2-Deoxyglucose Dendrimer-Enabled Niclosamide Delivery to FRβ-Expressing Macrophages Alleviates Endometriosis Progression and Associated Hyperalgesia

A.S. and K.H. designed the research; A.D., A.S., and R.S. designed and synthesized the dendrimers; M.S., M.H., and T.P. performed the animal experiments and/or analyzed the data; A.D., K.J.G., and A.I.D. performed quantitative biodistribution analysis; N.S.M., V.K.S., and P.S.C. collected/or prepared the human samples; M.S. performed single‐cell RNA sequencing and analysis; A.D., M.S., R.S., A.S., and K.H. wrote the paper; all authors read, reviewed, edited, and approved the manuscript.

All animal experiments were conducted at Washington State University in accordance with NIH guidelines for care and use of laboratory animals (protocol #6751).

This work was supported by NIH grant R01HD104619 (to Kanako Hayashi ) and a Washington Research Foundation Technology Commercialization Phase 1 award (to Kanako Hayashi and Anjali Sharma ).

All procedures involving human subjects were performed in accordance with ethical approval by the Institutional Review Board (IRB) guidelines for the University of Florida under IRB code NO. IRB202003131. All patients were recruited from the Division of Minimally Invasive Gynecologic Surgery of the University of Florida Health Shands Hospital, Gainesville, Florida. Participants were either previously diagnosed with EuE or suspected of having EuE, with symptoms including ovarian cysts, chronic pelvic pain, dysmenorrhea, dyschezia, dyspareunia, uterine bleeding, and myofascial pain syndrome. EuE is further confirmed by pathological analysis of the excised lesions obtained during surgery. Informed written consent was obtained from each participant before the procedure. In this study, peritoneal washings from four patients aged 19–37 years (Table S1 ) were collected at the beginning of the procedure. Briefly, the pelvic cavity was irrigated using approximately 100 mL of sterile saline, and approximately 80 mL of peritoneal washings containing pelvic exudate cells were collected from each patient and placed on ice immediately. Red blood cells were removed by resuspending cells in 500 µL Red Blood Cell Lysis Buffer (BioLegend, Cat.#420301). Cells were then counted and cryopreserved with freezing media containing 90% fetal bovine serum (FBS) and 10% dimethyl sulfoxide, and stored at −80°C for further analysis. To perform single‐cell RNA sequencing (scRNA‐seq), frozen pelvic exudate cells were thawed in a 37°C water bath and prepared following the 10x Genomics protocol named “Fresh frozen human peripheral blood mononuclear cells for single‐cell RNA sequencing” (document #CG00039 Rev E). Briefly, thawed cells were slowly transferred into a 50 mL conical tube and recovered by sequential 1:1 dilution with warmed FBS for a total of five times. Cell viability was assessed to ensure samples had viability greater than 80%. After centrifuging at 300 g for 5 min at room temperature, the cells were resuspended in phosphate‐buffered saline (PBS) with 0.04% bovine serum albumin (BSA) at a concentration of 900–1200 cells/µL. Cells were then placed on ice and immediately proceeded to library construction using a 10x Genomics Chromium Next GEM Single Cell 3’ Kit v3.1, following the standard kit user guide (document # CG000204 ) for a target of 6000 cells. Libraries were sequenced on Illumina NovaSeq 6000. FASTQ files were aligned to the human reference genome GRCh38 via Cell Ranger v7.1.0 [ 31 ], and an average of 107 903 reads per cell was achieved for each RNA library (Table S2 ). We integrated publicly available human scRNA‐seq datasets from eutopic endometrium with or without EuE, ectopic endometrium (EcE), and pelvic exudate cells. Processed Cell Ranger output files or raw FASTQ files were downloaded from the Gene Expression Omnibus (GEO), ArrayExpress, or the Sequence Read Archive (SRA). These datasets included: (1) Wang et al. ( GSE111976 ) [ 32 ], (2) Lai et al. ( GSE183837 ) [ 33 ], (3) Garcia‐Alonso et al. (E‐MTAB‐10287) [ 34 ], (4) Tan et al. ( GSE179640 ) [ 15 ], (5) Fonseca et al. ( GSE213216 ) [ 35 ], (6) Huang et al. ( GSE214411 ) [ 36 ], (7) Marečková et al. (E‐MTAB‐14039) [ 37 ], (8) Shin et al. (PRJNA932195) [ 38 ], (9) Han et al. ( GSE228030 ) [ 39 ], and (10) Zou et al. (PRJNA713993) [ 40 ], as shown in Figure  1A,B . For all datasets, only samples collected from women of reproductive age were included. For endometrial scRNA‐seq datasets from (1) to (8), patients labeled as undergoing hormonal treatments during sample collection were also excluded. Samples used in this study are summarized in Table S3 . scRNA‐seq analysis of FOLR2 + macrophages in endometrium and pelvic exudate cells in endometriosis. (A,B) Lists of human scRNA‐seq datasets analyzed and sample size per dataset in the groups of (A) eutopic endometrium with (EuE) and without (Cont) endometriosis, EcE (lesion, EcE), and (B) pelvic exudate cells from subjects with (Endo) and without (Cont) endometriosis. (C,D) UMAP plots showing major cell types identified in the endometrium (EM) and pelvic cells (P). (E,F) Violin plot representing marker gene expression for each major cell type identified in endometrial and pelvic cell datasets. (G,H) UMAP projections showing expression patterns of FOLR2 in endometrial and pelvic cells. (I,J) Bar plots showing enriched GO biological processes in FOLR2 + macrophages from endometrium and pelvic cells. (K,L) Proportions of endometrial and pelvic FOLR2‐expressing cells to total cells within each tissue type. Differences among groups were analyzed with one‐way ANOVA followed by the Tukey multiple comparisons test. ** p < 0.01 and *** p < 0.001. (M,N) Dot plot showing representative significantly DEGs involved in proinflammatory, angiogenesis, and adhesion processes among groups of endometrium and pelvic cells (adjusted p‐value <0.05). RNA matrices from both newly generated and existing public datasets were individually imported to Scanpy v.1.8.1 for further analysis [ 41 ]. Low‐quality cells were excluded using the following criteria: (1) genes that were detected in fewer than 3 cells, (2) cells with fewer than 300 genes, (3) a maximum of 100,000 UMIs, and (4) a maximum of 20% mitochondrial content. Doublets were detected and removed using Scrublet [ 42 ]. Filtered matrices from endometrial or pelvic cell scRNA‐seq datasets were combined separately and analyzed independently. For each combined endometrial or pelvic cell scRNA‐seq matrix, gene expression values were normalized with a global scaling factor of 10,000 and log‐transformed. The top 3000, or 2000, highly variable genes for the endometrial or pelvic cell datasets, respectively, were identified by Scanpy. The mitochondrial genes, haemoglobin genes, and ribosomal genes were excluded from the highly variable gene set [ 43 ]. Principal component analysis (PCA) was performed using the highly variable gene set. After Harmony (v.0.0.10) [ 44 ] batch effect correction, Harmony‐corrected components were used to generate the neighborhood graph and perform dimensionality reduction using uniform manifold approximation and projection (UMAP). The Leiden algorithm was then used for clustering analysis [ 45 ]. Marker genes of each cluster were identified using the Wilcoxon rank‐sum test [ 46 ] with the default setting (Table S4 ). Major cell types were determined by manually annotating cell clusters with previously established marker genes from the literature [ 15 , 39 , 47 , 48 ]. To understand the transcriptional profiles of folate receptor beta‐positive (FOLR2 + ) macrophages, cells with specific FOLR2 expression (FOLR2>0) were labeled and extracted from endometrial and pelvic cell matrices. Enriched genes in FOLR2 + cells were identified by the Wilcoxon rank‐sum test in a one‐versus‐rest fashion, with a minimum log 2 (fold change) of 2 (Table S5 ). Gene Ontology (GO) analysis was performed on the identified enriched genes in FOLR2 + endometrial or pelvic cells using clusterProfiler (v.4.2.2) [ 49 ]. The enriched GO list was further filtered with the simplify function (cutoff = 0.7) to remove redundant terms (Table S6 ). Differentially expressed genes (DEGs) among eutopic endometrium without (Control; Cont) or with EuE, and EcE in the endometrial FOLR2 + macrophages, and DEGs between control (Cont) and endometriotic (Endo) pelvic exudate cells, were identified by Wilcoxon rank‐sum test, with a minimum log 2 (fold change) of 0.5 (Supplementary Table S7 ). Experimental methods related to materials, instrumentation, synthesis protocols, characterization of compounds, in vitro drug release studies, and FA‐2DG‐D‐Niclo formulation stability studies are presented in the Supporting Information (Figures S2–S34 ). C57BL/6 mice were purchased from Inotiv and housed in an environment‐controlled animal facility (20°C–22°C, ∼40% relative humidity, and 12:12 light‐dark cycle) with ad libitum access to food and water. All animal experiments were conducted at Washington State University in accordance with NIH guidelines for care and use of laboratory animals (protocol #6751). An experimental mouse model of EuE was employed, using procedures previously described and adopted by us, with minor modifications [ 50 ]. Briefly, to induce EuE‐like lesions (ELL), female donor mice were injected subcutaneously with pregnant mare serum gonadotropin (PMSG; 5 IU; Sigma, St. Louis, MO) to stimulate an estrogenic response within the uterus. Uteri were harvested from these donors 41 h after PMSG injection. The endometrium was then scraped from the myometrium and dissected into fragments (1–2 mm per side), and introduced as the source of syngeneic mouse endometrium (50 mg of fragments) via injection (in 0.2 mL of PBS) into the peritoneal cavity in the ovary‐intact recipient females under anesthesia via inhaled isoflurane. Adult female mice ( n = 36, ∼3 months old) were randomly divided into three groups ( n = 3/group/timepoint): (1) sham controls administered with Cy5‐labeled dendrimer FA‐2DG‐D‐Cy5, (2) ELL‐induced mice administered with 2DG‐D‐Cy5, and (3) ELL‐induced mice administered with FA‐2DG‐D‐Cy5. The animals in the ELL groups received donor endometrium to induce endometriotic lesions, and the sham control mice received PBS. 1 week following lesion induction, all mice received FA‐2DG‐D‐Cy5 or 2DG‐D‐Cy5 injected into the tail vein, as listed in the above group description, at a dose of 50 mg/kg (300 µL). Mice were then euthanized at 1, 6, 24, and 72 h after injection. The whole skin was gently peeled off to acquire the whole‐body Cy5 fluorescence distribution using an in vivo imaging system (IVIS; Revvity). Lesions and/or major organs, including the brain, heart, spleen, lungs, liver and kidneys, were dissected to obtain ex vivo fluorescence images and stored at ‐80°C. The fluorescence intensity of the region of interest (ROI) was calculated with the Living Image software (4.7.4) accompanying IVIS. Amounts of dendrimer in major organs and lesions were further examined ex vivo using fluorescence intensity‐based quantification. Frozen organs (lesions, liver, spleen, heart, kidneys, brain, and lungs) were slowly thawed on ice and weighed. Tissue samples were then dissected to obtain known masses from each organ. Samples were homogenized in methanol (1 mL per 100 mg tissue) with stainless steel beads using a tissue homogenizer. The homogenized samples were centrifuged at 4°C, and the clear supernatants were transferred to protein LoBind Eppendorf tubes and stored at −80°C. For fluorescence quantification, thawed supernatants were centrifuged again before analysis. Fluorescence intensity was measured on a Varioskan Lux Multimode Microplate Reader (ThermoFisher), using λ _ex = 650 nm and λ _em = 670 nm for Cy5. Background fluorescence from control solvent was subtracted, and corrected fluorescence values were converted to 2DG‐D‐Cy5 and FA‐2DG‐D‐Cy5 concentrations using calibration curves generated at 5–10 slit widths. To evaluate the specificity of cellular uptake of the dendrimer, endometriotic lesions were induced in six female mice. On Day 7 following lesion induction, mice received an intraperitoneal (i.p.) injection of 2DG‐D‐Cy5, FA‐2DG‐D‐Cy5, or FA‐2DG‐D‐Cy5‐Niclo (50 mg/kg/bw, n = 3/group). Peritoneal cells, lesions, kidney, liver, and spleen were collected 2 weeks after dendrimer injection. Specific cellular uptake of the dendrimer was then assessed by immunostaining. The efficacy of targeted delivery of niclosamide with FA‐conjugated dendrimer (FA‐2DG‐D‐Niclo) was examined using the mouse model of EuE described above. On Day 7, ELL‐induced mice were administered a single i.p. injection of control (FA‐2DG‐D), or 25 mg or 50 mg/kg body weight (bw) of FA‐2DG‐D‐Niclo. Behavioral tests were performed on Day −1 (a day before lesion induction), Day 7 (before dosing), and Day 14 or 21 (1 or 2 weeks after dosing, n = 16 at Day −1 and Day 7, n = 8 at Day 14 and Day 21). Mice were then euthanized on Day 14 or 21 after the last behavioral test. Peritoneal fluid was recovered by lavage (4 × 2 mL of ice‐cold PBS with 3% FBS) for immune profiling. The number and volume of lesions were recorded. Lesions were fixed in 4% paraformaldehyde for further analysis. A standard behavioral (mechanical sensitivity) test was performed before sample collection, as described by our laboratory previously [ 51 , 52 ]. Mice ( n = 8/group/time point) were allowed to acclimate quietly in the testing room for 30 min under red light. Then, von Frey filaments (BIO‐VF‐M, Bioseb) were applied 10 times to the skin perpendicular to the lower abdomen and bilateral hind paws to evaluate local or systemic sensitivity, respectively. The force in