Lysosome-targeting endocytosis of NPsThe inherent susceptibility of NPs to lysosome-targeting endocytosis underlies the MONOTAB concept. To validate this rationale, we first explored the lysosomal delivery of NPs. A panel of cells (including B16F10, CT26, SKOV3, and MCF-7) was incubated with rhodamine B-labeled NPs (RBNPs, Supplementary Fig. 1a) for 2 or 4 h, washed and then analyzed by flow cytometry. The results revealed that over 90% of cells in each cell line were fluorescently positive after 4-h incubation (Fig. 1b). Moreover, we confirmed that nearly all the detected NPs were internalized into cells but not absorbed on the membrane via Trypan Blue quenching (Supplementary Fig. 1b and c). The fluorescence intensity measurements of RBNPs in the medium following co-incubation showed that ~22% of the given NPs were internalized within 2 h and ~28% within 4 h (Supplementary Fig. 1d and e). We further examined the subcellular distribution of RBNPs after 4 h incubation using confocal microscopy, which showed high-degree colocalization between the RB signal and lysosomes (Fig. 1c and Supplementary Fig. 1f), with Manders’ colocalization coefficients consistently exceeding 0.8 or even 0.9 (Supplementary Fig. 1g). Notably, despite distinct membrane proteomes among these cell lines22,23, minor variations in NP uptake and lysosomal localization were observed, implying that the lysosome-targeting endocytosis of NPs does not rely on specific surface receptors19.To identify the critical pathways involved in NP internalization, we next performed a set of endocytosis inhibition experiments. Cells were pre-treated with chemical endocytosis inhibitors or exposed to low temperatures (4 °C), followed by incubation with RBNPs for 1.5 h. Flow cytometry analysis showed that low-temperature treatment significantly inhibited RBNP uptake across all cell lines, pointing to an energy-dependent uptake process. Cholesterol sequestration (filipin)24 or inhibition of clathrin-coated pit (CCP) dynamics (Pitstop 2)25,26 did not influence NP uptake, while the tyrosine kinase inhibitor (genistein)27, actin polymerization inhibitor (cytochalasin D)28,29, and PI3K signaling inhibitor (wortmannin)30,31 showed mild inhibitory effects in a cell line-specific manner (Fig. 1d, Supplementary Fig. 2a and b). Notably, chlorpromazine significantly reduced NP uptake in all cell lines. Since chlorpromazine inhibits clathrin-mediated endocytosis but not specifically21, we used siRNA (small interfering RNA) to knock down clathrin (Supplementary Fig. 2c) to confirm the involvement of clathrin in the internalization process. This knockdown resulted in a significant decrease in NP uptake by ~60% (Supplementary Fig. 2d). These results suggest a major role of clathrin-mediated endocytosis in NP internalization.To ascertain the independence of NP uptake from specific surface receptors, we screened a panel of typical receptors associated with lysosomal trafficking, including insulin-like growth factor 2 receptor (IGF2R)4, asialoglycoprotein receptor (ASGPR)5,11,12, integrin αvβ3 (ITGAV:ITGB3)7, atypical chemokine receptor 3 (ACKR3/CXCR7)8, scavenger receptor (SCARB1)9, glucagon-like peptide 1 receptor (GLP-1R)32, transferrin receptor (TFRC)33, low-density lipoprotein receptor (LDLR)34, glucosylceramidase beta (GBA)35, and vesicle transport through interaction with t-SNAREs (VTI)36. Since Asgpr and Glp-1r are minimally expressed in B16F10 cells, we focused on the remaining to identify genes for which knockdown may ablate NP uptake. Targeted gene silencing via CRISPR interference (Supplementary Fig. 3a) or RNA interference (Supplementary Fig. 3b) did not impact the internalization of RBNPs (Fig. 1e and f), hinting at the independence of specific receptors. To further confirm this observation, we subjected