Self‐Assembly of Stimuli‐Responsive Peptide Enhances Therapeutics by Specifically Disrupting Hepatocellular Carcinoma Lysosomes In Vivo

ABSTRACT

Lysosome sequestration or drug‐triggered autophagic flux curtails antitumor drug potency in hepatocellular carcinoma (HCC) and can potentially be reversed with tumor cell‐specific lysosomal disruption. Here, we demonstrate that a chimeric peptide (RS‐FS), consisting of HCC‐targeting RS and nanostructure‐forming motifs (FS), self‐assembles into nanospheres at neutral pH and transforms into nanofibers under acidic and reductive conditions. These nanofibers specifically localize to tumors and disrupt tumor cell lysosomes, thus enhancing doxorubicin's activity in human HCC cells in vitro and orthotopic HCC mice in vivo after RS‐FS‐doxorubicin treatment. Importantly, intravenous RS‐FS potentiated oral Lenvatinib's antitumor activity up to 61‐fold, and eradicated tumors in orthotopic HCC mice via HCC cell‐specific lysosome disruption. Potent antitumor effects were also achieved with intravenous RS‐FS and oral Epimedium brevicornu Maxim. ‐derived extracellular vesicles in orthotopic HCC mice, with markedly reduced tumor growth and increased cytotoxic T infiltration, in which RS‐FS‐mediated lysosome disruption promoted drug release and autophagic flux blockade. Our study demonstrates that RS‐FS self‐assembles into nanospheres or nanofibers in response to stimuli and enables tumor cell‐specific lysosome disruption, resulting in enhanced drug release, autophagic flux blockade, and antitumor activities of diverse therapeutics in HCC mice, and thus provides a generalizable peptide adjuvant for sensitizing HCC‐targeted therapeutics.

A short linear peptide (RS‐FS) with HCC‐targeting, nanosphere‐ and nanofiber‐forming motif self‐assembles into nanospheres in circulation and transforms to nanofibers to specifically disrupt tumor lysosomes, releasing antitumor drugs and blocking autophagic flux to augment antitumor potency with minimal side effects in vitro or in vivo. RS‐FS can enhance antitumor drugs with or without co‐administration, which improves clinical deployability.

1. Introduction

Hepatocellular carcinoma (HCC) remains an increasing global burden, particularly in the Asia‐Pacific region [1]. Current clinical antitumor therapeutic agents such as atezolizumab, bevacizumab [2, 3], and multi‐targeted receptor tyrosine kinase inhibitor Lenvatinib have improved HCC prognosis; however, the response rate and progression‐free survival can still be improved [4, 5]. In particular, Lenvatinib stimulates autophagic flux via lysosome fusion with autophagosomes, thus compromising its therapeutic efficacy [6]. The efficacy of other chemotherapeutics, such as sorafenib, regorafenib, and doxorubicin [7, 8] or nanomedicines [9, 10], is also attenuated by lysosome sequestration or both lysosome sequestration and drug‐triggered autophagic flux. Thus, overcoming the tumor cell lysosome‐related hurdles specifically would improve the potency of antitumor therapeutics.

Recently, self‐assembled peptide‐based functional nanomaterials (SAPs) have gained popularity as antitumor therapeutics or delivery vehicles given the ease of synthesis, tunability, and good biocompatibility [11, 12]. Although SAPs have been extensively used for enhancing antitumor therapeutics, including biotherapy, photodynamic, and photothermal, as well as bioimaging [11, 12, 13], the production commonly depends on co‐assembling of drugs/ prodrugs or imaging agents with peptide vectors in vitro, which involves complex encapsulation or covalent conjugation process with detailed characterization required [13] and co‐assembly will alter the clinically‐approved administration protocol of drugs. Thus, a generalizable peptide vector that can augment therapeutics’ antitumor efficacy without complexation with antitumor drugs would be desirable.

To accomplish this, in this study, we combined our previously identified proprietary HCC‐targeting peptide (RS) [14], and a stimuli‐responsive fiber‐forming hexapeptide motif (VAELYL) derived from natural insulin protein (F) that can assemble into nanofibers in lysosomes to disrupt lysosomes to improve antitumor drug release [15, 16, 17, 18], and 5 valine hydrophobic tail (VVVVV) that facilitates the formation of nanospheres via hydrophobic interaction (S) [19]. Given the higher stability and potency of the D‐ than L‐ enantiomeric form of F as reported previously [15], we adopted the D‐amino acid for the F peptide in our peptide design. We hypothesized that RS provides tumor‐targeting specificity, hydrophobic S promotes peptide nanosphere formation in a neutral environment, whereas F enables nanospheres to transform into nanofibers in response to the acidic and reductive environment in tumor cell lysosomes, thus triggering lysosome disruption. Intravenous RS‐FS enabled effective tumor suppression in subcutaneous and orthotopic HCC mice when loaded with doxorubicin or simultaneously administered with oral Lenvatinib or Epimedium brevicornu Maxim. ‐derived extracellular vesicles (EEV). Our study demonstrates that RS‐FS potentiates therapeutics’ antitumor efficacy in HCC mice by promoting tumor cell‐specific lysosome disruption and subsequent drug release, thus providing a generalized tumor cell lysosome‐targeted peptide tool for antitumor therapy.

2. Results

2.1. Self‐Assembly of Stimuli‐Responsive RS‐FS Enables Human HCC Cell‐Specific Lysosome Disruption

We designed a generalized peptide vector (RS‐FS) that enabled tumor cell‐specific lysosome damage as RS‐FS nanospheres would transform to lysosome‐disrupting nanofibers in response to environmental stimuli (Figure 1a). RS‐FS self‐assembled into nanospheres with an average diameter of 147 nm under an oxidative environment at neutral pH, a condition simulating physiological bloodstream condition [20], and maintained the nanosphere morphology at pH6.5, simulating the mildly acidic tumor microenvironment [20], visualized by transmission electron microscopy (TEM) and nanoparticle tracking analysis (NTA) (Figure 1b,c). In contrast, RS‐FS appeared as heterogeneous vesicles at pH6.5 with reducing 10 mm glutathione (GSH), reflective of early endosome [20], and transitioned to nanofibers (Figure 1b,c) in response to acidic and reductive environment (pH5.0 with 10 mm GSH), a condition similar to lysosome [20]. Notably, RS‐FS without oxidation failed to maintain the morphologies and appeared as heterogeneous vesicles at pH6.5 with or without 10 mm GSH, though it transformed into nanofibers in a disorderly manner at pH5 with 10 mm GSH (Figure 1b,c). A sharp doxorubicin fluorescence increase observed in the solution of pH6.5 with GSH compared to pH7.4 and pH6.5, peaking at pH5 with GSH for RS‐FS‐doxorubicin under oxidative conditions, showed that the doxorubicin is only released in endosomal/lysosomal compartments (Figure S1a). Doxorubicin co‐assembled with RS‐FS without oxidation exhibited a substantial fluorescence increase at pH6.5 without GSH (Figure S1a). These results showed that oxidation is important for targeted intracellular release. Although RS‐S also assembled into nanospheres under pH7.4 and 6.5, it failed to form nanofibers under pH5 and a reductive environment (Figure 1b,c), confirming that the full RS‐FS peptide is required for functionality. Circular Dichroism (CD) spectra analysis revealed the conformational transition from predominantly random coils and weak α‐helixes at pH7.4, 6.5, and 6.5 with 10 mm GSH to β sheets at pH5 with 10 mm GSH conditions, reflected by the red shift of the positive peak and the concomitant blue shift of the negative extremum (Figure 1d). Relatively low critical micelle concentration (CMC) with negative zeta potential at pH7.4 indicated the relative stability of RS‐FS in circulation (Figure S1b). To determine the critical gelation concentration (CGC) under pH5 with 10 mm GSH, we mixed RS‐FS with eosin‐Y dye [21] to visualize the gelation process, and observed morphological changes at different concentrations of RS‐FS, with hydrogel formation appearing at 1 mg/mL, supported by TEM showing the phase transition from nanofiber to intertwined network (Figure 1e), indicating that the CGC for RS‐FS is 1 mg/mL. Consistently, rheological results showed the storage modulus (G′) remained almost the same as the loss modulus (G″) for RS‐FS under pH7.4, 6.5, and 6.5 with 10 mm GSH (below 30 Pa), whereas the value of G′ rose to 600 Pa, significantly greater than G″ under pH5 with 10 mm GSH (Figure 1f), showing the formation of stable hydrogel under an acidic and reductive environment. Hydrogel formation delayed diffusion of lysosome‐labeling LysoTracker dye in RS‐targeted human HCC LM3 cells treated with RS‐FS but not in normal human liver 7702 cells after fluorescence recovery after photobleaching (FRAP) assay [22] (Figure 1g), verified by dense amyloid‐like fibers inside enlarged lysosomes of LM3 cells and cytosolic release of cathepsin B triggered by lysosomal membrane permeabilization (LMP) [23] (Figure 1h,i), demonstrating that RS‐FS forms a hydrogel in HCC cells specifically. To further validate tumor‐specific lysosome disruption by RS‐FS in vivo, we intravenously injected Cy5‐labeled RS‐FS and a scrambled‐FS, in which the RS sequence was randomized, into day‐14 subcutaneous HCC mice. A much higher tumor/liver ratio was achieved with RS‐FS (4.73) than scrambled‐FS (1.08) (Figure S1c), confirming the tumor selectivity of RS‐FS conferred by RS. Concordantly, nanofibers were specifically formed in tumor cells from subcutaneous HCC mice treated with RS‐FS, but not with scrambled‐FS, in which nanofibers were found in both HCC and normal liver cells (Figure S1d). Altogether, the data demonstrate that RS‐FS assembles into nanospheres at neutral pH and transforms to nanofibers under acidic and reductive conditions with tumor cell selectivity, resulting in hydrogel formation and lysosome swelling in HCC cells specifically.