grams (g) of the filament evoking a withdrawal response (50% response count as sensitive) was recorded. Three behaviors were considered positive responses to filament stimulation: (1) sharp retraction of the abdomen, (2) immediate licking and/or scratching of the area of filament stimulation, or (3) jumping. All behavioral tests were performed with investigators blinded to treatment groups. These data were then subsequently analyzed by another blinded investigator. Single‐cell suspensions of peritoneal exudate cells were used for immune cell profiling by flow cytometry as described previously [ 51 , 52 ]. Briefly, peritoneal exudate cells were lysed using Red Blood Cell Lysis Buffer (BioLegend) and incubated at room temperature for 20 min with Zombie Aqua Fixable Viability dye (BioLegend). Cells were blocked on ice for 20 min with Fc Block anti‐CD16/CD32 (ThermoFisher) and stained with fluorochrome‐conjugated monoclonal antibodies for 1 h (Table S8 ). Samples ( n = 8/group) were acquired with the Attune NxT Acoustic Focusing Cytometer using Attune NxT software (ThermoFisher), and data were analyzed with FlowJo v10.4 software (FLOWJO). The peritoneal exudate cell suspension was first fixed with 4% paraformaldehyde at RT for 15 min, and blocked with 10% donkey serum at RT for 30 min. Cells were then incubated with FRβ primary antibody (1:200, Abcam, Cat.#ab302532) and a macrophage marker, F4/80 (1:200, Cell Signaling Technology, Cat. #19564) primary antibodies at 4°C overnight and Alexa Fluor 488‐ and 568‐conjugated F(ab’) secondary antibodies (Molecular Probe) at RT for 1 h. Following DAPI staining, the cells were resuspended in Fluoromount‐G (SouthernBiotech) and mounted on slides. The mounted slides were imaged using a Leica SP8‐X Confocal Microscope. Immunohistochemistry of FRβ and F4/80 on cross‐sections (5 µm) of paraffin‐embedded tissues, including lesion, kidney, and liver, was performed using the FRβ primary antibody (1:400) and F4/80 (1:400) primary antibodies and Alexa Fluor 488‐ and 568‐ conjugated F(ab’) secondary antibodies, or the VECTASTAIN ABC kit (Vector Lab). Immunostaining images were acquired by Leica SP8‐X Confocal Microscope or Leica DM4 B microscopy. FRβ‐positive cells in lesions were counted by ImageJ in an area of 0.07244 mm 2 , and the number of FRβ + cells per 1 mm 2 was shown. Statistical analyses were performed using GraphPad Prism (version 9.5). To compare differences in dendrimer organ uptake among groups, one‐way ANOVA followed by the Tukey multiple comparison test was used. To analyze behavioral test results, time‐dependent differences within a single group were compared using one‐way ANOVA followed by the Dunnett's multiple‐comparison test. To compare effects of different niclosamide dosages of niclosamide at each time point, one‐way ANOVA was used with Tukey multiple‐comparison tests. Differences in lesion number, volume, and FRβ + or CD45 + cell counts between groups at each time point were analyzed using one‐way ANOVA followed by the Tukey multiple‐comparison test. In all cases, data were tested for normal distribution using the Shapiro–Wilk normality test. If data were normally distributed, one‐way ANOVA was used to analyze differences among groups. If data were not normally distributed, the Kruskal–Wallis test was performed, as stated in the figure caption. Significance was defined as p < 0.05 (*), p < 0.01 (**), p < 0.001 (***), or p < 0.0001 (****).

We performed an integrated analysis with our newly generated pelvic exudate cell scRNA‐seq dataset from women with EuE (Tables S1 and S2 ), along with publicly available human datasets, including eutopic endometrium without or with EuE, EcE (lesion), and pelvic exudate cells (Figure  1A,B , Figure S1 , and Table S3 ). After data merging and quality control filtering, 748 509, and 76 187 total high‐quality endometrial and pelvic exudate cells were retained, respectively. Clustering analysis identified seven major cell types in the endometrium (EM), including B lymphocytes (CD79A + ), endothelial cells (PECAM1 + ), epithelial cells (EPCAM + ), myeloid cells (LYZ + ), perivascular cells (RGS5 + ), stromal cells (DCN + ), and T lymphocytes and natural killer (NK) cells (CD3E + and CD2 + ) (Figure  1C,E , Figure S1 , and Table S4 ). In the pelvic cavity (P), B lymphocytes (CD79A + ), dendritic cells (CD1C + ), macrophages (Mac, CD14 + ), mast cells (TPSAB1 + ), NK cells (GNLY + ), and T lymphocytes (CD3D + ) were assigned as overarching cell types (Figure  1D,F and Table S4 ). As expected, UMAP projection confirmed that FOLR2 + cells are localized within the myeloid cell clusters of endometrial and pelvic exudate cells (Figure  1G,H and Figure S1D,G ). In the pelvic macrophages, FRβ + macrophages are predominantly composed of tissue‐resident macrophages (cluster 0: TIM4 hi , cluster 1: APOE hi , and cluster 4: proliferating macrophages), while monocyte‐like (cluster 2: CCR2 hi monocyte‐converting macrophages) or newly differentiated macrophages (cluster 3: MARCO hi and cluster 6: CD3 hi ) show absent or low‐abundant FRβ expression, respectively (Figure S1B ). GO enrichment analysis indicated that both endometrial and pelvic FOLR2 + macrophages were primarily associated with myeloid migration and activation and immune regulation (Figure  1I,J and Table S6 ). In particular, biological processes associated with cytokine production, leukocyte cell–cell adhesion, and interleukin‐6 (IL‐6) production were predominantly enriched in the FOLR2 + macrophages of the endometrium. Stronger association with phagocytosis, cholesterol regulation, and response to bacteria was detected in the pelvic FOLR2 + macrophages, consistent with the primary roles of resident FOLR2 + macrophages in host defense and metabolic processes [ 53 , 54 , 55 ]. Comparative analysis showed a significantly higher abundance of FOLR2 + macrophages in EcE than in eutopic endometrium with (EuE) and without (Cont) EuE (Figure  1K ). In the ectopic and eutopic endometrium, FRβ expression is also shown in a specific population of CD14+ macrophages (Figure S1F ). Similarly, in pelvic exudate cells, significantly elevated numbers of FOLR2 + macrophages were observed in patients with EuE (Figure  1L ). Furthermore, DEG analysis of endometrial and pelvic FOLR2 + macrophages demonstrated significant upregulation of proinflammatory (CCL2, IL‐6R, and NFKB1), angiogenesis (FGFR1, HIF1A, and MMP2), and adhesion‐related genes (ITGAV, TGFBR1, and TGFBR2) in EuE‐associated endometrium (EuE and/or EcE) and pelvic cells (Endo) (Figure  1M,N and Table S7 ). Collectively, these results indicate an expansion of a pro‐disease FOLR2 + macrophage subpopulation in the EcE and pelvic cavity in EuE patients with pro‐disease functions. A major challenge in developing nanomaterial‐based drug delivery systems lies in the complexity and