B16F10 cells to trypsin treatment for 3 h to digest extracellular domains of membrane proteins. Subsequent co-incubation with RBNPs revealed only a mild decrease in the mean fluorescence intensity (MFI) of cells, with no change in the ratio of RB-positive cells (Fig. 1g). In contrast, co-incubation with biotinylated cRGD (biotin-cRGD) and Cy3Avidin (Supplementary Fig. 4), which are internalized via integrin-mediated endocytosis, showed a substantial reduction in the delivery of Cy3Avidin by biotin-cRGD (Supplementary Fig. 4). These results strongly suggest that NP uptake does not rely on specific surface receptors.Construction of the MONOTAB platformTo ease the preparation of MONOTAB and enable high modularity, we sought to develop an antibody mounting platform for the convenient installation of specific antibodies. Commercially available streptavidin-conjugated anionic NPs were chosen as the chassis, which permits the anchoring of biotinylated anti-IgG (Fc specific) antibodies (αFc) onto the NPs through the streptavidin-biotin interaction. The preparation involves simply mixing the NPs with biotinylated αFc and then centrifugation. As the Fc regions of IgG antibodies from the same host species are identical, the resulting αFc-tethered NPs (αFc-NPs), which serve as the antibody mounting platform, can specifically recognize and immobilize any targeted antibody containing the Fc fragment, allowing for rapid production of diverse MONOTABs towards various targets.To demonstrate the feasibility of this approach, we incubated NPs with a biotinylated anti-IgG antibody at a 1:1 molar ratio of streptavidin to biotin. Dynamic light scattering (DLS) measurements showed that the average hydrodynamic diameter of the resulting αFc-NPs was ~120 nm, ~20 nm larger than that of the NPs (Supplementary Fig. 5a and b). Transmission electron microscopy (TEM) and scanning electron microscopy (SEM) confirmed the particle size and the uniformly spherical structure of both NPs and αFc-NPs (Supplementary Fig. 5c and d). To verify the presence of αFc on NPs, we ran a reducing SDS-PAGE gel with αFc-NPs, αFc, and NPs (Fig. 2a). After Coomassie blue staining, the αFc-NP lane showed bands at ~17, ~25, ~50, and ~75 kDa (Fig. 2b), which correspond to monomeric streptavidin, the light chain of αFc, the heavy chain of αFc, and monomeric immunoglobulin, respectively, confirming the anchoring of αFc on the NP surface. Furthermore, the αFc-NPs demonstrated excellent stability, as evidenced by the negligible change in particle size over 2-week storage at 4 °C (Supplementary Fig. 5e).Fig. 2: Antibody anchoring does not alter the lysosome-targeting ability of NPs.a Putative fragments of αFc-NP, αFc, and NP yielded after protein denaturation with 2-mercaptoethanol. b Reducing SDS-PAGE gel of αFc-NP, αFc, and NP with Coomassie blue staining. The gel images are representative of n = 3 independent replicates. c TEM images of cells incubated with αFc-NP (50 μg mL−1) for 8 h. The TEM images are representative of n = 3 independent replicates. d Live-cell images of B16F10, CT26, SKOV3, or MCF-7 cells treated with αFc-RBNP (100 μg mL−1) for 4 h, respectively. Scale bar, 10 μm. The images are representative of n = 3 biological replicates. e Left panel: live-cell images of different cell lines treated with RB/Cy5CTRL-NPs (50 μg mL−1) for 10 h. Scale bar, 10 μm. Right panel: Fluorescence intensity profiles along the line drawn in the box of the left panel. The images are representative of n = 3 biological replicates. f Left panel: live-cell images of B16F10 cells captured at the indicated time points after incubation with Cy5CTRL-NPs (50 μg mL−1) for 10 h and then cell washing. Scale bar, 10 μm. Right panel: Fluorescence intensity profiles along the line drawn in the box of the left panel. The