FIGURE 1.

Characterization of RS‐FS in vitro. (a) Diagram for the stimuli‐responsive transformation of RS‐FS. RS‐FS consisted of HCC‐targeting RS peptide (green), an acid‐responsive self‐assembling “vealyl” peptide (blue), and a hydrophobic “VVVVV” peptide (yellow) in which the lowercase letter represents D‐amino acids. Black dots represent thiol groups on cysteine residues within the RS peptide. (b) Transmission electron microscopy (TEM) images of RS‐FS and RS‐S morphologies under different conditions, including pH 7.4 (blood circulation), pH 6.5 (tumor extracellular matrix), pH 6.5 plus 10 mm GSH (early endosomes), and pH 5.0 plus 10 mm GSH (lysosomes) (scale bar = 200 nm). (c) Nanoparticle Tracking Analysis (NTA) of RS‐FS and RS‐S size distribution under different conditions. N.A. means not available. (d) Circular Dichroism (CD) spectra of RS‐FS at different conditions. (e) Macroscopic (inverted vial) and microscopic (TEM) morphologies of RS‐FS at different concentrations in buffer pH5.0 plus 10 mm GSH (scale bar = 50 nm). (f) Rheological analysis of RS‐FS under different conditions. (g) Fluorescence recovery after photobleaching (FRAP) assays of lysosomes in human hepatocyte HL7702 and HCC LM3 cells before and after treatment with 100 µm RS‐FS for 24 h (scale bar = 5 µm) (n = 3, data are presented as mean ± SEM, ^** p < 0.01, two‐tailed t‐test). (h) TEM images showing the effect of RS‐FS on lysosome ultrastructure in HL7702 and LM3 cells. Black solid arrows point to primary lysosomes; black hollow arrows indicate secondary lysosomes; white arrows point to lysosomes disrupted by RS‐FS (scale bar = 500 nm). (i) Immunofluorescence for cathepsin B in human hepatocyte HL7702 and HCC LM3 cells before and after treatment with 100 µM RS‐FS for 24 h (scale bar = 10 µm). The solid arrowheads point to the punctate cathepsin B staining; hollow arrowheads point to the diffuse cathepsin B staining (scale bar = 10 µm).

2.2. RS‐FS Promotes Tumor Cell‐Specific Uptake, Lysosomal Escape, and Antitumor Activity of Doxorubicin in Human HCC Cells in vitro

We next examined the effect of RS‐FS on doxorubicin's antitumor activity in vitro. As expected, higher fluorescence signals were found in human LM3 cells treated with RS‐FS‐ and RS‐S‐doxorubicin than in human liver 7702 cells, whereas no difference was observed for free doxorubicin between LM3 and 7702 cells (Figure 2a). Intriguingly, significantly stronger fluorescence intensity was found with RS‐FS‐doxorubicin than the RS‐S group despite a similar percentage of doxorubicin‐positive LM3 cells (Figure 2b), suggesting that nanofiber‐mediated lysosome disruption might reduce doxorubicin exocytosis [24] and enhance doxorubicin release in the cytoplasm. In LM3 cells, co‐localization of doxorubicin and LysoTracker signal suggests that doxorubicin is probably trapped in lysosomes. Unlike RS‐S, RS‐FS delivery resulted in higher doxorubicin levels in LM3 cell nuclei (Figure 2c,d), indicating RS‐FS‐mediated doxorubicin release via lysosome disruption. This phenomenon was generalizable as a similar outcome was observed with human 97H and HepG2 HCC cells and murine Hepa1‐6 HCC cells (Figure S2a,b). Significantly reduced cell proliferation and migration were detected in RS‐FS‐doxorubicin‐treated LM3 cells compared to the RS‐S group (Figure 2e,f), confirming the enhanced antitumor efficacy facilitated by RS‐FS‐mediated lysosome disruption. Notably, RS‐FS alone also dampened tumor cell growth via lysosome disruption while showing minimal effect on human normal liver cells (Figure S2c), demonstrating synergism with doxorubicin rather than just delivery. Collectively, these findings demonstrate that RS‐FS enables doxorubicin's tumor cell‐specific uptake, lysosomal escape, and antitumor inhibition in HCC cells in vitro.

FIGURE 2.

In vitro evaluation of DOX‐loaded RS‐FS. (a) Flow cytometric analysis and quantification of the percentage of DOX‐positive cells (n = 5; ^* p < 0.05, ^** p < 0.01, One‐way‐ ANOVA post hoc Student‐Newman‐Keuls test). DOX@RS‐FS refers to DOX‐loaded RS‐FS. (b) Mean fluorescence intensity (MFI) of DOX in HL7702 and LM3 cells treated with DOX@RS‐FS (n = 5; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (c) Intracellular localization of DOX (green) in HL7702 and LM3 cells treated with free DOX, DOX@RS‐S, or DOX@RS‐FS. Lysosomes were stained with LysoTracker (red) (scale bar = 5 µm). The nuclear region was outlined by the dashed line. (d) Quantitative analysis of colocalization of DOX with nuclei or lysosomes (n = 5; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (e) Cell proliferation inhibition assays for HL7702 and LM3 cells treated with free DOX, DOX@RS‐S, or DOX@RS‐FS (n = 3; ^* p < 0.05, ^** p < 0.01, one‐way ANOVA post hoc Student‐Newman‐Keuls test). (f) Cell migration inhibition assays for HL7702 and LM3 cells treated with free DOX, DOX@RS‐S, or DOX@RS‐FS (n = 5; ^* p < 0.05, ^** p < 0.01, one‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 100 µm). Data are presented as mean ± SEM.

2.3. RS‐FS Enhances Doxorubicin's Circulation, Tumor‐Specific Delivery, and Antitumor Potency in HCC Mice In Vivo

As nanoparticles generally have long circulatory half‐lives [25], we intravenously injected RS‐FS‐doxorubicin into day‐7 subcutaneous HCC mice once and examined doxorubicin fluorescence signals in circulation. Mice treated with RS‐FS‐doxorubicin exhibited 2.7 times higher fluorescence than free doxorubicin in plasma 2 h after injection (Figure 3a). Doxorubicin, which usually accumulates in the liver, was redirected to the tumor by RS‐FS, resulting in a 6.3‐fold tumor/liver ratio increase compared to free doxorubicin (Figure 3b,c). This tumor/liver ratio was durably 6‐fold higher in day‐7 orthotopic HCC mice treated with RS‐FS‐doxorubicin than free doxorubicin (Figure 3d,e), indicating tumor‐specific delivery and accumulation of doxorubicin conferred by RS‐FS.

FIGURE 3.

Stability and biodistribution of DOX@RS‐FS in HCC mice in vivo. (a) Measurement of plasma concentration of DOX at 2 h post‐injection in day‐10 subcutaneous HCC mice treated with free DOX or DOX@RS‐FS (n = 4; ^** p < 0.01, two‐tailed t‐test). (b) Tissue distribution and (c) quantitative analysis of DOX fluorescence signals in day‐10 subcutaneous HCC mice at 6 h post‐injection (n = 4; ^** p < 0.01, two‐tailed t‐test). B‐brain; Q‐quadriceps; Lu‐lung; S‐spleen; K‐kidney; H‐heart; Li‐liver; and Tu‐tumor. (d) Tissue distribution and (e) quantitative analysis of DOX fluorescence signals in orthotopic HCC mice at 6 h post‐injection (n = 4; ^** p < 0.01, two‐tailed t‐test). Data are presented as mean ± SEM.