limited reproducibility of synthetic procedures, scalability constraints, and poor batch‐to‐batch consistency. To overcome these limitations, we used an efficient copper(I) catalyzed alkyne‐azide cycloaddition (CuAAC) click reaction to design and synthesize the FA‐2DG‐D conjugate capable of selectively targeting FRβ + macrophages for treating EuE. This click chemistry‐based approach offers a robust and versatile strategy for achieving precise, reproducible, and site‐specific conjugation of targeting ligands or therapeutic molecules to polymeric nanocarriers, including dendrimers [ 28 , 56 , 57 , 58 ]. FA was chemically conjugated to the dendrimer via a CuAAC click reaction (Figure  2A ). To enable this conjugation, FA ( 1 ) was functionalized with an azide group to enable its participation in the CuAAC reaction. The folate receptor recognizes FA through its pterin and p ‐aminobenzoate moieties via multiple hydrogen bonds and electrostatic interactions within the receptor pocket [ 59 ]. To preserve these recognition elements, conjugation with the azide linker was performed through the γ‐carboxylic acid group of the glutamic acid residue, forming a stable amide linkage with the terminal amine of the linker. The carboxylic acid in the ligand was activated using dicyclohexylcarbodiimide (DCC) and N ‐Hydroxysuccinimide (NHS), and the resulting NHS‐activated intermediate was reacted with azido‐PEG5‐amine ( 2 ) in the presence of DIPEA to yield the desired FA‐Azide ( 3 ). Successful formation of the azide‐functionalized FA was confirmed by 1 H NMR spectroscopy, which showed characteristic PEG proton signals from the azide linker at δ 3.0 (2H), 3.4 (2H), and 3.5–3.7 (20H) ppm, along with a new amide proton signal at δ 7.3 ppm (Figure  2B and Figure S2 ). HPLC analysis confirmed the purity of the product, exhibiting a single peak with a retention time of 13.3 min and more than 98% purity (Figure S4 ). To enable the subsequent CuAAC coupling, terminal alkyne groups were introduced on the surface of 2DG‐D. The primary hydroxyl groups of the surface‐exposed 2DG moieties were utilized for this modification. 2DG‐D ( 4 ) was reacted with 5‐hexynoic acid ( 5 ) in the presence of 1‐ethyl‐3‐(3‐dimethylaminopropyl)carbodiimide hydrochloride (EDC) and 4‐(dimethylamino)pyridine (DMAP) to afford 2DG‐D‐Hexyne ( 6 ) (Figure  2A ). 1 H NMR spectrum showed new methylene proton signals from the hexynoic linker in the aliphatic region at δ 1.75‐1.68 (8H), 2.46‐2.34, and 2.74‐2.58 (8H) ppm along with 52 distinct protons from the 2DG‐D around 8 ppm corresponding to 24 surface triazole protons in 2DG‐D (sharp signal), 8 interior triazole protons connecting the dendrons/arms to the core of the dendrimer and 20 amide protons (broad signal) from the core of the 2DG‐D dendrimer (Figures  2B and S6 ), confirming the attachment of 4 hexyne arms. HPLC analysis showed a shift in retention time from 14.0 min for 2DG‐D ( 4 ) to 19.7 min for 2DG‐D‐Hexyne ( 6 ), confirming successful surface modification (Figures S8 and S16 ). Finally, the CuAAC reaction between FA‐Azide ( 3 ) and 2DG‐D‐Hexyne ( 6 ) yielded the FRβ + macrophage‐targeted platform FA‐2DG‐D ( 7 ). Previous studies have demonstrated that dendrimers bearing a single FA ligand exhibit monovalent, reversible binding with folate‐binding protein (FBP), whereas dendrimers conjugated with two or more FA molecules engage in multivalent interactions, resulting in stronger receptor binding [ 60 , 61 , 62 ]. Given the high hydrophobicity of FA, conjugating more than two ligand molecules could potentially reduce the dendrimer's aqueous solubility. Our CuAAC click chemistry approach ensured desired loading of FA on the dendrimer. The number of attached molecules was confirmed through the comparative integration of aromatic protons from the dendrimer at δ7.2 ppm (16H), and the aromatic and amide protons of FA at δ 8.7 ppm (2H), 7.7 ppm (4H), 7.0 ppm (4H), and 6.7 ppm (4H). The HPLC chromatogram showed a retention time shift from 19.7 min for 2DG‐D‐Hexyne ( 6 ) to 16.1 min for FA‐2DG‐D ( 7 ) (Figures S9 and S11 ). The purity of FA‐2DG‐D (7) was determined to be more than 96% by HPLC (Figure S11 ). Dynamic light scattering (DLS) analysis revealed a hydrodynamic diameter of 6.6 ± 0.4 nm and a zeta potential of −5.8 ± 0.3 mV (Figures S12 and S13 ). Preparation and structural elucidation of FA‐2DG‐D‐Cy5 conjugate. (A) Schematic representation for the synthesis of FA‐2DG‐D‐Cy5 using a CuAAC reaction. Reagents and conditions: (i) DMSO, DCC, NHS, DIPEA, RT, 12 h, 70% (ii) DMF, EDC.HCl, DMAP, RT, 12 h, 80% (iii) CuSO 4 .5H 2 O, Sodium ascorbate, DMF, DI Water, 60°C, 10 h, 84% (iv) CuSO 4 .5H 2 O, sodium ascorbate, DMF, DI Water, 40°C, 6 h, 88%. (B) 1 H NMR spectra showing characteristic proton signals of 2DG‐D, 2DG‐D‐Hexyne, Folic Acid, FA‐Azide, FA‐2DG‐D and FA‐2DG‐D‐Cy5 at each step of the synthesis. To enable visualization and assessment of uptake of the dendrimer in FRβ + macrophages in EuE, the dendrimer was further functionalized with a near‐infrared dye, cyanine 5 (Cy5). The FA‐2DG‐D ( 7 ), containing approximately two terminal alkynes on its surface, was conjugated with Cy5‐azide ( 8 ) via a CuAAC reaction, yielding the fluorescently labeled FA‐2DG‐D‐Cy5 ( 9 ). Successful conjugation was confirmed by the appearance of characteristic Cy5 proton signals in the 1 H NMR spectrum (Figure  2B ). The number of Cy5 molecules attached to the dendrimer surface was estimated by proton integration. Comparison of the dendrimer aromatic protons at δ 7.2 ppm (16H) with the Cy5 aromatic protons at δ 8.4 ppm (4H), 8.2 ppm (4H), 7.4 ppm (4H), 6.6 ppm (2H), and 6.4 ppm (2H) indicated the attachment of approximately two Cy5 molecules (Figure  2B and Figure S14 ). The HPLC chromatogram of FA‐2DG‐D‐Cy5 ( 9 ) exhibited a retention time shift from 16.1 to 17.7 min following Cy5 conjugation, with a purity exceeding 97% (Figures S15 and S16 ). In addition, the Cy5‐labeled dendrimer displayed a characteristic UV absorbance at 650 nm, corresponding to Cy5, which further confirmed successful conjugation. A Cy5‐labeled 2DG‐D dendrimer (2DG‐D‐Cy5), lacking the FA targeting ligand, was prepared as a nontargeted control for uptake studies following our previously reported procedure [ 27 ]. We developed a niclosamide‐labeled dendrimer platform, FA‐2DG‐D‐Niclo, by conjugating niclosamide to the FA‐2DG‐D, to enable targeted delivery of niclosamide to disease‐driving FRβ + macrophages as a therapy for EuE. To achieve high conjugation efficiency and precise drug loading, CuAAC was employed. A niclosamide bearing an azide linker (Niclo‐Azide) was first synthesized by functionalizing the phenolic ─OH group of niclosamide (Figure  3A ). Several linker architectures were explored to identify a construct that combined physiological stability with controlled intracellular