images are representative of n = 3 biological replicates. g Western blots (left) and quantitative analysis (right) of EEA1, LAMP1, and RAB7 in B16F10 cells after incubation with 50 μg mL−1 CTRL-NPs or NPs for 10 h (n =  3 biologically independent experiments). Data are presented as mean ± SD where relevant. P values were determined by one-way ANOVA with Dunnett’s post hoc test. ns, no significance; *p < 0.05; ***p < 0.001. Source data are provided as a Source Data file.We next investigated whether the surface anchoring of αFc would alter the biological performance of the NPs, particularly the lysosome-targeting endocytosis. The cell counting kit-8 (CCK8) assay proved the minimal cytotoxicity of both NPs and αFc-NPs to B16F10 cells at serial concentrations ranging from 0.01 ng mL−1 to 100 μg mL−1 (Supplementary Fig. 5f). Flow cytometry analysis showed that the endocytosis in B16F10 cells was concentration-dependent for both NPs and αFc-NPs (Supplementary Fig. 5g), and the uptake efficiency of αFc-RBNP was comparable to that of RBNP after co-incubation for 4 or 8 h, reaching 87% and 96%, respectively (Supplementary Fig. 5h). We examined the cell samples with negative-stain electron microscopy and observed abundant accumulation of spherical nanoparticles within membrane-enclosed compartments (Fig. 2c), which were subsequently identified as lysosomes through live-cell confocal microscopy (Fig. 2d, Supplementary Fig. 5i and j). The endocytosis inhibition experiments indicated that clathrin-mediated endocytosis contributed the most to αFc-NP internalization, which was consistent with the findings of NPs (Supplementary Fig. 5k). These results demonstrate that the inherent lysosome-targeting endocytosis of NPs was not altered by the surface modification with αFc.We next incubated αFc-NPs with an IgG control antibody produced from the matching host species to generate control MONOTAB (CTRL-NP). At a 1:1 molar ratio of IgG:αFc, the average hydrodynamic diameter of CTRL-NPs was ~30 nm larger than that of αFc-NP, suggesting the successful immobilization of the IgG control onto αFc-NPs (Supplementary Fig. 6a and b). Cryo-TEM confirmed the size, morphology, and well-dispersed state of CTRL-NPs (Supplementary Fig. 6c and d). We then asked if this layer of antibody would influence the uptake efficiency. To this end, we prepared a series of CTRL-NPs with molar ratios of αFc:IgG varying from 1:0.1 to 1:2 (Supplementary Fig. 6e) and then incubated them with different cell lines for 4 h. Flow cytometry analysis showed that the varying αFc:IgG ratios did not impact NP uptake in all cell lines (Supplementary Fig. 6f). Furthermore, we confirmed that the internalization of these CTRL-NPs did not need the interactions between IgG and Fc receptors (FcRs) by using FcR-preblocked B16F10 cells (Supplementary Fig. 6g). To enable more POI binding, we selected the αFc:IgG ratio of 1:2 in the following studies.Next, we proceeded to investigate whether the CTRL-NPs retained the lysosome-targeting ability. Cy5-labeled IgG (Cy5IgG) was immobilized on the surface of αFc-RBNPs to form RB/Cy5CTRL-NPs. Co-incubation of cells with RB/Cy5CTRL-NPs led to the colocalization of Cy5IgG and RBNPs within the lysosomes (Fig. 2e and Supplementary Fig. 6h). To ensure that the lysosomal delivery of Cy5IgG occurred specifically due to its binding to αFc-NPs, we incubated B16F10 cells with Cy5IgG alone, Cy5IgG plus αFc or NPs, or Cy5IgG plus αFc-NPs (namely Cy5CTRL-NPs). A remarkable 660-fold increase in cellular fluorescence was observed when Cy5IgG was co-incubated with αFc-NPs, whereas no increase was observed when co-incubated with αFc or NPs (Supplementary Fig. 7a, b). Furthermore, even after extended incubation for an additional 8 or 24 h following washing, the Cy5 signal remained localized within