluorescence signal intensity was quantitatively assessed with Living Image.
Furthermore, the tumor-bearing mice were euthanized at 6 h or 24 h after intravenous injection of the NPs. The major organs, including the heart, liver, spleen, lungs, kidneys, and tumors, were subsequently isolated for fluorescence imaging to assess the distribution of the fluorescence signal.
Phase transition of the NPs in vivo and ultrasound imaging
Nine tumor-bearing mice with tumors measuring approximately 1 cm in diameter were selected for the experiment. The mice were intravenously injected with 200 μL of DTX/PFP@Lipid, aPDL1-DTX/PFP@Lipid, or PBS (control group) (n=3). At 6 h or 24 h postinjection, the mice were anesthetized with 400 mg/kg tribromoethanol via intraperitoneal administration. The tumor site was subsequently exposed to LIPUS irradiation (2.5 W/cm2, 3 min). Ultrasound images were captured with a Canon i800 diagnostic ultrasound instrument with an i18LX5 line array probe at the following time points: preinjection and 6 h postinjection + LIPUS and 24 h postinjection + LIPUS, after which the B-mode and CEUS image signal intensities were analyzed.
Evaluation of the in vivo antitumor efficacy of combined NP and LIPUS treatment
Pan02 cells were inoculated into the right abdominal subcutis of the mice. When the tumor volume reached approximately 60 mm3, the mice were randomly divided into 7 groups (n=5): model, ultrasound irradiation alone, free DTX, free aPD-L1, DTX/PFP@Lipid, aPDL1-DTX/PFP@Lipid, DTX/PFP@Lipid+LIPUS, and aPDL1-DTX/PFP@Lipid+LIPUS. The NPs or free drugs were injected at the same dose (DTX: 30 mg/kg; aPD-L1: 5 mg/kg) via the tail vein. Twenty-four hours after intravenous drug administration, the tumor site was irradiated with LIPUS (2.5 W/cm2, 5 min). The treatment was repeated every 5 days for a total of 3 times. Tumor growth was monitored regularly. After 1 week of observation following the last treatment, the mice were euthanized by cervical dislocation, and the tumors were removed and weighed to calculate the relative rate of tumor growth inhibition, as follows.
Histopathological and immunohistochemical analyses of tumor tissues
Each tumor sample was fixed in tissue fixative, dehydrated, embedded in paraffin, and then cut into serial sections (4 μm thick). These sections were subjected to hematoxylin and eosin (H&E) and immunofluorescence chemical staining to detect the expression of Pan CK, nuclear proliferation-associated antigen (Ki67), and α-SMA and CD8+ T-cell infiltration. Additionally, immunohistochemical analysis was performed to assess FoxP3, CD206, and CD86 expression.
Statistical analyses
Statistical analysis was conducted via GraphPad Prism 9.5.1 software. The data are presented as the means ± standard deviations (SDs). Comparisons between two groups were performed via an independent samples t test, whereas one-way ANOVA was used for comparisons among more than two groups. Statistical significance was considered at P < 0.05, with levels of significance denoted as follows: no significance (NS), *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.
Characterization of the NPs
The aPDL1-DTX/PFP@Lipid had a size of 61.63 ± 0.55 nm with a narrow, positively skewed distribution, a PDI of 0.24 ± 0.01, and a zeta potential of -22.63 ± 0.87 mV (Fig. 1A). TEM revealed that these nanoparticles had a core‒shell structure with a spheroidal shape, as reflected by their uniform size distribution (Fig. 1B). Over 5 and 10 days of in vitro observation, there was a marginal decrease in particle size and a slight increase in the absolute value of the zeta potential, although these differences did not reach statistical significance (P > 0.05) (Fig. 1C-D). The EE and LC of DTX were determined to be 91.9% and 14.8%, respectively.
In vitro phase transition capability of the NPs
The appearance of the nanoemulsion changed from transparent to milky white as the water bath temperature increased. A gradual increase in the number of phase-transition microbubbles was observed under a microscope (Fig. 2A), with a marked increase observed at 55°C (Fig. 2B). This led to a phase transition in which the MB size distribution changed from sparse and uneven to small, dense, and uniform, with a diameter of approximately 7.32 ± 3.72 µm (Fig. 2C). After the phase change occurred, the volume increased by 120-fold compared with that of the nanoparticles before the phase transition. The ultrasound echo signal intensity of the phase-transition nanoemulsion exhibited a temperature dependence, peaking at 55°C (Fig. 2D-E) and increasing 125-fold compared with that before the phase change. After the ultrasound-triggered phase change, the echo signal intensity of the nanoemulsion increased with increasing ultrasound irradiation intensity and time and reached the maximum intensity after 3 minutes of ultrasound irradiation at 2.5 W/cm2 (Fig. 2F-G).