Corroborating with tumor‐specific accumulation of doxorubicin, RS‐FS‐doxorubicin achieved significantly stronger tumor inhibition than free doxorubicin or RS‐FS alone in subcutaneous HCC mice (Figure S3a–c), indicating augmented antitumor effects of doxorubicin conferred by RS‐FS. Compared to multiple injections, a single intravenous injection of RS‐FS‐doxorubicin showed less profound antitumor effects (Figure S3d), we chose multiple repeated injections over a single injection for subsequent studies. Dynamic monitoring of tumor growth via Magnetic Resonance Imaging (MRI) revealed that tumor growth was strikingly retarded in orthotopic HCC mice treated with RS‐FS‐doxorubicin, with approximately 40‐fold reduction in tumor weight, compared to free doxorubicin, which showed incremental tumor size in a time‐dependent manner (Figure 4a–d). There was no bodyweight change between mice treated with RS‐FS‐doxorubicin and untreated controls, whereas gradual weight loss was observed after free doxorubicin treatment (Figure 4e), a commonly reported side effect for doxorubicin [26]. TdT‐mediated dUTP Nick‐End Labeling (TUNEL) and histological staining showed increased DNA fragmentation and nuclei degradation with atrophic nuclei in tumor tissues from mice treated with RS‐FS‐doxorubicin compared to free doxorubicin (Figure 4f,g), indicating RS‐FS enhanced doxorubicin‐mediated tumor cell apoptosis. We intravenously administered RS‐FS‐doxorubicin at 10 mg/kg doxorubicin daily for 3 days (Figure S4a) [26] with no decrease observed in bodyweight and survival in wild‐type mice treated with RS‐FS‐doxorubicin, whereas free doxorubicin caused sharp weight loss and reduced survival (Figure S4b,c), demonstrating that RS‐FS alleviates systemic toxicity caused by high doses of doxorubicin. Dramatic heart function deterioration was found in doxorubicin‐treated mice, indicated by declines in left ventricular end diastolic diameter, left ventricular end systolic diameter, and stroke volume via echocardiography (Figure S4d) and heart rates via electrocardiogram (Figure S4e), but these were not observed in RS‐FS‐doxorubicin‐treated mice. Similarly, cardiac cell nuclei aggregation, hepatic sinusoid enlargement, and renal tubular epithelial cell swelling with cytoplasmic vacuolization (Figure S4f) and significantly elevated levels of circulating lactate dehydrogenase (LDH), aspartate transaminase (AST), and urea (Figure S4g), reflective of heart, liver, and kidney side effects linked to doxorubicin [27], were not detected in mice treated with RS‐FS‐doxorubicin. These results demonstrate significantly reduced doxorubicin‐related toxicity conferred by RS‐FS [26, 28] and widening of the therapeutic window. In summary, these results showed that complexing with RS‐FS enables extended circulation, tumor‐specific delivery, enhanced antitumor activity, and reduced systemic toxicity of doxorubicin in HCC mice in vivo.

FIGURE 4.

Evaluation of therapeutic efficacy of DOX@RS‐FS in orthotopic HCC mice. (a) Diagram for dosing regimen of DOX or DOX@RS‐FS in orthotopic HCC mice. I.V. means intravenous injection. (b) Real‐time MRI monitoring of tumor growth in day‐7 orthotopic HCC mice treated with PBS, DOX, or DOX@RS‐FS at different time‐points. Tumors were circled. (c) Quantitative analysis of tumor volume of orthotopic HCC mice treated with PBS, DOX, or DOX@RS‐FS (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (d) Representative tumor images and quantitative analysis of tumor weight of orthotopic HCC mice treated with PBS, free DOX, or DOX@RS‐FS (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 1 cm). Tumors were circled. (e) Body weight curves of orthotopic HCC mice treated with PBS, free DOX, or DOX@RS‐FS at different timepoints (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (f) TUNEL staining of genomic DNA damage in tumor tissues from treated orthotopic HCC mice (scale bar = 100 µm). (g) H&E staining of tumor tissues from orthotopic HCC mice treated with PBS, DOX, or DOX@RS‐FS. Black arrowheads indicate atrophic nuclei; hollow arrowheads indicate areas of nuclear degradation. Data are presented as mean ± SEM.

2.4. Intravenous RS‐FS Potentiates Oral Lenvatinib's Effects, Resulting in Marked Tumor Regression in HCC Mice In Vivo

New molecular targeted drugs targeting tumor‐specific receptors, such as first‐line multiple kinase receptor inhibitor Lenvatinib, show certain tumor cell selectivity [29, 30], but can also trigger autolysosome formation and autophagic flux with resultant reduced potency [6]. RS‐FS's tumor cell‐specific lysosomal disruption may enhance the antitumor potency of drugs of this kind without the need to co‐assemble, thus acting like an adjuvant and avoiding alteration on Lenvatinib's clinically‐approved oral administration route. To this end, we administered intravenous RS‐FS at 50 mg/kg with oral Lenvatinib at 30 mg/kg [31] simultaneously for four times at 4‐day intervals into subcutaneous HCC mice (Figure 5a). Strikingly, tumor volume reduced 47‐fold in mice treated with co‐administration of RS‐FS and Lenvatinib compared to Lenvatinib alone, with one mouse being tumor‐free (Figure 5b). Remarkably, 50% of day‐7 orthotopic HCC mice had tumors eradicated on day 23 via MRI after intravenous RS‐FS and oral Lenvatinib at the same dosage as used in subcutaneous HCC mice, and tumor weight was reduced 61‐fold lower than Lenvatinib alone (Figure 5c–e). TEM demonstrated in tumor cells from HCC mice treated with RS‐FS and Lenvatinib (1) primary and secondary lysosomes disappeared, and (2) swollen mitochondria with reduced cristae, increased mitochondrial area, and circularity accumulated. In contrast, the number of autolysosomes increased, and mitochondria appeared damaged after Lenvatinib alone as reported previously [6] (Figure 5f). This suggests that RS‐FS selectively depletes the lysosome pool, resulting in blockade of autophagic flux and accelerating Lenvatinib‐mediated mitochondria damage with disappeared cristae and increased area in tumors [5]. Cytochrome c diffused into the cytoplasm due to the disappearance of cristae in damaged mitochondria [32] after RS‐FS and Lenvatinib treatment, with no change found between the Lenvatinib and PBS groups (Figure 5g). Markers for apoptosis via mitochondrial pathway [33], including cleaved caspase‐9 and caspase‐3, increased, and cell proliferation biomarker Ki67 decreased (Figure 5h) [34], tumor cell apoptosis (Figure 5i), and atrophic nuclei (Figure 5j) increased, as reflected by TUNEL and H&E staining, respectively, after combination treatment compared to Lenvatinib alone. The data collectively showed that the combination therapy with intravenous RS‐FS and oral Lenvatinib can dramatically reduce or eliminate tumors in orthotopic HCC in vivo, and RS‐FS functions by selectively disrupting tumor cell lysosomes, leading to blockade of autophagic flux and accumulated damaged mitochondria in tumor cells.

FIGURE 5.

Investigation of the effect of intravenous RS‐FS on oral Lenvatinib in HCC mice in vivo. (a) Diagram for dosing regimen of LEN or LEN plus RS‐FS in subcutaneous and orthotopic HCC mice. (b) Representative tumor images and quantitative analysis of tumor volume and weight in subcutaneous HCC mice treated with PBS, Lenvatinib (LEN) alone, or LEN plus RS‐FS (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). Real‐time MRI monitoring of tumor growth (c) and quantitative analysis (d) in day‐7 orthotopic HCC mice treated with PBS, LEN, or LEN plus RS‐FS at different time‐points (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). Tumors were circled. (e) Representative tumor images and quantitative analysis of tumor weight in orthotopic HCC mice treated with PBS, LEN, or LEN plus RS‐FS (n = 6; ^** p<0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 1 cm). Tumors were circled. (f) TEM images of tumor tissues from treated orthotopic HCC mice showing mitochondrial and autolysosome ultrastructure (scale bar = 1 µm) and quantification of Cristae density, mitochondrial perimeter, area, and circularity (n = 25; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). M‐mitochondria; PL‐primary lysosomes; AL‐autolysosomes; Arrowheads indicate damaged mitochondria. Cristae density was determined by the number of cristae within a mitochondrion divided by the total mitochondrial length. Circularity = 4π × area/ perimeter². (g) Immunofluorescence staining for Cytochrome c (Cyt‐C) in tumor tissues from treated orthotopic HCC mice (scale bar = 10 µm). Solid arrowheads point to the punctate Cyt‐C staining; hollow arrowheads point to the diffuse Cyt‐C staining. (h) Immunohistochemical staining and quantitative analysis of Cleaved Caspase‐9, Cleaved Caspase‐3, and Ki67 in tumor tissues from treated orthotopic HCC mice (n = 5; ^** p<0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 50 µm). TUNEL apoptosis (i) and H&E (j) staining of tumor tissues from treated orthotopic HCC mice (scale bar = 100 µm). Arrowheads indicate atrophic nuclei. Data are presented as mean ± SEM. N.s. means not significant.