release. Initially, aliphatic and PEG‐based carboxylic acids were coupled to niclosamide through EDC/DMAP‐mediated esterification, aiming to generate ester linkages that are labile under intracellular conditions and stable at physiological pH. However, due to the proximity of the ester group to the aromatic ring, the resulting phenolic esters exhibited instability and underwent rapid cleavage during subsequent CuAAC reactions [ 63 ]. To circumvent this issue, a previous report on a disulfide‐linker‐based niclosamide prodrug was followed with some modifications to design and synthesize niclosamide‐azide (niclo‐azide) containing a disulfide linker [ 64 ]. It was reported that the disulfide linker remains stable under extracellular conditions but undergoes selective cleavage in the intracellular environment, thereby releasing free niclosamide [ 64 ]. Preparation and structural elucidation of FA‐2DG‐D‐Niclo conjugate. (A) Schematic representation for the synthesis of (i) Niclo‐Azide and (ii) FA‐2DG‐D‐Niclo using CuAAC reaction. Reagents and conditions: (i) DCM, HCl, Dioxane, RT, 3 h (ii) DCM, TEA, Triphosgene, −25°C, 1 h (iii) DCM, TEA, DMAP, RT, 16 h, 49% (iv) DMF, EDC.HCl, DMAP, RT, 12 h, 82% (v) CuSO 4 .5H 2 O, Sodium ascorbate, DMF, DI Water, 60°C, 10 h, 85% (vi) CuSO 4 .5H 2 O, Sodium ascorbate, DMF, DI Water, 40°C, 10 h, 75% (B) 1 H NMR spectra showing characteristic proton signals of FA‐2DG‐D’, Compound 10, Niclosamide, Niclo‐Azide and FA‐2DG‐D‐Niclo at each step of the synthesis. The synthesis of niclo‐azide was carried out through a multistep route (Figure  3A ). The Boc‐protected disulfide derivative ( 10 ) was synthesized using a published procedure [ 64 ]. It was then deprotected using 4 N HCl in dioxane to afford compound 11 as the amine hydrochloride salt, which was directly converted to the corresponding carbamoyl chloride intermediate 12 using triphosgene, following coupling with niclosamide ( 13 ) under mild basic conditions to afford the desired niclo‐azide ( 14 ) (Figure  3A ). Successful formation of the product was confirmed by comparative integration in 1 H NMR spectroscopy, which showed characteristic aromatic protons from the niclosamide scaffold at δ 8.4‐7.3 ppm (6H) and the aliphatic protons from the disulfide linker between δ 3.7‐2.7 ppm (11H) (Figure  3B and Figure  S19 ). The product purity (∼ 95.9%) was verified by HPLC, which showed a chromatogram with a retention time of 14.7 min (Figure  4A and Figure S21 ). To enable subsequent CuAAC coupling, 2DG‐D (4) was partially modified with 5‐hexynoic acid. Approximately 12 hydroxyl groups per dendrimer were modified to yield 2DG‐D‐Hexyne′ (15 ) (Figure  3A ). The formation of the required product was confirmed by the appearance of characteristic methylene proton signals from the hexynoic linker in the aliphatic region of the 1 H NMR spectrum at δ 1.7 (24H), 2.4 (24H), and 2.2 ppm (Figure  3B and Figure S23 ), which confirmed the presence of 12 hexyne arms. HPLC analysis of 2DG‐D‐Hexyne’ (15) showed the chromatogram at 10.4 min with a purity exceeding 97% (Figure  4A ). Subsequently, a CuAAC reaction between FA‐Azide (3) and 2DG‐D‐Hexyne′ (15) yielded FA‐2DG‐D′ (16) . The 1 H NMR spectrum of FA‐2DG‐D′ (16) showed characteristic aromatic resonances corresponding to FA, confirming successful conjugation of two molecules per dendrimer (Figure S26 ). The resulting construct retained approximately 10 free terminal alkyne groups per dendrimer. The HPLC chromatogram exhibited a retention time shift from 10.4 min for 2DG‐D‐Hexyne′ to 9.5 min for FA‐2DG‐D′ (16) , with a purity of ∼99% (Figure  4A and Figure S28 ). Lastly, FA‐2DG‐D’ (16) was conjugated with niclo‐azide ( 14 ) through CuAAC to obtain FA‐2DG‐D‐Niclo (17) . Consistent with our previous studies [ 56 , 58 ], we aimed to maintain the drug loading between 10% and 15% to ensure aqueous solubility of the dendrimer conjugate. Drug loading was calculated by 1 H NMR spectroscopy through comparison of the dendrimer aromatic protons at δ 7.2 ppm (16H) with the newly aromatic signals at δ 8.4 (10H), 8.3 (10H), and 8.2‐7.8 ppm, along with distinct amide protons of niclosamide at δ 10.43 ppm (10H), confirmed the conjugation of approximately 10 niclosamide molecules per dendrimer, corresponding to ∼12% drug loading by weight (Figure  3B and Figure S29 ). HPLC analysis revealed a shift in retention time from 9.5 min for FA‐2DG‐D′ (17) to 11.2 min for FA‐2DG‐D‐Niclo (17) , with a final purity of more than 98% (Figure S31 ). DLS analysis indicated a hydrodynamic diameter of 7.6 ± 0.6 nm and a zeta potential of −7.7 ± 0.6 mV (Figure  4C and Figures S32 and S33 ). Characterization of FA‐2DG‐D‐Niclo conjugate and intermediates: (A) HPLC chromatograms of intermediates at various synthesis steps and percentage purity of FA‐2DG‐D‐Niclo. (B) (i) Hydrodynamic diameter, and (ii) zeta potential of FA‐2DG‐D analyzed by dynamic light scattering (DLS) in triplicate. (C) (i) Hydrodynamic diameter and (ii) zeta potential of FA‐2DG‐D‐Niclo analyzed by DLS in triplicate. The physicochemical properties of FA‐2DG‐D‐Niclo (17) are presented in Figure  5A . While niclosamide is practically insoluble in water (∼5 µg/mL) [ 65 ], the dendrimer–drug conjugate exhibited excellent aqueous solubility (∼150 mg/mL). At a drug loading of 12%, this corresponds to an effective niclosamide solubility of approximately 18 mg/mL in water, representing a ∼3600 fold increase in solubility compared to free niclosamide (Figure  5A ). Characterization of FA‐2DG‐D‐Niclo conjugate and intermediates: (A) Table showing the physicochemical properties of FA‐2DG‐D‐Niclo and image showing the increase in solubility of niclosamide after conjugation with FA‐2DG‐D scaffold (FA‐2DG‐D‐Niclo). (B) The figure shows the reproducibility in the synthesis of multiple 400 mg‐scale batches of FA‐2DG‐D‐Niclo via 1 H NMR and HPLC. The 1 H NMR spectra of three batches (400 mg scale) show characteristic protons from FA‐2DG‐D‐Niclo. The HPLC chromatographs show reproducibility in the purity of these batches, with all these batches showing >99% purity. (C) In vitro drug release profile of FA‐2DG‐D‐Niclo conjugate in PBS and 0.1 mM GSH, intracellular conditions at 37°C after 15 days, and in (100%) human plasma after 120 h. The shelf stability of FA‐2DG‐D‐Niclo (17) (1 mg/mL) was assessed in PBS at 4°C, room temperature (25°C), and 40°C over a 30 day period. The formulation remained stable throughout the study, retaining more than 96% purity by HPLC at all tested temperatures (Figure S34 ). No shifts in retention time were observed over 30 days, confirming the chemical stability of the formulation under these conditions. To investigate the cellular uptake of the drug‐loaded dendrimer, a fluorescently labeled FA‐2DG‐D‐Cy5‐Niclo ( 18 ) was synthesized by reacting FA‐2DG‐D′ ( 16 ) first with Cy5‐azide (8) , followed by