In vitro NPs targeting and uptake
PD-L1 expression (green fluorescence) was detected in Pan02 cells via fluorescence microscopy (Fig. 3A). Flow cytometry revealed that PD-L1 expression in Pan02 cells was approximately 96.5% (Fig. 3B). Moreover, PDAC cells exhibited significant binding to DiI-labeled aPDL1-DTX/PFP@Lipid NPs, as indicated by the red fluorescence surrounding the cells. However, NP binding was reduced in the presence of free aPD-L1 antibody (Fig. 3C), and semiquantitative analysis of the fluorescence intensity revealed that the difference was significant (P < 0.001).
Furthermore, fluorescence microscopy (Fig. 4A) revealed that the cellular uptake of the nanoparticles (red fluorescence) was dependent on both intensity and time. Significant differences in the increase in fluorescence intensity were observed at the 3 h, 6 h, and 9 h time points (P < 0.05) (Fig. 4B). Flow cytometry analysis (Fig. 4C) confirmed that the cellular uptake efficiency of the aPDL1-DTX/PFP@Lipid NPs increased over time, with uptake rates of 15.4%, 76.1%, 91.7%, and 97.5% at 3 h, 6 h, 9 h, and 12 h, respectively.
ROS production and NP cytotoxicity
Following combination treatment with the NPs and LIPUS irradiation, a substantial increase in the ROS signals was observed in the tumor cells (Fig. 5A). The CCK-8 results revealed a decrease in overall cell viability as the concentration of DTX or nanoparticles increased (P < 0.0001). Intergroup analysis revealed that the free DTX group presented the lowest cell viability (P < 0.001) (Fig. 5B). After ultrasound irradiation, a significant decrease in cell viability was observed in the drug-loaded nanoparticle group, with the aPDL1-DTX/PFP@Lipid group showing the most prominent effect. At an NP concentration of 25 µg/mL, the cell viability decreased to 31.14%, indicating substantial cytotoxicity (P < 0.05) (Fig. 5C).
In vivo biodistribution and metabolism of the NPs
In the targeted nanoparticle group, the fluorescence signal intensity at the tumor site clearly and continuously increased within the first 12 h, peaked at 24 h, and then plateaued until 48 h. Clearance of the signal and a reduction in its intensity were observed at 96 h. Throughout the observation period, the fluorescence intensity in the targeted group at various time points significantly exceeded that in the nontargeted group (P < 0.001). Even after 196 h, the average fluorescence signal in the targeted group remained 1.85 times greater than that in the nontargeted group (Fig. 6A, 6C).
At 6 h and 24 h after intravenous nanoparticle injection, three mice per group were euthanized for analysis. The results showed that at 6 h, both groups presented the highest fluorescence signals in the liver and spleen, followed by the kidneys. Moreover, the tumor signal intensity in the targeted group was significantly greater than that in the nontargeted group (P < 0.01). By 24 h, the tumor signal intensity in the targeted group had continued to increase significantly compared with that in the nontargeted group (P < 0.001). Interestingly, analysis revealed that the fluorescence signal intensity per unit area in the tumor was significantly greater than that in the spleen (P < 0.001), which was a smaller difference than that in the liver (P < 0.01) (Fig. 6B, 6D).
In vivo phase transition and ultrasound imaging of the NPs
LIPUS irradiation was performed at the tumor site at 6 h and 24 h after material injection. In the targeted group, the ultrasonic echo signal was greater at 24 h than at 6 h, and both the grayscale and CEUS signals were significantly greater in the targeted group than in the nontargeted group (P < 0.05) (Fig. 7).
Antitumor effects of the nanoparticles in vivo
With continued treatment, the tumor volume in the model group of mice continued to increase, whereas the aPDL1-DTX/PFP@Lipid + LIPUS group exhibited a significant trend toward tumor growth inhibition. By the 14th day, the tumor growth inhibition rate reached 88.91% in the aPDL1-DTX/PFP@Lipid + LIPUS group (Fig. 8).