the lysosomes (Fig. 2f). These data indicate that NPs could efficiently hijack the tethered IgG into lysosomes and the tethered protein could be stably trapped without lysosomal escape.Effects of MONOTAB on lysosomal functionAfter co-incubation with Cy5CTRL-NPs, we observed an unexpected increase in the LysoTracker signal (Supplementary Fig. 7a and c). This observation raises the possibility that MONOTAB might promote lysosomal biogenesis. To test this hypothesis, we examined the expression levels of endo-lysosome markers, including LAMP1 (lysosome), EEA1 (early endosome), and RAB7 (late endosome), and found that all the tested markers were upregulated after the CTRL-NP treatment (Fig. 2g). Immunofluorescence assay further confirmed the increase of LAMP1 (Supplementary Fig. 7d and e). These results are consistent with a previous report indicating that internalization of anionic polystyrene nanoparticles results in activation of the transcription factor EB, a master regulator of lysosome biogenesis, and increased lysosomal degradation capacity37.As an increase in LysoTracker staining or lysosome markers may also be observed upon lysosomal dysfunction, one may question if the nanoparticles could potentially impair lysosomal health, thereby activating lysosome biogenesis as a compensatory response. To clarify this question, we performed the DQ Green BSA assay to evaluate lysosomal degradation capacity. Untreated B16F10 cells and cells treated with NPs, αFc-NPs, or CTRL-NPs exhibited bright green fluorescence, indicating the effective hydrolysis of the DQ Green BSA into single, dye-labeled peptides by lysosomal proteases. In contrast, no fluorescent signal was observed in cells treated with Bafilomycin A1 (BafA1), an established lysosomal inhibitor (Supplementary Fig. 7f and g). We also examined lysosome membrane stability with acridine orange (AO), a fluorescent dye that emits red fluorescence when protonated in intact lysosomes and green fluorescence when deprotonated in the cytoplasm. Strong green fluorescence was detected in cells treated with chloroquine (CQ, a lysosome-permeability enhancer), while untreated cells and cells treated with NPs, αFc-NPs, or CTRL-NPs exhibited red fluorescence only (Supplementary Fig. 7h and i). These results imply that MONOTAB may promote lysosomal biogenesis without affecting lysosomal health, which promises higher protein degradation potential.MONOTAB-mediated degradation of membrane-associated protein PD-L1We next aimed to evaluate the efficacy of the MONOTAB strategy on clinically relevant targets. We first targeted programmed death-ligand 1 (PD-L1), a membrane-associated protein that is often overexpressed on the surface of tumor cells and facilitates their immune evasion38,39. PD-L1-targeted MONOTABs were constructed by incubating αFc-NP with either FITC-labeled or unlabeled anti-PD-L1 antibody (αPD-L1). Co-incubation of cells with FITCαPD-L1-NPs (Fig. 3a) led to lysosomal delivery of FITCαPD-L1 (Fig. 3b and Supplementary Fig. 8a), replicating our findings with CTRL-NPs. We reasoned that αPD-L1-NPs could induce the degradation of PD-L1 by enriching PD-L1 molecules in lysosomes. B16F10 cells were treated with PBS, αPD-L1, αFc-NP, or αPD-L1-NP for 24 h and then assayed for PD-L1 levels. Western blot analysis showed substantial degradation of both total and membrane-associated PD-L1 with 3.3 nM (αPD-L1-equiv. concentration) of αPD-L1-NP (Fig. 3c and d). Immunofluorescence (IF) microscopy revealed nearly complete removal of PD-L1 from cell membranes following the 24-h treatment with αPD-L1-NP, as opposed to the treatments with PBS or αPD-L1 alone (Fig. 3e and Supplementary Fig. 8b). These results highlight the great potential of MONOTAB in inducing robust degradation of membrane proteins.Fig. 3: Degradation of PD-L1 