2.5. RS‐FS Augments EEV's Tumor‐Suppressing Ability by Promoting Tumor Cell‐Specific Lysosomal Escape and Autophagic Flux Blockage in HCC Mice In Vivo

Encouraged by the potentiating effect of RS‐FS on molecular targeted drug Lenvatinib, we wished to evaluate the generalizability by combining RS‐FS with EEV, EVs from Epimedium brevicornu Maxim., a medicinal herb whose extracts were shown to effectively inhibit HCC tumor growth in vivo [35]. More importantly, EEV is enriched with icariin, which can be easily converted to icaritin, an anti‐HCC traditional Chinese medicine clinically approved in China [36]. Similar to Lenvatinib [30], icaritin was reported to have tumor cell‐selective toxicity, whereas EEV bearing icaritin can potentially overcome icaritin's poor aqueous solubility and cell permeability with EV's favorable bioavailability, cellular uptake, and stability [37]. Unsurprisingly, icariin was effectively converted to icaritin in EEV by snailase [38] without altering the morphology and size of EEV (Figure S5a–c). Consistent with the potent and tumor cell‐selective inhibitory effect of icaritin, EEV after snailase treatment exhibited stronger inhibition on LM3 cell proliferation than untreated EEV (Figure S5d). Strikingly, EEV induced significant tumor inhibition, with 16‐ or 8‐fold higher antitumor potency observed for equal particles of EEV with or without snailase treatment than the equivalent amount of icaritin, respectively (Figure S5d), confirming that EEV is important for the bioavailability of icaritin. RS‐FS further enhanced EEV's antitumor inhibition and tumor apoptosis in human LM3 cells in vitro (Figure S6a,b) and in day‐7 subcutaneous HCC mice in vivo (Figure S6c), validating the synergistic effect of RS‐FS and EEV in tumor suppression.

Co‐localization of PKH67‐labeled EEV with LysoTracker signals decreased in LM3 cells after RS‐FS and EEV treatment compared to EEV alone (Figure 6a), whereas no difference was found in H7702 cells (Figure S7a), ascertaining RS‐FS‐mediated tumor cell‐specific lysosome disruption and lysosomal escape of EEV. Administration of intravenous RS‐FS at 50 mg/kg and oral EEV at 5 (10¹¹ particles/kg for 3 weeks at every other day intervals gradually shrank the tumor in orthotopic HCC mice, with one sixth of mice being tumor‐free, whereas EEV alone failed to halt tumor growth (Figure 6b–e). We posited that RS‐FS‐facilitated lysosomal release of icaritin‐bearing EEV accelerates production of reactive oxygen species (ROS) and mitochondrial damage, resulting in enhanced tumor cell apoptosis, mobilization of dendritic cells, and cytotoxic T cell infiltration as reported for icaritin [39] (Figure 6f). Indeed, levels of ROS were significantly elevated (Figure 6g), the number of autolysosomes significantly declined, damaged mitochondria significantly rose (Figure 6h), and circulating mitochondrial DNA leakage increased (Figure S7b) after combination treatment compared to EEV alone. As accumulated ROS triggers permeabilization of mitochondrial membrane [40], RS‐FS‐mediated blockade of autophagic flux can synergistically accelerate mitochondria damage triggered by EEV. Increased apoptosis and reduced proliferation of tumor cells (Figure 6i) and significantly elevated expression of biomarkers for dendritic cell (DC) maturation [41]‐major histocompatibility complex I (MHC I), CD86, and CCR7‐ in intratumoral DCs (Figure 6j) in mice treated with combination therapy compared to EEV alone demonstrate that leaked mitochondrial DNA promoted tumor cell apoptosis and DC maturation [42]. Correspondingly, significantly increased infiltration of CD8⁺ T cells and reduced immunosuppressive Foxp3⁺ Tregs were found after combination therapy compared to EEV alone (Figure 6k). These results demonstrated that RS‐FS augments EEV's antitumor potency by selectively disrupting tumor cell lysosomes with resultant blockade of autophagic flux and acceleration of mitochondria damage, leading to enhanced tumor apoptosis and T cell infiltration in orthotopic HCC mice in vivo.

FIGURE 6.

Systemic evaluation of intravenous RS‐FS on oral EEV in orthotopic HCC mice in vivo. (a) Subcellular localization of EEV (green) in LM3 cells treated with EEV or EEV plus RS‐FS (scale bar = 5 µm). Lysosomes were stained with LysoTracker (red) (n = 5; ^** p < 0.01, two‐tailed t‐test). (b) Diagram for dosing regimen of EEV or EEV plus RS‐FS in orthotopic HCC mice. Real‐time MRI monitoring (c), quantitative analysis (d) of tumor growth in day‐10 orthotopic HCC mice treated with PBS, EEV, or EEV plus RS‐FS at different timepoints (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). Tumors were circled. (e) Representative tumor images and quantitative analysis of tumor weight in orthotopic HCC mice treated with PBS, EEV, or EEV plus RS‐FS (scale bar = 1 cm) (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). Tumors were circled. (f) Illustration of the potential mechanism of RS‐FS‐mediated drug release, apoptosis, and antitumor immunity. (g) Measurement of ROS levels in tumor tissues from orthotopic HCC mice treated with PBS, EEV, or EEV plus RS‐FS (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (h) TEM images of tumor tissues showing autolysosomes and mitochondrial ultrastructure (scale bar = 1 µm) and quantitative analysis of autolysosomes and mitochondria per 100 µm² cytoplasmic area (n = 6; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). M‐mitochondria; PL‐primary lysosomes; AL‐autolysosomes; Arrowheads indicate damaged mitochondria. AM‐mitochondria in autophagy; DM‐damaged mitochondria; and NM‐normal mitochondria. (i) Immunohistochemical staining and quantitative analysis of cleaved Caspase‐9 and Ki‐67 in tumor tissues from treated orthotopic HCC mice (n = 5; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 100 µm). (j) Flow cytometric analysis of CD86, MHC‐I and CCR7 in intratumoral DCs from treated orthotopic HCC mice (n = 4; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test). (k) Immunohistochemical staining and quantitative analysis of CD8⁺ T cells and Foxp3⁺ Tregs in tumor tissues (n = 5; ^** p < 0.01, One‐way ANOVA post hoc Student‐Newman‐Keuls test) (scale bar = 100 µm). Data are presented as mean ± SEM.

3. Discussion

One of the major obstacles for HCC chemotherapy or molecular targeted therapy is that tumor cells can self‐regulate antitumor therapeutics either by lysosome sequestration or lysosome‐facilitated autophagic flux [6, 7, 24], thus tumor‐specific lysosome disruption would represent a viable path to enhance antitumor therapeutics’ efficacy. By combining HCC‐targeting peptide with nanofiber‐ and nanosphere‐forming motifs liable to environmental stimuli, we demonstrated that a 23‐aa short linear peptide RS‐FS can self‐assemble into nanospheres or transform to nanofibers in response to different environmental stimuli with tumor cell selectivity, resulting in tumor cell‐specific lysosome disruption and subsequent drug lysosomal release and blockade of autophagic flux, and augmented antitumor potency with minimal effects on normal liver cells in human HCC cells in vitro or HCC mice in vivo. RS‐FS can enhance therapeutics’ potency by either encapsulating drugs such as doxorubicin or potentiating therapeutics after co‐administration without altering the drugs’ administration route, as shown with Lenvatinib and EEV. Thus, we provide a generalizable peptide vector and adjuvant to enhance antitumor therapeutics’ potency with reduced side effects without complex synthesis, conjugation, or encapsulation, and alteration of the clinically‐approved drugs’ administration route. The versatility makes this approach easily translatable clinically.