Pathological examination
H&E staining (Fig. 9A) revealed that tumor cell nuclei in the control group were deeply stained, varied in size, and displayed features of nuclear division. Conversely, in the aPDL1-DTX/PFP@Lipid + LIPUS group, most tumor cell nuclei appeared condensed and necrotic, indicating potent tumor cell killing effects. Confocal laser microscopy revealed positive Pan CK expression in the epithelial cells of the PDAC tissues, which colocalized with the DAPI-labeled cell nuclei. Furthermore, there was a significant increase in CD8 + T-cell infiltration in the aPDL1-DTX/PFP@Lipid + LIPUS group (P < 0.0001). The expression of the antigen Ki67, which is associated with tumor cell proliferation, and α-SMA, which is linked to cancer-related fibroblasts, was markedly reduced in the aPDL1-DTX/PFP@Lipid + LIPUS group (P < 0.0001) (Fig. 9B-D). Immunohistochemical analysis revealed a decrease in FoxP3 and CD206 expression, whereas CD86 expression increased in the aPDL1-DTX/PFP@Lipid + LIPUS group (Fig. 10).
Microbubble ultrasound contrast agents can resonate with ultrasound waves, resulting in strong backscattering and significantly amplified echoes [15, 16]. This technology has been widely utilized for clinical disease diagnosis, marking a significant advancement in ultrasound imaging, and is frequently considered the third revolution in this field. However, because the average particle size of microbubble ultrasound contrast agents currently used in clinical practice is 2.5 µm, they cannot pass through the approximately 150 nm inner diameter of endothelial gaps and extravasate from blood vessels [17, 18], which prevents their development for use in tumor-targeted molecular diagnosis. The use of liquid–vapor phase-transition nanoparticles represents a novel approach for further molecular ultrasound imaging of tumors. NPs are not easily visualized in B-mode or by CEUS imaging, but they can undergo a phase transition to form microbubbles upon the application of ultrasound, which results in noticeable echo enhancement [19]. These smaller nanoparticles can then pass through gaps in vascular endothelial cells and, if they are endowed with specific modifications, can directly attach to tumor cells. Moreover, when nanoparticles undergo a phase change upon heating or applying ultrasound irradiation, the formed ultrasonography-visible microbubbles can be utilized for molecular tumor diagnosis.
The phase-transition ability and targeted imaging effect of liquid–vapor phase-transition nanoparticles have been a focus of research. Many nanoparticles mentioned in the literature have relatively large particle sizes (ranging from 100–500 nm) [20–22], limited targeting capabilities, and suboptimal phase-transition properties, which leads to low echo signal intensities in contrast ultrasound. To increase the contrast ultrasound echo intensity in specific targeted tissues and achieve true molecular ultrasound diagnosis, it is essential to focus on improving the selection of nanoparticle materials, refining the preparation methods, controlling the particle size, and implementing targeted modifications.
Phospholipids are commonly selected as shell materials for lipid nanoparticles because of their favorable elasticity and ductility [23]. In this research, synthetic phospholipids were utilized for the shell, and cholesterol was added in an appropriate proportion to increase the rigidity and stability of the nanoparticles. The core of the nanoparticles contained a liquid fluorocarbon, which is crucial for achieving phase transition [24]. PFP, with a boiling point of 29°C, transitions from liquid to gas at the body's normal temperature of 36–37°C. However, the actual boiling point of PFP within the nanoparticles is greater because of the Laplace pressure [25, 26]. The Laplace pressure is inversely related to the radius of the nanoparticles [27, 28], meaning that smaller nanoparticles experience higher Laplace pressures, resulting in nanoparticles with a more stable structure. The nanoparticles produced in this study have a diameter of approximately 60 nm and a phase change temperature of 50–55°C, which is significantly higher than the normal temperature of the human body. These features improve the stability of the nanoparticles upon intravenous injection into the bloodstream, reducing the risk of gas embolism caused by spontaneous phase changes in the nanoparticles within the circulation.
The nanoparticles were prepared by thin film hydration and ultrasonic emulsification [11, 21, 29, 30]. Traditional ultrasonic emulsification typically involves the use of instruments such as an ultrasonic cell crusher [9, 20], which requires the insertion of a cone-shaped amplitude rod into the solution, causing the solution container to seal properly. Moreover, the high energy at the head end of the amplitude rod results in an uneven distribution of ultrasonic energy, causing a premature phase change of the liquid fluorocarbon during nanoparticle preparation and leading to variations in particle size. To address these issues, a noncontact ultrasonic crusher was employed in this study.