mediated by αPD-L1-NP.a Schematic illustration of live-cell confocal microscopy assay. b Live-cell images of B16F10 cells treated with FITCαPD-L1 (3.3 nM) or FITCαPD-L1-NP (FITCPD-L1-equiv., 3.3 nM) for 4 h. Scale bar, 10 μm. The images are representative of n = 3 biological replicates. c, d Western blot analysis of total (c) and membrane-associated (d) PD-L1 in B16F10 cells receiving different treatments for 24 h. αPD-L1-equiv. concentration, 3.3 nM; NP-equiv. concentration, 25 μg mL−1. The blots are representative of n = 3 biological replicates. e IF of surface PD-L1 in B16F10 cells treated with αPD-L1 (3.3 nM) or αPD-L1-NP (PD-L1-equiv., 3.3 nM) for 24 h. Scale bar, 20 μm. The IF images are representative of n = 3 biological replicates. f Western blot analysis of B16F10 cells treated with αPD-L1-NP (3.3 nM) for 12 or 24 h in the presence or absence of 0.1 mg mL−1 leupeptin (LPT). The blots are representative of n = 3 biological replicates. g–j Western blot analysis of PD-L1 in B16F10 cells treated with αPD-L1-NP at the indicated concentrations for 24 h (g and h) or at 3.3 nM for the indicated durations (i and j) The blots are representative of n = 3 biological replicates. k–o In vivo antitumor study of αPD-L1-NP in B16F10 tumor-bearing C57BL/6 mice. Mice (n = 5 mice per group) were treated intratumorally (i.t.) with PBS, αPD-L1 (2.0 mg kg−1), or αPD-L1-NP (αPD-L1-equiv. dose, 2.0 mg kg−1) for three times, respectively. k Schematic diagram outlining the experimental design. l Tumor growth curves of mice receiving different treatments. m Image of tumors resected after animal euthanasia. n Immunofluorescence staining of PD-L1 in tumor sections. Scale bar, 100 μm. o Western blot analysis of PD-L1 levels in tumor lysates. The blots are representative of n = 3 biological replicates. All the uncropped blots are included in the Source Data file. Data are presented as mean ± SD. Statistical significance was calculated via one-way ANOVA with Dunnett’s post hoc test. **p < 0.01; ***p < 0.001. Source data are provided as a Source Data file.To confirm whether the degradation occurs in lysosomes, B16F10 cells were treated with αPD-L1-NPs in the presence or absence of leupeptin (LPT), a commonly used lysosomal protease inhibitor. The LPT treatment significantly diminished the degradation of PD-L1, indicating the involvement of lysosomal proteases in the MONOTAB-mediated degradation (Fig. 3f). Further data of total PD-L1 levels following the 24-h treatment with different concentrations of αPD-L1-NP unveiled a concentration-dependent degradation profile without the hook effect (Fig. 3g and h), echoing the monofunctional modality of MONOTAB. Notably, significant degradation of PD-L1 was already detectable with αPD-L1-NP at a subnanomolar concentration (0.7 nM) and virtually complete elimination was achieved at 6.7 nM. Time-course experiments showed that MONOTAB-mediated PD-L1 degradation occurred in a time-dependent manner, with the levels of PD-L1 persistently decreasing to complete depletion by 48 h (Fig. 3i and j). These results indicate that the MONOTAB approach can efficiently direct surface proteins to lysosomes for degradation.To validate our approach further, we conducted a comprehensive comparison between MONOTAB and existing approaches using published data on PD-L1 degradation (Supplementary Table 1). Additionally, we benchmarked MONOTAB against two established methods based on bifunctional chimeras, integrin-facilitated lysosomal degradation (IFLD) and GalNAc-LYTAC. PD-L1-targeted BMS-L1-RGD (IFLD type) and αPD-L1-GalNAc (GalNAc-LYTAC type) were synthesized following the described procedures5,7. As opposed to the near-complete degradation of PD-L1 observed with 6.7 nM of αPD-L1-NP, treatment with 50 nM of BMS-L1-RGD for the same duration led to only ~46% of PD-L1 degradation (Supplementary Fig. 8c). On the other hand, to ensure a fair comparison between αPD-L1-NP and αPD-L1-GalNAc, considering GalNAc’s avid binding to ASGPR predominately expressed on hepatocytes, we used Hepa1–6 cells, a murine hepatoma cell line, as the cell model. At a low concentration of 1.3 nM, both αPD-L1-NP and αPD-L1-GalNAc induced similar levels of PD-L1 degradation. However, at higher concentrations, αPD-L1-GalNAc exhibited a typical hook effect due to its bifunctional nature, while αPD-L1-NP caused even more substantial degradation (Supplementary Fig. 8d).Next, we evaluated the in vivo antitumor effects of αPD-L1-NP. C57BL/6 mice bearing subcutaneous B16F10 tumors were treated with PBS, αPD-L1, or αPD-L1-NP, respectively, and tumor size was measured (Fig. 3k). Compared with the control and αPD-L1 groups, tumor growth was significantly inhibited by the treatment with αPD-L1-NP (Fig. 3l, m and Supplementary Fig. 8e) and no body weight loss was observed during the experiment (Supplementary Fig. 8f). Immunofluorescence analysis of PD-L1 expression in tumor sections revealed a markedly reduced PD-L1 level in the αPD-L1-NP group (Fig. 3n and Supplementary Fig. 8g), which was further corroborated by Western blot analysis (Fig. 3o and Supplementary Fig. 8h). These results underscore the therapeutic potential of MONOTAB in vivo.MONOTAB-mediated degradation of secreted protein MMP2Having demonstrated the target scope of MONOTAB towards the membrane-associated protein, we next proceeded to assess its efficacy in degrading secreted proteins. Matrix metalloproteinase 2 (MMP2), which is highly expressed in various tumors and crucial for tumor invasion and metastasis40,41, was chosen as the target protein. The anti-MMP2 MONOTAB (αMMP2-NP) was prepared similarly. B16F10 cells were treated with PBS, αMMP2, or αMMP2-NP for 12 h, and the culture media were then assayed for the MMP2 activity and content (Fig. 4a). Gelatin zymography revealed a substantial decrease in MMP2 activity following the αMMP2-NP treatment. Given that the treatment with αMMP2 also mildly reduced MMP2 activity relative to PBS control, to rule out the possibility that the observed decrease by αMMP2-NPs was exclusively due to the inhibition of MMP2 catalytic activity, we further ran Western blot to quantify the MMP2 content in the media. Similarly, a significant reduction in MMP2 content was observed with αMMP2-NPs. Moreover, αMMP2-NPs substantially elevated the intracellular MMP2 content, especially upon the inhibition of lysosomal proteases with LPT (Fig. 4b), demonstrating the ability of MONOTAB to redirect secreted proteins into lysosomes for degradation. Further analysis of MMP2 activity and content following the treatment with varying concentrations of αMMP2-NP demonstrated a concentration-dependent degradation profile without the hook effect (Fig. 4c). Similar results were observed in other cell lines (Supplementary Fig. 9a and b).Fig. 4: Degradation of MMP2 mediated by αMMP2-NP.a Gelatin zymography and Western blot assay of cell culture media for MMP2 activity and content. Media were collected after B16F10 cells were treated with PBS, αMMP2 (12 nM), or αMMP2-NP (α-MMP2-equiv. 12 nM) for 12 h. The gels and blot are representative of n = 3 biological replicates. b Western blot assay of MMP2 inside (IN) or outside (OUT) of B16F10 cells treated with αMMP2-NP (α-MMP2-equiv. 12 nM) for 12 h in the presence or absence of 0.1 mg mL−1 LPT. The gel and blots are representative of n = 3 biological replicates. c MMP2 activity and content in the culture media of B16F10 cells treated with varying concentrations of αMMP2-NP for 12 h. The gels and blot are representative of n = 3 biological replicates. d, e Schematic illustration (d) and results (e) of the wound-healing assay. B16F10 cells were treated with PBS, αMMP2 (12 nM), or αMMP2-NP (α-MMP2-equiv. 