Most importantly, unlike other peptide designs, which require pretreatment in vitro [11, 13, 36], linear RS‐FS can naturally self‐assemble into nanospheres or nanofibers under different environmental conditions. Unlike chloroquine, a classical lysosome‐tropic agent that functions through pH neutralization and decreased autophagosome‐lysosome fusion without cell selectivity [43], RS‐FS can selectively disrupt tumor cell lysosomes with minimal off‐target effects on other cells, resulting in a favorable therapeutic window. Cytosolic release of cathepsin B also showed RS‐FS triggered lysosomal membrane permeabilization in HCC cells. However, no degradation of LAMP2 was found with RS‐FS (data not shown), similar to LMP induced by chloroquine [44], possibly due to the lack of enzymatic cleavage activity such as sialidase as reported for hemagglutinin‐neuraminidase (HN) protein of Newcastle disease virus [45], although other possibilities cannot be excluded. As Good Manufacturing Practice (GMP) production of linear peptides is well‐established [46], this approach is easily translatable to manufacturing. Additionally, the advent of molecular targeted therapeutics that are already tumor‐selective, such as Lenvatinib [30], intravenous RS‐FS can sensitize oral Lenvatinib by triggering tumor cell‐specific lysosome disruption, thus blocking the enhanced autophagic flux induced by liver‐accumulated Lenvatinib [47]. Intrinsic liver accumulation of Lenvatinib also explains significantly greater antitumor efficacy of RS‐FS with Lenvatinib in orthotopic than in subcutaneous HCC mice. Along the same line, icaritin also shows tumor cell selectivity, though suffering from low biocompatibility and cell permeability [37]. In contrast, EEV‐enriching icaritin after snailase conversion exhibits favorable biocompatibility, cell permeability, and stability. Likewise, intravenous RS‐FS enhanced oral EEV's antitumor potency by specifically disrupting tumor cell lysosomes, leading to lysosomal escape and blockade of autophagic flux and accelerating EEV‐mediated mitochondria damage. These results show that a linear peptide vector can augment antitumor therapeutics’ efficacy without the need for drug loading. Therefore, our findings may have significant implications for the development of targeted therapy in HCC, with extension to other tumors, given the extensive use of molecular targeted therapeutics as front‐line treatments for cancer patients with reported side effects [2].

While fluorescent doxorubicin was used as a model drug because it could be easily monitored in vitro and in vivo, we acknowledge that doxorubicin is no longer employed for HCC in the clinic [48], but this approach can be exploited for other tumors [49] with the appropriate targeting ligand in place of RS, considering the large number of clinically‐approved chemotherapeutics [7]. Although RS‐FS is a tumor‐targeted drug delivery vehicle, tumor cell‐specific lysosome disruption makes RS‐FS a promising non‐classical drug adjuvant to enhance targeted therapeutics for HCC and other tumors in the clinic [2, 50], especially as lysosomes play a critical role in regulating drug potency [7, 9, 24]. Also, further detailed studies on the functionality of RS‐FS in drug‐induced resistance models, such as Lenvatinib, are warranted in the future [31]. Nevertheless, this is the first proof‐of‐concept study to demonstrate that a short multifunctional peptide amplifies antitumor drug potency by specifically disrupting tumor cell lysosomes, thus providing a new peptide tool for tumor‐targeted therapy in a wide array of cancers.

In summary, we demonstrated that RS‐FS, a linear 23‐aa peptide consisting of hydrophilic HCC‐targeting peptide, hydrophobic nanosphere‐ and nanofiber‐forming motifs, enables amplified antitumor therapeutics’ potency via tumor cell‐specific lysosome disruption, resulting in lysosomal escape and blockade of autophagic flux triggered by antitumor drugs, with minimal effects on normal liver cells, in HCC models in vitro and in vivo, thus providing a new peptide tool for targeted therapeutics in HCC without complex synthesis, conjugation, or encapsulation, and alteration of clinically‐approved drugs’ administration routes.

4. Materials and Methods

4.1. Animals and Injections

C57BL/6 wild‐type mice (6–8‐week‐old) were used in all experiments (the number used was specified in Figure legends). Mice were housed under specific pathogen‐free conditions in a temperature‐controlled room. The experiments were carried out in the Animal unit, Tianjin Medical University (Tianjin, China), according to procedures authorized by the institutional ethical committee (Permit Number: SYXK2023‐0004). Mice were sacrificed by CO₂ inhalation or cervical dislocation at desired time‐points, and tissues were fixed with Bouin's solution (Sigma, St. Louis, MO, USA) and embedded with paraffin for immunohistochemical and histological studies.

4.2. Cell Culture

Murine HCC cell line Hepa1‐6 was purchased from Boster Biological Technology Ltd. (Wuhan, China). The human liver cell line HL7702 and human HCC cell lines MHCC‐LM3 and MHCC‐97H were obtained from BeNa Culture Collection (Beijing, China), and HepG2 was sourced from the American Type Culture Collection. Cell lines were cultured in Dulbecco's modified Eagle's medium (DMEM) supplemented with 2 mm glutamine and 10% fetal bovine serum.

4.3. Establishment of Tumor‐Bearing HCC Mouse Models

To establish mouse models bearing subcutaneous HCC tumors, Hepa1‐6 (1 × 10⁶) cells were suspended in 70 µL PBS and injected into C57BL/6 mice subcutaneously. For orthotopic HCC mice, subcutaneous Hepa1‐6 tumors with a longitudinal diameter of 1 cm were collected from subcutaneous mice after cervical dislocation. Tumor tissues were washed in D‐Hanks buffer, and necrotic tumor tissues were removed; the remaining tumor tissues were cut into about 1mm³ pieces. The recipient mice were anesthetized with isoflurane, and the skin was sterilized with iodophor three times before surgery as described previously [51]. Tumor tissues (3–5 pieces) were implanted in the left lobe of the liver in the recipient mice under anesthesia, and the peritoneum and skin were closed with 4–0 sutures. DOX (5 mg/kg), RS‐FS (50 mg/kg), or DOX‐loaded RS‐FS (5 mg/kg DOX and 50 mg/kg RS‐FS) was administered intravenously to mice bearing subcutaneous or orthotopic Hepa1‐6 tumors. The treatment commenced on day 7 after tumor inoculation, with injections given every 4 days for a total of four doses. To assess the acute toxicity of DOX and DOX‐loaded RS‐FS, both formulations were administered intravenously at a daily dose of 10 mg/kg (based on DOX amount) for 4 consecutive days. Subsequently, blood samples were collected via the retro‐orbital plexus for serum biochemical analysis of urea, lactate dehydrogenase (LDH), aspartate aminotransferase (AST), and creatine kinase (CK). For Lenvatinib experiments, Lenvatinib (30 mg/kg, suspended in PBS with 1% Tween 80) was administered orally every 4 days for a total of four doses, starting on day 7 after tumor inoculation. In the combination group, mice received the same amount of Lenvatinib orally with intravenous RS‐FS (50 mg/kg) simultaneously. For Icaritin and EEV experiments, Icaritin (50 mg/kg suspended in PBS with 1% Tween 80) and EEV (5 × 10¹¹ particles/kg suspended in PBS with 10 mm EDTA) were administered orally on day 7 for subcutaneous HCC mice and on day 10 for orthotopic HCC mice every other day for 15 or 23 days, respectively. For the EEV and RS‐FS combination group, mice received oral EEV and intravenous RS‐FS (50 mg/kg) under identical dosing regimens as described above. Tumor volume was measured at different timepoints as specified in the figure legends. For subcutaneous HCC models, tumor volume (length‐L and width‐W) was measured using calipers and calculated as (L × W²)/2. For orthotopic HCC models, tumor volume was determined with Magnetic resonance imaging (MRI) images.

4.4. Magnetic Resonance Imaging (MRI)

MRI images of orthotopic HCC mice were acquired using T2 propeller sequence [51] with the following parameters: slice thickness of 1.0 mm, slice spacing of 0.5 mm, TR/TE of 3494/70.7 ms, matrix of 256 × 160 and FOV of 8 × 8 cm (3.0 Tesla MR scanner, Signa Excite HDx; GE healthcare, Milwaukee, WI, US) with a small animal coil in Tianjin Medical University General Hospital. During the examination, mice were anesthetized with pentobarbital sodium and fixed to minimize body motion.

4.5. Pharmacokinetics and Biodistribution

C57BL/6 wild‐type mice (6–8‐week‐old) were intravenously administered 20 mg/kg DOX or DOX‐loaded RS‐FS. Blood was collected via the retro‐orbital plexus at 2 h post‐injection. Plasma was recovered by centrifugation and added to 96‐well Black Polystyrene Medium Bind Stripwell plates and imaged with the IVIS spectrum (PerkinElmer IVIS Spectrum, USA). For the biodistribution, subcutaneous and orthotopic HCC tumor‐bearing mice were injected with 20 mg/kg free DOX or DOX‐ loaded RS‐FS, or subcutaneous HCC tumor‐bearing mice were injected with 5 mg/kg Cy5‐labeled RS‐FS or scrambled‐FS peptide or with 50 mg/kg RS‐FS or scrambled‐FS intravenously for a single injection. The mice were terminally anesthetized and perfused with 50 mL cold PBS 6 or 24 h post‐injection with tissues and tumors harvested for imaging with the IVIS spectrum imaging system.