Phospholipids and PFP were enclosed in a centrifuge tube, and ultrasonic waves were then applied from the outside to the inside of the tube, ensuring uniform energy distribution. The closed space generates a specific pressure, which prevents the spontaneous phase change of PFP [31]. Consequently, the prepared nanoparticles contained an adequate amount of PFP that was evenly distributed. The size of the gap between cancer blood vessels has been estimated to be 100–600 nm. By adjusting the ultrasound energy and irradiation mode, nanoparticles with a particle size of less than 100 nm were obtained that could easily pass through blood vessel endothelial gaps, which ensures their good stability [32].
Currently, the two primary methods widely used for droplet phase change are heating and ADV [26, 33]. ADV utilizes ultrasonic irradiation, which offers a high penetration depth and controllable energy, increasing its practicality [34]. ADV-mediated nanoparticle vaporization is influenced by various factors. Lipid shells exhibit excellent mechanical elasticity, allowing for repeated expansion and contraction [35], along with high biocompatibility, good loading capacity, and controllable biological properties [36, 37]. The energy required to reach the vaporization threshold of phase-transition nanoparticles increases proportionally with the length of the lipid acyl chain, and the lipid composition modulates the acoustic characteristics of the nanodroplets [14, 38]. On the other hand, cholesterol maintains relatively stable biofilm fluidity across various temperatures without significantly impacting the phase change temperature, thus increasing the stiffness of the phase-transition nanoparticles.
When the incident ultrasound frequency matches the natural oscillation frequency of the microbubble, resonance occurs, resulting in the highest acoustic energy output and optimal CUES performance. The resonant frequency is inversely correlated with the microbubble diameter; for example, a 3 µm microbubble resonates at 2.4 MHz, whereas a 5 µm microbubble resonates at 1.3 MHz. The conversion of PFP from a liquid to a gas causes a volume expansion of approximately 125 times. If the particle size of a prepared nanoparticle exceeds 200 nm, its extravasation from blood vessels for effective targeting and binding to tumor cells may be hindered. On the other hand, after phase transition, the microbubble particle size can reach 25 µm, exceeding the resonant frequency of diagnostic ultrasound commonly used in clinical settings and potentially compromising the efficacy of ultrasound imaging. In the experiments here, the nanoparticle transition to microbubbles led to a 120-fold expansion in vitro to approximately 7 µm. In vivo, considering constraints such as interstitial pressure, the size of the microbubbles was controlled and closely mimicked that of traditional ultrasound contrast agents. This enhancement facilitates an up to 125-fold increase in echo intensity for effective nanoparticle-mediated contrast-enhanced molecular ultrasound diagnosis.
Following intravenous injection, the nanoparticles were readily cleared by the reticuloendothelial system in vivo. The aim of this study was to reduce uptake by reticuloendothelial cells in vivo by adding PEG to the nanoparticle shell and controlling the nanoparticle size [30]. Additionally, an aPD-L1 antibody was conjugated to the nanoparticle surface via carbodiimide chemistry, enabling targeted binding to PDAC cells expressing PD-L1. Fluorescence imaging analysis demonstrated enhanced binding of the nanoparticles to PDAC cells, particularly those at the tumor site.
The unique and complex immune TME of PDAC contributes to unfavorable clinical treatment outcomes and prognosis [39]. This research demonstrated that intravenous nanoparticle injection led to significant antitumor effects in mice with PDAC. This treatment regimen resulted in a significant decrease in α-SMA expression, which is linked to cancer-associated fibroblasts and fibroplasia inhibition in cancerous tissues. The PD-L1 antibody-modified nanoparticles not only displayed targeting effects but also blocked the PD-L1/PD-1 pathway in PDAC, thus reducing immune suppression [40, 41]. Following nanoparticle administration, there are fewer Treg cells (FoxP3+) and M2 macrophages (CD206+) in PDAC tissues, along with more CD8 + T cells and M2 macrophages (CD86+) at the tumor site, which remodel the PDAC immune microenvironment to a certain extent [13, 39, 42]. Additionally, the release of oxygen from the PFP core of the nanoparticles improved the hypoxic conditions in the PDAC microenvironment [43].
liquid–vapor phase-transition NPs have been shown to bind specifically to PDAC cells, and these NPs experience transient changes in size during ADV; this process generates shear forces that can increase the permeability of the cell membrane [44]. Additionally, the instantaneous energy released when microbubbles burst leads to acoustic cavitation effects, further disrupting the cell membrane and allowing for increased drug entry into the cells [45]. DTX has poor water solubility and high systemic toxicity when it is administered systemically [46]. However, by encapsulating DTX in targeted nanobubbles and applying UTMD, these limitations can be overcome, ultimately enhancing the antitumor effects of DTX.