12 nM) for 12 h. Scale bar, 200 μm. The images are representative of n = 3 biological replicates. f, g Schematic illustration (f) and results (g) of the transwell cell invasion assay. CT26 cells were treated with PBS, αMMP2 (12 nM), or αMMP2-NP (α-MMP2-equiv. 12 nM). Scale bar, 200 μm. These images are representative of n = 3 biological replicates. All the uncropped gels and blots are included in the Source Data file. Source data are provided as a Source Data file.Considering that MMP2 degrades a wide range of extracellular matrix components and facilitates cell migration42, we next investigated whether MONOTAB-mediated MMP2 degradation could translate into decreased cell mobility. A wound-healing assay was performed to evaluate the effect of αMMP2-NP on cell migration. B16F10 cells were pre-incubated in serum-free media for 12 h, and the media were collected and re-applied to the cells along with PBS, αMMP2, or αMMP2-NP after wound creation and wash (Fig. 4d). To eliminate the impact of cell proliferation on wound healing, the cells were pre-treated with mitomycin C, a mitotic inhibitor. The αMMP2-NP treatment led to a significantly slower scratch closure rate than observed with αMMP2 or PBS (Fig. 4e), demonstrating the high potency of αMMP2-NP in inhibiting cell mobility.To further validate our findings, we carried out the cell invasion assay using a transwell apparatus. The serum-free culture media collected after different treatments were incubated in the apical chambers coated with Matrigel for 24 h at 37 °C, followed by cell seeding in the apical chambers with a fresh serum-free medium. A full medium was added to the basolateral chambers (Fig. 4f). Following 12-hour incubation, cells beneath the membrane of the inner chamber were visualized with crystal violet staining. In contrast to PBS or αMMP2, the αMMP2-NP treatment essentially prohibited cell migration towards the basolateral side of the membrane, persisting for at least 24 h (Fig. 4g). This observation was consistent with the results in the wound-healing experiment, further confirming the functional cellular consequences produced by MONOTAB-mediated protein degradation.MONOTAB-mediated degradation of extracellular vesiclesInspired by the potent ability of MONOTAB to induce extracellular protein degradation, we next asked whether the target range could be extended to non-protein targets such as extracellular vesicles (EVs). EVs are nanoscale lipid-bound vesicles released by cells and play crucial roles in intercellular signaling and pathological processes. Despite their emerging significance as therapeutic targets, selective degradation of EVs has yet to be achieved. Given the lack of EV-specific proteins for targeting, we explored the possibility of using phosphatidylserine (PS), a molecule commonly exposed on the outer leaflet of EVs but not on viable cells43, as a target for designing the EV-targeted MONOTAB. In light of the strong interaction between PS and Annexin V, we prepared Annexin V-NPs for EV capture by incubating NPs with biotinylated Annexin V. DLS measurement confirmed the binding of EVs to Annexin V-NPs, which resulted in an enlarged particle size (~274 nm) after co-incubation (Supplementary Fig. 10a).Next, we investigated if Annexin V-NPs could facilitate the uptake of EVs into cells. To ease the detection, we used an ECDHCC1-PalmGRET stable cell line to produce EGFP-labeled EVs (EGFPEVs), which emitted stable fluorescence at pH as low as 5.0 (Supplementary Fig. 10b). Co-incubation of B16F10 cells with Annexin V-NPs and EGFPEVs resulted in a ~70-fold increase in EV uptake compared to EGFPEVs alone, Annexin V plus EGFPEVs, or NPs plus EGFPEVs (Fig. 5a and b). Moreover, co-incubation with