4.6. Cardiac Function Analysis

Cardiac function was assessed by non‐invasive transthoracic echocardiography and electrocardiography (Vevo2100, FUJIFILM Visual Sonics, USA) in anesthetized mice (induced with 2% isoflurane and maintained on 1.5% isoflurane in 100% oxygen) as reported before [26]. Core temperature was maintained at 37.0°C ± 0.5°C with a heating pad. Heart rate and a lead II electrocardiogram (ECG) were continuously monitored via limb electrodes. Two‐dimensional echocardiographic images and M‐mode tracings were acquired using a high‐frequency transducer to obtain parasternal long‐axis and short‐axis views at the level of the papillary muscles. Left ventricular end‐diastolic diameter (LVEDD) and left ventricular end‐systolic diameter (LVESD) were measured from the M‐mode recordings. The corresponding LV volumes at end‐diastole (Ved) and end‐systole (Ves) were then calculated, respectively, using the cubic formula (V = 1.047 × LVID³). Stroke volume (SV) was derived as SV = Ved −Ves.

4.7. Microenvironment Mimicking Buffers

Buffers simulating the pH and redox conditions in the blood circulation, extracellular matrix of tumor cells, early endosome, and lysosome were prepared for in vitro analysis of RS‐FS. Buffer A for mimicking blood circulation: 0.1 m Phosphate Buffer Solution (PBS) (pH 7.4). Buffer B for mimicking the extracellular matrix of tumor cells: 0.1 m 2‐(N‐Morpholino) ethanesulfonic acid (MES) buffer (pH 6.5). Buffer C for mimicking early endosome: 0.1 m MES buffer (pH 6.5) with 10 mm GSH. Buffer D for mimicking lysosome: 0.1 m acetate buffer (pH 5) with 10 mm GSH. All buffers were rendered isotonic by adding NaCl to match the osmolarity of physiological conditions (300 mOsm/L).

4.8. Peptide Assembly

Self‐assembled RS‐FS (VVVVVvealylCGRCKCCNGERS), RS‐S (VVVVVvCGR CKCCNGERS), Cy5‐labeled RS‐FS (VVVVVvealylCGRCKCCNGERS), and Cy5‐ labeled scrambled‐FS (VVVVVvealylSCGNCRCERCGK) were synthesized by China ‐Peptides (Shanghai, China) with 95% purity, in which the lowercase letter represents D amino acid. Peptides (2 mg) were dissolved in 10 mL PBS buffer (0.1 m, pH 7.4) at room temperature (RT). To oxidize, the solution was supplemented with H₂O₂ at a final concentration of 10 mm and incubated for 24 h at 37°C. To prevent oxidation, the solution preparation and incubation were conducted under a nitrogen atmosphere using nitrogen‐saturated buffers.

4.9. Epimedium Brevicornu Maxim.‐Derived Extracellular Vesicles (EEV) Isolation

EEV were isolated with the Phyto‐EVpure method [52]. Briefly, fresh leaves of Epimedium brevicornu Maxim. (purchased from Anguo Traditional Chinese Medicine Market, Hebei, China) were washed and surface‐sterilized with 70% ethanol for 30 s with agitation. After five rounds of rinse with distilled water, tissues were soaked for 8 h in a digestion buffer (20 mmol/L MES, 2 mmol/L CaCl₂, 0.1 mol/L NaCl, pH 5.5) supplemented with antibiotics (100U/mL penicillin, 0.1 mg/mL streptomycin, and 0.25 µg/mL Amphotericin B). Then sterilized plant tissues were digested with the enzyme solution containing 1% cellulase (MERYE, China) and 0.5% pectinase (Yuanye, China) in the same antibiotic‐supplemented buffer at 37°C with shaking at 200 rpm for 4 h in which enzymes were centrifuged at 15 000 g for 15 min and filtered through a 0.22 µm filter, followed by ultra‐centrifugation at 100 000 g for 1 h to remove endogenous EV and particulates. The supernatants after digestion were filtered through sterile gauze and subjected to centrifugation at 3000 g for 20 min and 10 000 g for 40 min at 4°C to remove cellular debris. Finally, EEV were pelleted by ultracentrifugation at 100 000 g for 70 min at 4°C, resuspended in sterile PBS, and stored at −80°C for subsequent use.

4.10. Snailase Treatment of EEV

To convert icariin in EEV to active aglycone metabolite Icaritin, EEV were treated with snailase (10 mg/mL, Solarbio, China) in PBS (pH 5.8) with 0.1% saponin at 42°C overnight as reported previously [37]. Snailase‐treated EEV were pelleted by ultra‐centrifugation at 100 000 g for 70 min at 4°C, resuspended in sterile PBS, and stored at −80°C. To verify the conversion, pre‐ and post‐treated EEV with snailase (1 × 10¹¹ particle) were lysed with methanol (2 mL) and vortexed for 1 min and sonicated on ice for 15 min. The mixture was then centrifuged at 12 000 g for 10 min at 4°C to remove insoluble debris, and the supernatant was carefully collected for liquid chromatography‐mass spectrometry (LC‐MS) analysis (Agilent 7500ce, Agilent Technologies, Waldbronn, Germany).

4.11. Nanoparticle Tracking Analysis

The size distribution of RS‐ F Sand EEV was measured by Nanosight (NS300, Malvern, UK) as per the manufacturer's instructions. Briefly, the camera level was set at level 14 as all particles were distinctly visible without signal saturation at this level. Captures and analyses were achieved by using the built‐in NanoSight Software (NTA3.3.301, Malvern, UK). For each measurement, five consecutive 60 s videos were recorded under the following conditions: cell temperature at 25°C; syringe speed at 30 µL/s.

4.12. Circular Dichroism (CD) Spectrum Analysis

CD spectra of peptide molecules under different conditions were by measured by a Circular Dichroism spectrometer (J‐1700, JASCO, Japan) using 1 mm quartz cuvette. The concentration of the sample is 0.1 mm in buffer A, B C, and D. CD spectra of samples were analyzed by DichroWeb (http://dichroweb.cryst.bbk.ac.uk/), an open‐access website for calculating protein secondary structure from circular dichroism spectroscopic data.

4.13. Rheological Measurement

The rheological measurement of RS‐FS (5 mg/mL) in buffer A, B, C, and D, respectively, was determined by a rheometer (ARES‐G2, TA‐Waters, USA) using a plate with a diameter of 25 mm under a 0.5 mm gap. The storage modulus (G′) and loss modulus (G′′) were measured at a fixed frequency of 1 Hz and a fixed strain amplitude of 1% at 25°C.

4.14. Critical Gelation Concentration (CGC) Measurement

The CGC of RS‐FS in buffer D was measured by the vial inverse method as reported previously [15]. Briefly, RS‐FS was dissolved in buffer D with 1% eosin‐Y (E8090, Solarbio, Beijing, China) at final concentrations of 0.25, 0.5, 1, 2, 4, and 8 mg/mL in transparent glass vials. After being sealed and incubated at 37°C for 6 h, the vials were inverted to determine the gelation extent. The minimum concentration of RS‐FS for gelation was considered as the CGC.

4.15. Critical Micelle Concentration (CMC) Measurement

To measure the critical micelle concentration (CMC), Rhodamine 6G was mixed with RS‐FS in buffer A, B, C, or D at a final concentration of 5 µm [15]. After incubation at RT for 24 h, the maximum absorption wavelength (λmax) of Rhodamine 6G was measured using a Microplate Reader (Varioskan Lux, ThermoFisher, USA). The CMC value was determined by plotting the λmax against the logarithm of the peptide concentration, identifying the intersection of two linear regression lines.

4.16. Zeta Potential Measurement

The zeta potential of RS‐FS was determined with Zetasizer Nano‐ZS (Malvern Instruments, Worcestershire, UK). Briefly, oxidized RS‐FS was mixed with buffer A, B, C, or D at a final concentration of 0.2 mg/mL and incubated for 6 h at 37°C before measurement. Nanosphere samples (RS‐FS in buffer A, B, or C) were measured directly using standard capillary cells. The nanofiber samples (RS‐FS in buffer D) were measured in low‐field mode with a folded capillary cell.