In this study, targeted drug-loaded liquid–vapor phase-transition nanoparticles, aPDL1-DTX/PFP@Lipid NPs, which possess an optimal particle size that enables them to penetrate vascular endothelial cell gaps and bind specifically to PDAC cells, were developed. Upon ultrasound irradiation, these NPs undergo a phase change, transforming into microbubbles that exhibit strong diagnostic ultrasound resonance to significantly enhance the ultrasound imaging efficacy and enable targeted molecular ultrasound diagnosis of PDAC. Furthermore, the biological effects induced by the ultrasound-mediated disruption of the microbubbles increased the permeability of the PDAC cell membranes, facilitating drug penetration into the cells. The combination of UTMD, PD-L1 antibodies and DTX effectively inhibits PDAC cells and reshapes the TME through mechanisms such as suppressing fibrous tissue proliferation, reversing immune suppression, and ameliorating hypoxia. This approach allows the precise targeted diagnosis and treatment of PDAC through molecular ultrasound imaging and shows considerable potential for clinical application.
PDAC: Pancreatic ductal adenocarcinoma; TME: tumor microenvironment; CEUS: contrast-enhanced ultrasound; NPs: nanoparticles; MBs: microbubbles; PFP: perfluoropentane; ROS: reactive oxygen species; UTMD: ultrasound-targeted microbubble destruction; aPDL1-DTX/PFP@Lipid: aPD-L1-modified liquid‒vapor phase-transition nanoparticles carrying docetaxel; LIPUS: low-intensity pulsed ultrasound; aPD-L1: anti-PD-L1 antibody; DTX: docetaxel; DSPC: 1,2-distearoyl-sn-glycero-3-phosphocholine; DSPE-PEG2000: 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N‒ [methoxy(ethylene glycol)-2000]; DSPE-PEG2000-COOH: 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[carboxy(polyethylene glycol)-2000]; EDC: ethyl-3-(3-dimethyl-aminopropyl) carbodiimide; NHS: N-hydroxysuccinimide; DTX/PFP@Lipid:
DTX-loaded lipid phase-transition nanoparticles; EE: encapsulation efficiency; LC: drug loading capacity; ADV: acoustic droplet vaporization; PBS: phosphate-buffered saline.
Ethics approval and consent to participate
Animal Care was in accordance with institution guidelines. All animal studies were approved by the Animal Ethical Committee of Fujian Medical University (grant No. IACUC FJMU 2022-0721).
Consent for publication
Not applicable.
Data Availability
The datasets used and/or analyzed during current study are available from corresponding author on reasonable request.
Competing interests
The authors declare that they have no competing interests.
Funding
This research was supported by Joint Funds for the Innovation of Science and Technology, Fujian Province (Grant number: 2019Y9066).
Authors' contributions
Zhikui Chen involved in the design, planning, experimental guidance, funding support, and final decision-making of the entire research；Yi Tang and Qingling Shen directly involved in research design, experimental progress, and paper writing；Peng Lin provided experimental design direction and revised the manuscript； Minling Zhuo analyzed the data and drawn the schematic diagramt；Yajiao Gan wrote the manuscript；Yixi Su assisted with data search and organizing；Qingfu Qian provided technical assistance in material construction；Liwu Lin participated in project design guidance and paper revision； Ensheng Xue provide financial support for publishing the paper; All authors reviewed and edited the manuscript.
Acknowledgments
We thank the members of the Fujian Institute of Ultrasonic Medicine for their helpful comments and suggestions. The authors thank the Public Technology Service Center Fujian Medical University and the Central Laboratory of Union Hospital affiliated with Fujian Medical University for their help and technical support.
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Scheme 1 is available in the Supplementary Files section.
No competing interests reported.