Annexin V-NPs significantly reduced the fluorescence intensity of EGFPEVs in the medium (Fig. 5c), approaching the blank level. Colocalization analysis demonstrated that the internalized EVs were transported to lysosomes as well. Similar results were also observed in other cell lines (Supplementary Fig. 10c and d). To ensure that the internalized EVs were indeed trapped in but not fused with lysosomes, we examined the cell samples using negative-stain electron microscopy. As demonstrated in Fig. 5d, treatments with Annexin V-NPs and EGFPEVs showed intact EVs closely associated with NPs within lysosomes. Conversely, when treated with NPs plus EGFPEVs, only NPs were observed within lysosomes. This observation indicates that Annexin V-NPs effectively hijacked EVs into cells and transported them to the lysosomes.Fig. 5: Degradation of extracellular vesicles mediated by Annexin V-NP.a Live-cell cell images of B16F10 cells incubated with EGFPEVs, Annexin-V + EGFPEVs, NPs + EGFPEVs, or Annexin-V-NPs + EGFPEVs for 8 h. The concentrations of EV, Annexin-V, and NP were 1.3 × 108 particles mL−1, 0.4, and 50 μg mL−1, respectively. Scale bar, 10 μm. The images are representative of n = 3 biological replicates. b Fold changes in MFI of EGFR signal in a (n = 3 biologically independent experiments). c Fluorescence intensity of EGFPEVs in the medium after co-incubation for 8 h (n = 6 biological replicates). d Representative TEM images of cells incubated with NPs + EVs or Annexin-V-NPs + EVs for 8 h. Scale bars are defined on the panel. White arrow points to the internalized EVs. The images are representative of n = 3 biological replicates. e Fluorescence intensity measurement of exocytosed EGFPEVs in the medium (n = 5 biological replicates). f Real-time tracking of EGFP signal in individual cells after incubation with EVs (1.3 × 108 particles mL−1) and Annexin V-NPs (50 μg mL−1) for 8 h. Scale bar, 10 μm. Cells 1–3 represent randomly tracked n = 3 single cells. g Fluorescence intensity of EGFP signal in individual cells at the indicated time points. Data are presented as mean values ± SD where relevant. Statistical significance was calculated via one-way ANOVA with Dunnett’s post hoc test. ns, no significance; **p < 0.01; ****p < 0.0001. Source data are provided as a Source Data file.Afterward, we aimed to clarify the fate of the internalized EVs. We first explored the possibility of exocytosis. B16F10 cells treated with EGFPEVs, Annexin-V plus EGFPEVs, NPs plus EGFPEVs, or Annexin-V-NPs plus EGFPEVs were further cultured in fresh serum-free medium. Following incubation for 10 h, the fluorescence intensity of EGFP in the medium was measured, which showed no significant difference across all groups compared to the EGFPEV-free group (Fig. 5e). This suggests that the internalized EVs may not undergo exocytosis. We next asked whether the endocytosed EVs could be degraded. B16F10 cells were incubated with Annexin V-NPs and EGFPEVs for 8 hours, followed by washing for real-time imaging with time-lapse confocal microscopy. We assumed that EGFPEV degradation would be accompanied by the degradation of EGFP, and thus, the diminishment of EGFP signal within the cells could reflect the degree of EV degradation. Live-cell imaging indeed showed a time-dependent decrease in intracellular EGFP signal (Fig. 5f and g), indicating EV degradation. To rule out the contribution of photobleaching, we tracked the EGFP signal in ECDHCC1-PalmGRET cell debris under continuous laser exposure, and only minimal change was detected (Supplementary Fig. 10e and f). These results underscore the potential of using the MONOTAB platform for selective degradation of non-protein targets such as EVs, opening up possibilities for targeting intercellular communication mechanisms mediated by these vesicles.