4.17. Fluorescence Recovery after Photobleaching (FRAP) Assay

The FRAP assay was adopted to assess cell lysosomal membrane permeability [22, 53]. Cells were pre‐treated with RS‐FS (100 µm) for 24 h, and lysosomes were labeled with Lyso‐Tracker Red (GC19882, GlpBio, USA). Region of interest (ROI) within a lysosome was photobleached using a 561 nm laser at 100% power. The recovery of fluorescence into the bleached area was then monitored by time‐lapse imaging at 20s intervals.

4.18. Transmission Electron Microscopy (TEM)

Pre‐treated RS‐FS or RS‐S (0.1 mm) were dropped on a carbon‐coated grid and then stained with 2% uranyl acetate for 1 min. The morphologies of peptide molecules were measured by a transmission electron microscope (HT7700, Hitachi, Japan). Extracellular vesicles derived from Epimedium brevicornu Maxim. were combined with an equal volume of 4% paraformaldehyde in PBS for fixation [54]. A 20 µL aliquot of the mixture was then placed onto a sheet of parafilm, followed by adsorption onto a carbon‐coated copper grid at RT for 20 min. The grid was subsequently rinsed twice with PBS, followed by two sequential washes with ddH₂O for 5 min each time, to minimize ionic background. Next, the grid was immersed in 100 µL of 1% glutaraldehyde for 5 min and then washed eight times with 100 µL ddH₂O for 2 min per wash. Subsequently, the grid was placed into 100 µL uranyl oxalate solution (pH 7.0) for 5 min, followed by immersion in 100 µL a methyl cellulose‐uranyl acetate mixture (prepared by combining 100 µL of 4% uranyl acetate with 900 µL of 2% methyl cellulose) on ice for 10 min. Excess liquid was carefully removed, and the sample was allowed to dry before examination using TEM (HT7700, Hitachi, Japan). MHCC‐LM3 and HL7702 cells were seeded in 10 cm dishes at a density of 2 × 10⁶ cells/dish for 24 h, and then fresh culture medium containing 100 µm RS‐FS peptides was added and incubated for another 24 h. After washing with PBS for three times, cells were fixed by 2.5% glutaraldehyde for 5 min, and then scraped off with a scraper and collected by centrifugation. Cells were incubated in 2% osmium tetroxide, stained with 2% uranyl acetate, dehydrated in graded ethanol, and embedded in Spurr's resin. Ultrathin sections were obtained using a diamond knife. After being contrasted with solutions of 3% uranyl acetate and lead citrate, ultrathin sections of MHCC‐LM3 and HL7702 cells were examined with TEM. Likewise, tumor tissues were held in 2.5% glutaraldehyde in 0.1 m phosphate buffer (pH 7.4) for 10 min. Then, the tissues were cut into 2 × 2 × 2 (mm) cubes and immersed in 2.5% glutaraldehyde overnight at 4°C. Tissues were rinsed in PBS and treated with 2% osmium tetroxide for 1 h. The tissues were then dehydrated in a series of escalating concentrations of ethanol and embedded in Spurr's resin. Ultrathin sections were obtained using a diamond knife. After being contrasted with solutions of 3% uranyl acetate and lead citrate, ultrathin sections were examined with TEM.

4.19. Doxorubicin (DOX) Loading and Release

DOX‐loaded RS‐FS or ‐RS‐S were prepared by co‐dissolving peptides (10 mg) with DOX (2 mg) in DMSO (5 mL), followed by dialysis with regenerated cellulose (RC) dialysis bags (SP133336, MWCO 20 kDa, Shanghai Yuanye Bio‐Technology Co. Ltd, Shanghai, China) against PBS buffer (0.1 m, pH 7.4) for 24 h with buffer changes every 4 h to remove DMSO and unloaded DOX. To obtain oxidized formulations, the dialysis buffer was supplemented with H₂O₂ (10 mm) for an additional 24 h. For non‐oxidized formulations, all procedures, including solution preparation and dialysis, were conducted under a nitrogen atmosphere using nitrogen‐saturated buffers to prevent oxidation. An in vitro drug release assay was carried out at physiological temperature (37°C) by sequentially immersing the dialysis bag in 100 mL of buffer A, buffer B, buffer C, and buffer D under gentle stirring. The amount of DOX released from nanospheres was measured by a luminescence spectrometer (Biotek Synergy HT) at the emission wavelength of 595 nm and excitation wavelength of 485 nm. After the completion of the entire release study, the residues after drug release were taken out and freeze‐dried. Then the dried sample was pressed into powder and dissolved in 1 mL of DMSO. The amount of residual DOX was determined through a luminescence spectrometer (Biotek Synergy HT). The total amount of DOX loaded in the initial nanoparticles was calculated as the sum of the DOX released from the sample and the residual DOX. The cumulative release percentage was calculated as the ratio of the amount of DOX released from the nanoparticles to the total amount of DOX initially loaded.

4.20. Cellular Uptake

Confocal laser scanning microscopy (CLSM) and flow cytometry were employed to examine cellular uptake and intracellular distribution of DOX for DOX‐loaded nanospheres. Briefly, human HL7702, MHCC‐LM3, MHCC‐97H, HepG2, and murine Hepa1‐6 cells were cultured in 35 mm confocal dishes (CellVis, USA). Cells were treated with 5 µm D DOX or DOX‐loaded nanospheres with an equal amount of DOX and 100 µm RS‐FS. After 2 h incubation, the culture media was removed, and the cells were rinsed twice with PBS to remove free nanospheres or DOX. Then the lysosomes were labeled with LysoTracker Red (GC19882, GlpBio, USA). Cells were observed with CLSM (FV1000, Olympus, Japan) with DOX excited at 485 nm at the emission of 595 nm, and LysoTracker Red was excited at 577 nm with the emission at 590 nm. Cellular uptake was measured with Flow cytometry (FACSVerse, BD, USA) with excitation and emission wavelengths of 488 and 595 nm, respectively. For EEV cellular uptake, HL7702 and LM3 cells were incubated with PKH67‐labeled snailase‐treated EEV (5 × 10⁹ particles/mL) and RS‐FS (100 µm) for 2 h. Subsequently, cells were washed with PBS and stained with LysoTracker Red, followed by visualization using CLSM at the excitation wavelength of 488 or 577 nm and emissions at 502 or 590 nm for PKH67 and LysoTracker Red, respectively.

4.21. 3‐(4,5‐Dimethylthiazol‐2‐yl)‐2,5‐Diphenyltetrazolium Bromide (MTT) Assay

MTT assay was performed to evaluate cell cytotoxicity in vitro. Cells were seeded in 96‐well plates at a density of 2000 cells per well and cultured for 24 h. For RS‐FS‐alone, cells were treated with RS‐FS at 20, 100, or 200 µm. For DOX tests, cells were treated with DOX‐loaded RS‐FS or ‐RS‐S formulations at 100 µm peptides containing varying concentrations of DOX. HL7702 and MHCC‐LM3 cells were treated with 50, 200, or 400 nm DOX, DOX‐loaded RS‐S or ‐RS‐FS formulations. To test EEV, HL7702 and MHCC‐LM3 cells were treated with 2.5, 5, 10, 20, and 50 µm Icaritin, 5 × 10⁸, 1 × 10⁹, 5 × 10⁹, and 1 × 10¹⁰ particles/mL pre‐treated EEV or snailase‐ treated EEV, or snailase‐treated EEV plus RS‐FS (100 µm). After 48 h incubation, 10 µL of MTT (5 mg/mL in PBS) solution was added to each well and incubated for 4 h. The medium was then replaced with 100 µL DMSO to dissolve formazan crystals, with absorbance measured at 570 nm using a microplate reader (Tecan, Sunrise, Switzerland).

4.22. Transwell Assay

Cell migration was evaluated using Transwell chambers (8 µm pore size). HL7702 and MHCC‐LM3 cells were treated with 50 nm DOX, DOX‐loaded RS‐S (50 nm DOX with 100 µm RS‐S), or DOX‐loaded RS‐FS (50 nm DOX with 100 µm RS‐FS) for 48 h, followed by washing, trypsinization, and centrifugation to remove free drugs. Cells were seeded in the upper chamber at 8 × 10⁴ cells per well in serum‐free medium, while lower chambers contained complete medium. After 24 h incubation at 37°C, cells were fixed with 4% paraformaldehyde, with non‐migrated cells removed with cotton swabs in the upper chamber, and migrated cells in the lower chamber were stained with 1% crystal violet. Images were captured using an inverted microscope (Olympus BX51, Japan) and analyzed with ImageJ software (NIH, Bethesda, USA).