niclo‐azide ( 14 ) via a CuAAC reaction. Successful conjugation was confirmed by the appearance of characteristic Cy5 and niclo proton signals in the 1 H NMR spectrum (Figure S35 ). HPLC analysis of FA‐2DG‐D‐Cy5‐Niclo (18) showed a purity greater than 99% (Figure S36 ). The Cy5‐labeled dendrimer exhibited a characteristic UV absorbance at 650 nm, providing additional confirmation of the successful conjugation. Ensuring synthetic reproducibility and scalability remains a central barrier in advancing nanoparticle‐based therapeutics toward clinical applications. To overcome these challenges, we established a streamlined and scalable synthetic workflow for FA‐2DG‐D‐Niclo (17) that enables rigorous characterization of intermediates at each stage. To assess batch‐to‐batch consistency, three independent 400 mg‐scale batches were synthesized and evaluated by 1 H NMR and HPLC. The 1 H NMR spectra of all three batches (Figure  5B , left) showed identical chemical profiles and consistent drug and ligand loading. Similarly, HPLC analysis revealed overlapping chromatogram peaks with retention times of ∼11.2 min and purities exceeding 98%, confirming robust batch‐to‐batch reproducibility (Figure  5B , right). We further assessed the in vitro drug‐release profile of the conjugate under conditions mimicking both the extracellular environment (PBS + 0.1 mM GSH and 100% human plasma) and the intracellular microenvironment (pH 5.5 citrate buffer + 10 mM GSH) (Figure  5C ). Since niclosamide is attached to the dendrimer through a disulfide linkage, it enables glutathione (GSH) responsive cleavage [ 66 ]. Upon cleavage, the generated free thiol is known to undergo an intramolecular nucleophilic attack on the tertiary carbamate, ultimately yielding free niclosamide [ 64 ]. When conjugated to the targeted FA‐2DG‐D, this design facilitates controlled release within the intracellular environments while limiting premature systemic exposure outside the lesion sites. In PBS and 0.1 mM GSH, the conjugate was found to be relatively stable. Less than 40% of the drug was observed to be released in around 2 weeks, with <10% drug release detected for the first 5 days. Notably, less than 5% of the drug was released within 8 h. When incubated in human plasma, less than 35% of the drug was released over 5 days, indicating stability in physiological conditions (Figure  5C ). Under intracellular conditions (pH 5.5 citrate buffer + 10 mM GSH), the conjugate showed a sustained release profile, with ∼45% released in the first 24 h, ∼80% by two days, and reaching ∼95% over 12 days (Figure  5C ). The targeted and sustained intracellular drug release profile is important to maintain the efficacy of the drug for a long time and to avoid off‐target effects. We first evaluated the selective FRβ + macrophage targeting capability of FA‐2DG‐D‐Cy5 in ELL‐induced mice at 1, 6, 24, and 72 h ( n = 3) and compared it with sham controls ( n = 3). To evaluate the contribution of FA toward lesion targeting, biodistribution studies of nontargeted 2DG‐D‐Cy5 in ELL‐induced mice ( n = 3) were also performed under identical conditions (Figure  6 ). One week after lesion induction, 50 mg/kg bw of fluorescently labeled (FA‐2DG‐D‐Cy5) or nontargeted (2DG‐D‐Cy5) was injected into the tail vein. Cy5 fluorescent signals were calculated via the IVIS system at 1, 6, 24, and 72 h. FRβ + macrophages in the lesions showed a pronounced elevation in fluorescent (Cy5) signal starting at 1 h post‐injection (green arrows), which persisted above that of the 2DG‐D‐Cy5 group over the 72 h observation period (Figure  6A ). The ex vivo fluorescent results at 72 h showed specific uptake of FA‐2DG‐D‐Cy5 in the lesion, although some nonspecific uptake of FA‐2DG‐D‐Cy5 and 2DG‐D‐Cy5 was observed primarily in the liver and kidneys (Figure  6B,C ). For more precise quantification, frozen tissue samples from the 1, 6, 24, and 72 h groups were homogenized, and fluorescence intensity was measured to assess quantitative dendrimer uptake in lesions and major organs. Both FA‐2DG‐D‐Cy5 and 2DG‐D‐Cy5 showed strong, persistent accumulation within EuE lesions beginning at 1 h post‐injection and sustained through 72 h (Figure  6D and Figure S37 ). By 72 h, lesion uptake of FA‐2DG‐D‐Cy5 was significantly higher than that of 2DG‐D‐Cy5, suggesting its targeted accumulation in FRβ + macrophages that are abundant in endometriotic lesions (Figure  6D,E and Figure S37 ). Moreover, FA‐2DG‐D‐Cy5 demonstrated minimal accumulation (< 5% injected dose (% I.D.)) in other major organs, indicating rapid clearance from off‐target tissues (Figure  6E ). Importantly, unlike many nanoparticle systems that exhibit up to ∼80% hepatic accumulation, FA‐2DG‐D‐Cy5 showed only ∼4% liver accumulation, comparable to the previously characterized 2DG‐D‐Cy5 platform, demonstrating its favorable pharmacokinetic profile [ 27 ]. In vivo and ex vivo fluorescence imaging and whole‐body biodistribution of FA‐2DG‐D‐Cy5. (A) Whole‐body in vivo fluorescence images of mice following intravenous (tail vein) administration of FA‐2DG‐D‐Cy5 in sham controls (left) and ELL‐induced mice (right), or 2DG‐D‐Cy5 administered to ELL‐induced mice (center) at 1, 6, 24, and 72 h ( n = 3/group/timepoint). (B) Ex vivo fluorescence images of major organs collected at 72 h post‐injection. Organs are arranged from top to bottom as: brain, heart, spleen, lungs, liver, kidneys, and lesion. (C) Quantitative analysis of in vivo fluorescence signals in major organs at 72 h ( n = 3). (D) Heat map showing the mean percentage of injected dose (I.D.) in lesions and major organs at 1, 6, 24, and 72 h after FA‐2DG‐D‐Cy5 administration in sham controls and ELL‐induced mice, or ELL‐induced mice treated with 2DG‐D‐Cy5 ( n = 3/group). A heatmap was generated with the ComplexHeatmap package of R. (E) Quantitative biodistribution of FA‐2DG‐D‐Cy5 and 2DG‐D‐Cy5 in lesions and major organs at 72 h in sham and ELL‐induced mice ( n = 3). Data for (D) and (E) were obtained by fluorescence spectroscopy of homogenized tissues and are shown as the percentage of the injected dose per organ. Differences in each tissue among groups for data in (C) and (E) were analyzed by one‐way ANOVA followed by the Tukey multiple‐comparison test. p < 0.05 (*), p < 0.01 (**). ELL, endometriosis‐like lesion. We next examined the targeting ability, including selective cellular uptake, localization, and retention of FA‐2DG‐D‐Cy5‐Niclo, FA‐2DG‐D‐Cy5, or 2DG‐D‐Cy5 in ELL‐induced mice 2 weeks after dendrimer administration (Figure  7 ). Confocal imaging showed that FA‐2DG‐D‐Cy5‐Niclo and FA‐2DGD‐Cy5 (cyan) were selectively targeted and taken up by FRβ + (green) F4/80+ (red, a marker of macrophages) macrophages (not only in lesions but also in peritoneal exudate cells (Figure  7A,B , white arrows), where FRβ + macrophages were elevated in ELL‐induced mice. In