4.23. Cell Apoptosis Assay

EEV‐ or EEV plus RS‐FS‐induced cell apoptosis was measured with an Annexin V and Propidium Iodide (PI) Apoptosis Analysis Kit (Tianjin Sungene Biotech Co, Tianjin, China) as per manufacturer's instructions. Briefly, MHCC‐LM3 cells were seeded in 12‐well plates at a density of 1 × 10⁵ cells per well and cultured for 48 h. Cells were then treated for 48 h with PBS (control), EEV (1 × 10¹⁰ particles/mL), or a combination of EEV (1 × 10¹⁰ particles/mL) and RS‐FS (100 µm). After treatment, cells were collected by trypsinization without EDTA, followed by washing twice with PBS, and resuspended in 1 × Binding Buffer. Cell suspensions were incubated with Annexin V‐FITC and propidium iodide (PI) for 15 min at RT in the dark. Apoptosis was analyzed immediately using a flow cytometer (FACSVerse, BD, USA).

4.24. Flow Cytometry

To evaluate dendritic cell (DC) maturation, cell suspensions were prepared from tumor tissues using a murine Tumor Dissociation Kit (Miltenyi Biotec, Germany) as per the manufacturer's instructions. The dissociated cells were filtered through a 70 µm strainer and subjected to density gradient centrifugation using 40% Percoll (Cytiva) to enrich for immune cells. Cells were then stained with antibodies, including APC‐anti‐mouse CD86 (1:250), APC‐Cy7‐anti‐mouse MHC‐I (1:250; eBioscience, USA), PE‐Cy7‐anti‐mouse CCR7 (1:250), PE‐anti‐mouse CD11c (1:500), FITC‐ anti‐mouse CD45 (1: 1000), BV421‐anti‐mouse MHC‐II (1:500), and BV650‐anti‐mouse XCR1 (1:250; BioLegend, USA) on ice for 30 min. After washing with PBS, cells were analyzed by flow cytometry (FACSVerse, BD, USA).

4.25. Immunohistochemistry and Histology

For immunohistochemistry, antigens were retrieved by boiling tissue sections in sodium citrate buffer, and the endogenous peroxidase activity was blocked by incubation in 3% hydrogen peroxide solution. The paraffin sections (5 µm) were incubated with PBS buffer containing 3% bovine serum albumin (BSA) for 30 min at RT to block nonspecific binding, followed by staining with rabbit primary antibodies including anti‐Cleaved Caspase‐3 (1:500; 9661); anti‐Cleaved Caspase‐9 (1:200; 9509), anti‐CD8α (1:500; 98941), anti‐Foxp3 (1:500; 12653, Cell Signaling Technology, USA) or anti‐Ki67 (1:1000, ab15580, Abcam, USA) at 4(C overnight. Subsequently, sections were thoroughly washed and incubated with goat anti‐rabbit IgG‐HRP (1:1000; BM3894, Boster, Wuhan, China) for 1 h at RT. These sections were stained with 3,3′‐diaminobenzidine (DAB) staining (HY‐W025920, MedChemExpress, USA) and counterstained with hematoxylin (ST2067, Beyotime, China) for 1 min, followed by visualization under a light microscope (BX51, Olympus, Japan). For staining Cytochrome C (Cyt C), 8 µm cryosections were fixed with 4% paraformaldehyde and blocked with PBS containing 5% BSA, followed by staining with rabbit primary Cytochrome C antibody (1:200; 11940, Cell Signaling Technology, USA) and then incubated with AlexaFluor594‐conjugated goat anti‐rabbit IgG (1:200; A‐11037, Thermo Fisher Scientific, USA) after washing. Nuclei were counterstained with 4,6‐ diamino‐2‐phenyl indole (DAPI) and samples were analyzed by fluorescence confocal microscopy (LSM700, Zeiss, Germany). For Hematoxylin and Eosin (H&E) staining, the heart, liver, spleen, lung, kidney, and tumor tissues were embedded in paraffin and cut into 5 µm thick slices and subjected to H&E (C0105, Beyotime, China). For cathepsin B (CTSB) staining, HL7702 or MHCC‐LM3 cells were treated with 100 µm RS‐FS for 48 h and fixed with 4% PFA for 30 min at 4°C, followed by permeabilization with 0.1% Triton X‐100 for 30 min and blocking with 5% goat serum for 2 h at RT. Subsequently, cells were stained with a primary rabbit monoclonal antibody to CTSB (1:250; ab227811, Abcam, USA) at 4°C overnight, and then stained with AlexaFluor594‐conjugated goat anti‐rabbit IgG (1: 200; A‐11037, Thermo Fisher Scientific, USA) in PBS with Tween 20 (PBST) for 1 h at RT, followed by counterstaining with DAPI for 15 min at RT. Cells were photographed using a confocal laser scanning microscope (LSM700, Zeiss, Germany).

4.26. Reactive Oxygen Species (ROS) Detection

Levels of ROS in frozen HCC sections were measured with a DHE assay kit (S0064, Beyotime, China). Briefly, sections were incubated with 1X DHE working solution at 37°C for 20 min, followed by washing with PBS, and mounted with neutral gum (G8590, Solarbio, China). DHE fluorescence (Ex/Em = 535/610 nm) was visualized by confocal microscopy (FV1000, Olympus, Japan) and quantified with ImageJ (NIH, Bethesda, USA).

4.27. Plasma mtDNA Quantification

Circulating cell‐free mitochondrial DNA (mtDNA) was isolated from mouse plasma using the QIAamp DNA Blood Mini Kit (51104, Qiagen, Germany) and quantified by qPCR. Amplification was performed with SYBR Green Master Mix (04913914001, Roche, Swiss Confederation) using specific primers for the mitochondrial ND1 gene F: 5’‐CTAGAAACCCCGAAACCAAA‐3’; R: 5’‐CCAGCTATCACCAAGC TCGT‐3’) and the nuclear reference gene GAPDH F: 5’‐TTGGGTTGTACATCC AAGCA‐3’; R: 5’‐AACCTGCAGCCATCAGCTA‐3’. Relative mtDNA levels were normalized to GAPDH and calculated using the 2^(−ΔΔCt) method.

4.28. TUNEL Assay

Apoptotic cells in frozen tissue sections were identified by detecting DNA fragmentation with the TUNEL Apoptosis Assay Kit (T2130, Solarbio, China). Briefly, fresh‐frozen sections were air‐dried and fixed in 4% paraformaldehyde for 30 min at RT. After permeabilization with 0.1% Triton X‐100 in PBS for 10 min, sections were incubated with the TUNEL working solution (prepared by mixing Component A and Component B as per the kit's instructions) in a humidified chamber for 60 min at 37°C in the dark. Following incubation, sections were washed thoroughly with PBS to remove unincorporated reagents. Nuclei were counterstained with Hoechst (Component C diluted in PBS). The stained sections were mounted with an anti‐fade mounting medium and visualized under a fluorescence microscope (IX70, Olympus, Tokyo, Japan).

4.29. Statistical Analysis

All data are reported as mean values ± standard error of the mean (SEM). Sample size was determined by PASS software (version 11; NCSS, UT, USA). Normality of data distribution was assessed using the Shapiro‐Wilk test, and equality of variances was evaluated using Levene's test to guide the selection of parametric or nonparametric methods. Outliers were identified using the Grubbs' test. For comparisons involving two groups, either the two‐tailed Student's t‐test (parametric) or the Mann‐Whitney rank sum test (nonparametric) was used. For comparisons involving more than two groups, one‐way or two‐way analysis of variance (ANOVA) was applied for parametric data, followed by post‐hoc multiple comparisons using the Student‐Newman‐Keuls method. For nonparametric data, the Kruskal‐Wallis test was used, followed by Dunn's post‐hoc test for pairwise comparisons. Specific statistical tests used for each experiment were also indicated in the corresponding figure legends. All statistical analyses were performed using SigmaStat (version 3.5; Systat Software, Inc., USA), with statistical significance set at a two‐sided alpha level of p < 0.05. The sample size (n) for each experiment, all of which were ≥3, was also specified in the corresponding figure legends.

Author Contributions

H.Y. and R.J. designed the project. R.J., X.K., J.Z., J.L., X.H., Z.Y., L.Z., and Q.W. carried out the experiments. Y.W. helped with the diagram and graphical abstract drawing. Y.S. helped with writing. R.J. and H.Y. analyzed the data. H.Y. and R.J wrote the paper with the input from all authors.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Supporting File: adma73468‐sup‐0001‐SuppMat.docx.

Data Availability Statement

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