contrast, cellular uptake of 2DG‐D‐Cy5 without FA ligand was limited in the lesion and peritoneal cells (Figure  7A,B ). Given the potential uptake of nontargeted 2DG‐D‐Cy5 by activated macrophages [ 28 ], we further quantified nonspecific FA‐2DG‐D‐Cy5 internalization in peritoneal exudate cells. The uptake was an average of 0.59%, indicating minimal nonspecific internalization. No dendrimer retention was observed in the kidneys, liver, or spleen in ELL‐induced mice 2 weeks following dendrimer administration (Figure S38 ), indicating that FA‐2DG‐D‐Cy5‐Niclo and FA‐2DG‐D‐Cy5 underwent complete clearance, and off‐target accumulation in major organs was negligible. These results suggest that FA‐2DGD‐Cy5 dendrimer is internalized only by ligand–receptor endocytosis in FRβ + macrophages in the lesions and peritoneal cells. As it remains stable within cells for at least 2 weeks, it functions as an excellent nanocarrier for drug delivery. Cellular uptake of 2DG‐D‐Cy5, FA‐2DG‐D‐Cy5, and FA‐2DG‐D‐Cy5‐Niclo at 2 weeks following intraperitoneal administration. Representative images showing selective uptake of 2DG‐D‐Cy5, FA‐2DG‐D‐Cy5, or FA‐2DG‐D‐Cy5‐Niclo (A) in peritoneal cells and (B) in endometriosis‐like lesions (ELL) from ELL‐induced mice ( n = 3). Peritoneal cells were costained with FRβ (green), F4/80 (red), and DAPI (blue). The Cy5 signal (Cyan) was directly acquired using a confocal microscope. White arrows denote Cy5 + signal in FRβ‐ and F4/80‐double positive cells. Lastly, we assessed the efficacy of FA‐2DG‐D‐Niclo in EuE disease progression and EuE‐associated hyperalgesia. After the lesion induction, a single dose of FA‐2DG‐D‐Niclo dendrimer (25 or 50 mg/kg bw of niclosamide) or FA‐2DG‐D without niclosamide control was administered i.p. in ELL‐induced mice at Day 7, as shown in Figure  8A . Then, the efficacy of dendrimers was examined 7 (Day 14) and 14 (Day 21) days after dendrimer administration. We first confirmed the suppressive capability of FRβ + macrophages in the lesions and peritoneal cavity by FA‐2DG‐D‐Niclo dendrimer (Figure  8B,C ). Single dosing of FA‐2DG‐D‐Niclo (50 mg/kg bw) significantly reduced FRβ + macrophages in the lesions at Day 21, although the reduction of FRβ + macrophages was not observed at Day 14 (Figure  8B ). The peritoneal FRβ + macrophage population was reduced by both doses (25 and 50 mg/kg bw) of FA‐2DG‐D‐Niclo (Figure  8C ). The results indicate that 7 days after dendrimer administration may not be sufficient to reduce FRβ + macrophages in both lesions and the peritoneum. Indeed, lesion number and volume were also significantly reduced at Day 21 by FA‐2DG‐D‐Niclo at both doses, but there were no significant differences in lesion number or volume at Day 14 (Figure  9A ). However, we realized that three mice treated with both doses and examined at Day 14 did not have lesions. Furthermore, four and five mice, at doses of 25 and 50 mg/kg bw of niclosamide, respectively, did not have lesions by Day 21, either. Thus, a single dose of FA‐2DG‐D‐Niclo administration completely inhibited lesion growth and progression in 37.5%–62.5% of mice, whereas all control mice developed lesions on Days 14 and 21. While reduced FRβ + macrophages in the remaining lesions were observed only at Day 21, a few mice at Day 14 lacked lesions for examination, suggesting that FA‐2DG‐D‐Niclo was likely effective for lesion growth 7 days after administration. In support of this implication, abdominal and hindpaw hypersensitivities were already improved at Day 14 (Figure  9B ). The abdominal and hindpaw sensitivity 7 days after lesion induction was high compared to before lesion induction (Day −1). At Day 14, FA‐2DG‐D‐Niclo at a dose of 50 mg/kg bw of niclosamide significantly reduced the abdominal sensitivity, which is considered indicative of peripheral visceral pain. Both doses of FA‐2DG‐D‐Niclo improved hindpaw sensitivity, which is likely to be affected by peripheral and central sensitization. By Day 21, FA‐2DG‐D‐Niclo at both doses further improved abdominal and hindpaw sensitivities. The results clearly showed that a single dose of FA‐2DG‐D‐Niclo inhibited disease progression and EuE‐associated hyperalgesia. Notably, a single dose of FA‐2DG‐D‐Niclo remained effective for 2 weeks after administration. The current FDA‐approved niclosamide treatment requires a high dose (2 g/day) due to its low water solubility and is limited to oral administration for 7 days. Niclosamide treatment for patients who have EuE would need to be longer due to the chronic nature of the disease. Long‐term systemic treatment with niclosamide could cause off‐target actions. Therefore, the 2DG‐D‐based delivery of niclosamide and targeting of disease‐specific FRβ + macrophages is highly translational, as FA‐2DG‐D can deliver niclosamide intracellularly into FRβ + macrophages, thereby enhancing efficacy, reducing side effects, and treating chronic diseases such as EuE. Quantitative analysis of FRβ + cells in the lesion and peritoneum following dendrimer treatments. (A) Study design as described in Methods. (B) Representative immunohistochemical images (left) and quantification of FRβ + cells (right) in lesions at 14 or 21 days after lesion induction ( n = 3–8, as dendrimer‐Niclo inhibited lesion development). One‐way ANOVA followed by the Tukey multiple‐comparison test was used to assess differences among groups. (C) Representative flow plots (left) and quantification of peritoneal FRβ + cells (right). Differences in peritoneal FRβ + cells among groups at each time point were assessed by the Kruskal–Wallis test. Data in all figures are shown as the mean ± SEM ( n = 8). p < 0.05 (*), or p < 0.01 (**). Efficacy of FA‐2DG‐D‐Niclo in treating endometriosis. (A) Quantification of lesion numbers and volumes in mice at Days 14 and 21 after lesion induction. Lesion volumes were calculated as length × width 2 . Differences in lesion number and volume among groups at each time point were assessed by the Kruskal–Wallis test. (B) Evaluation of endometriosis‐associated hyperalgesia. Abdominal (left) and hindpaw (right) withdrawal thresholds were assessed by the von Frey test. Data represent the group mean threshold at the corresponding time point ( n = 16 at Day −1 and Day 7, n = 8 at Day 14 and Day 21). Red arrows show the time of i.p. administration of FA‐2DG‐D or FA‐2DG‐D‐Niclo. The time course differences within each treatment group were compared using a Kruskal–Wallis test followed by the Dunnett multiple‐comparison test. The colored stars or ns (nonsignificant) represent the differences at the corresponding time point relative to Day −1 within the treatment group. To compare dosing effects of niclosamide at each time point among groups, the Kruskal–Wallis test was used. Data are shown as mean ± SEM ( n = 8). p < 0.05 (*), p < 0.01 (**), p < 0.001 (***), or p < 0.0001 (****).

With integrated single‐cell transcriptomic analysis, we further identified a significant recruitment of pro‐disease FRβ + macrophages within human endometriotic lesions and the pelvic cavity, mirroring our observations in the mouse model. Importantly, overexpression of FRβ on these macrophages provides a ligand‐guided entry point, enabling receptor‐specific targeting for both imaging and therapeutic interventions. Leveraging this receptor overexpression, we engineered a FA‐conjugated dendrimer (FA‐2DG‐D) to exploit FRβ‐mediated uptake for targeted imaging and drug delivery in endometriotic lesions. FA‐2DG‐D offers several critical advantages for translational development. The dendrimer architecture enables precise, reproducible conjugation of targeting ligands and therapeutic payloads via click chemistry, resulting in well‐defined nanostructures, excellent aqueous solubility, and robust batch‐to‐batch consistency. Conjugation of niclosamide to generate FA‐2DG‐D‐Niclo markedly enhanced the solubility of this poorly water‐soluble drug, enabled controlled, intracellularly triggered release via a disulfide linker, and minimized premature systemic exposure. Importantly, the formulation demonstrated high stability under physiological and storage conditions. In a mouse model of EuE, the Cy5‐labeled platform demonstrated lesion‐ and peritoneal cell‐specific accumulation and intracellular internalization, along with a favorable biodistribution with minimal off‐target organ accumulation. FA‐2DG‐D‐Niclo enabled targeted delivery to disease‐driving macrophages. Administration of FA‐2DG‐D‐Niclo led to significant reductions in lesion number and volume, and improved hyperalgesia, highlighting both therapeutic efficacy and immunomodulatory activity. This work demonstrates the potential of the fa‐2DG‐D platform to target FRβ + macrophages for the treatment of EuE. By addressing key limitations of current hormonal and systemic treatments, this approach establishes a non‐hormonal, FRβ + macrophage‐focused strategy for the treatment of EuE.

Endometriosis (EuE) affects an estimated 10% of reproductive‐aged women, roughly 190 million individuals worldwide, and remains one of the most debilitating gynecological disorders [ 1 ]. The disease is characterized by chronic pelvic pain, dysmenorrhea, dyspareunia, and infertility, all of which severely diminish the quality of life for patients and their families [ 2 , 3 , 4 ]. In the United States alone, the annual economic burden is estimated at $78–119 billion, encompassing healthcare costs, productivity losses, and reduced workforce participation [ 5 ]. Despite its prevalence and impact, EuE remains incurable, and the therapeutic landscape has changed little over the past several decades. Current medical management relies on hormonal suppression of estrogen production or action [ 6 , 7 ], such as oral contraceptives, progestins, and/or gonadotropin‐releasing hormone (GnRH) agonists/antagonists [ 6 , 8 ], as well as surgical excision of lesions. However, hormonal treatments are limited in efficacy, have high recurrence rates, and cause significant side effects. Surgical excision offers temporary relief of symptoms, yet nearly half of surgically treated women experience recurrence within five years, and ∼12% fail to achieve meaningful pain reduction after surgery [ 9 , 10 ]. Thus, there is an urgent need to identify new therapeutic targets to develop novel nonhormonal treatments in EuE. A growing body of evidence implicates immune dysfunction, particularly macrophage dysregulation, in driving lesion establishment, chronic inflammation, angiogenesis, and neurogenesis in EuE [ 11 , 12 , 13 ]. Recently, our single‐cell transcriptomic analysis identified a unique FRβ‐expressing macrophage population, differentiated from monocyte‐derived proinflammatory macrophages and characterized by resident macrophage traits in mouse EuE [ 14 ]. This population of macrophages is also identified in the lesions of EuE patients [ 15 ]. The selective upregulation of FRβ + on these macrophages can be used as an accessible and disease‐restricted cellular target for ligand‐guided targeted drug delivery. Our prior studies demonstrated that niclosamide significantly suppressed endometriotic lesion growth, alleviated aberrant peritoneal inflammation, and decreased vascularization and innervation in lesions using two distinct mouse models of EuE without compromising fertility [ 13 , 14 , 16 ]. Niclosamide (5‐chloro‐ N ‐(2‐chloro‐4‐nitrophenyl)‐2‐hydroxybenzamide) is a chlorinated salicylanilide derivative that is FDA‐approved as an anthelmintic drug. Currently, multiple clinical trials are ongoing to repurpose it for the treatment of other diseases, including cancer and metabolic diseases [ 17 ]. However, the clinical use of niclosamide is significantly limited by its poor aqueous solubility and correspondingly low oral bioavailability. To address these challenges, researchers are actively developing innovative formulation strategies such as water‐soluble derivatives, solid dispersions, and nanocarrier‐based delivery systems to enhance its solubility, pharmacokinetics, and overall therapeutic potential [ 18 ]. Additionally, systemic niclosamide treatment inhibits several pathways, including STAT3, NFκB, and WNT signaling, across all cell types [ 11 , 12 , 16 ], including other macrophages [ 14 ]. Disrupting these essential pathways across all cell types would lead to unwanted side effects, and this poses a significant barrier to therapeutic use of niclosamide. Importantly, niclosamide can also reduce elevated FRβ + macrophages and restore the perturbed microenvironment in EuE by finely regulating macrophage dynamics [ 14 ], suggesting that FRβ + macrophages can serve as a viable therapeutic target in EuE. Therefore, targeted delivery of niclosamide to FRβ + macrophages offers a strategy to maximize therapeutic efficacy while minimizing systemic toxicity. In this context, dendrimer nanotechnology has emerged as a powerful tool for targeted therapeutic delivery, enabling selective engagement of pathogenic cells while minimizing off‐target toxicity [ 19 , 20 , 21 , 22 , 23 , 24 ]. Dendrimers are monodisperse, highly branched macromolecules whose multivalent surface architecture and tunable chemistry allow precise conjugation of targeting ligands, therapeutic agents, and imaging probes [ 25 , 26 , 27 ]. Building on our previously synthesized 2‐deoxy‐glucose (2DG) functionalized dendrimer (2DG‐D) [ 28 ], which incorporates 2DG moieties to enhance solubility and macrophage interaction, we engineered a folic acid (FA), a ligand of FRβ, conjugated dendrimer (FA‐2DG‐D) by covalently coupling FA to the dendrimer surface through orthogonal click chemistry. This design exploits ligand‐FRβ binding to selectively direct dendrimer uptake into FRβ + macrophages within endometriotic lesions and peritoneum [ 29 , 30 ]. To achieve targeted delivery, we further developed the niclosamide‐conjugated FA‐2DG‐D (FA‐2DG‐D‐Niclo). Here, we report the design, synthesis, and in vivo evaluation of FA‐2DG‐D and FA‐2DG‐D‐Niclo in a mouse model of EuE. Our results demonstrate that FA‐2DG‐D has FRβ + macrophage‐specific targeting in the lesions and peritoneum and delivers niclosamide into FRβ + macrophages with ligand–receptor internalization. This targeted dendrimer platform offers a promising strategy for therapeutic delivery in EuE, addressing a critical unmet need for more precise and effective interventions.

The authors A.S, K.H, R.S, A.D, M.S, M.E.H, and A.I.D have pending patents and invention disclosures on 2DG and/or FA‐2DG dendrimers and their conjugates.

The chemistry experimental details and supplementary figures related to characterization, stability studies, and biodistribution are provided in the Supporting Information . Supporting File 1 : adhm71661‐sup‐0001‐SuppMat.docx. Supporting File 2 : adhm71661‐sup‐0002‐SuppMat.xlsx.