This study was approved by the Institutional Review Committee of the Affiliated Cancer Hospital of Hainan Medical University (approval no. 2024.39). A total of 157 patients with pathologically confirmed primary lung adenocarcinoma (LUAD) who underwent surgical resection at Hainan Cancer Hospital between January 2016 and December 2024 were enrolled. The cohort comprised 69 male and 88 female patients (male-to-female ratio, 1:1.28), with a mean age of 63.1±8.5 years (range, 37–80 years). Paired tumor tissues and adjacent non-tumorous lung tissues (obtained ≥5 cm from the tumor margin) were collected at the time of surgery. Clinical and pathological data, including age at diagnosis, TNM stage (16), pathological subtype and histological grade, were retrieved from medical records. Written informed consent was obtained from all participants prior to enrollment.

TMAs were constructed using paired tumor and adjacent non-tumorous tissues. Specimens were fixed in 10% neutral-buffered formalin at room temperature for 24 h, dehydrated through a graded ethanol series (50–100%; 1 h each), cleared in xylene (three changes, 30 min each), infiltrated with paraffin (three changes, 1 h each) and embedded in paraffin blocks (melting point, 58°C). Cylindrical cores (2 mm in diameter) were extracted from representative regions of each donor block and transferred to recipient paraffin blocks using a tissue arrayer. Sections (4 µm thick) were cut from the TMA blocks, baked at 72°C for 30 min and deparaffinized by sequential immersion in xylene (three changes) and a descending ethanol series (100, 90, 80, 70 and 50%), 5 min each. Antigen retrieval was performed in EDTA buffer (pH 9.0; cat. no. K8004; Quantum Analytics, Inc.) using an automated IHC pretreatment device at 97°C for 18 min, followed by gradual cooling to 65°C. Endogenous peroxidase activity was quenched with 3% hydrogen peroxide for 10 min at room temperature. The sections were then incubated with an anti-HDAC1 primary antibody (dilution, 1:100; cat. no. 34589; Cell Signaling Technology, Inc.) at room temperature for 1 h. A two-step horseradish peroxidase-conjugated detection system (cat. no. PV-9000; ZSGB-Bio, Inc.) was subsequently applied. The sections were incubated with reaction enhancement solution (Reagent 2) at room temperature for 20 min, followed by enhanced enzyme-labeled secondary antibody (Reagent 3) at 37°C for 20 min. Immunoreactivity was visualized using 3,3′-diaminobenzidine (dilution, 1:20; cat. no. ZLI-9018; ZSGB-Bio, Inc.), and the sections were counterstained with hematoxylin at room temperature for 10–15 sec. After dehydration, clearing and mounting, images were captured under a light microscope (magnification, ×40 and ×400; Olympus BX53; Olympus Corporation). All slides were evaluated independently by two pathologists blinded to the clinical data.

HDAC1 expression was scored semi-quantitatively based on the staining intensity (0, negative; 1, weak; 2, moderate; 3, strong) and the proportion of positively stained cells (0, 0%; 1, <10%; 2, 10–50%; 3, 51–80%; 4, >80%). The final immunoreactive score (IRS) was calculated by multiplying the staining intensity score by the proportion score, yielding a range of 0–12. The cases were dichotomized into low-(IRS, 0–4) and high-expression (IRS, 6–12) group.

Human NSCLC cell lines A549, H1299 and H1975 were obtained from the Cell Bank of the Type Culture Collection of the Chinese Academy of Sciences. Cells were maintained in Roswell Park Memorial Institute-1640 (RPMI-1640) medium (cat. no. C11875500BT; Gibco; Thermo Fisher Scientific, Inc.) supplemented with 10% fetal bovine serum (FBS; cat. no. ST30-3302; PAN-Biotech GmbH) and 1% penicillin-streptomycin (cat. no. 15140-122; Gibco; Thermo Fisher Scientific, Inc.) at 37°C in a humidified 5% CO₂ incubator.

HDAC1-knockdown (HDAC1 K/D), HDAC1-overexpressing (HDAC1 O/E) and β-catenin-overexpressing (β-catenin O/E) stable cell lines were generated in A549, H1299 and H1975 cells using lentiviral transduction. For HDAC1 K/D, validated HDAC1-targeting short hairpin RNA (shRNA) sequences were cloned into the pLKO.1-GFP-puro lentiviral vector (Generay Biotech Co., Ltd.), with the empty pLKO.1 vector serving as the negative control. For HDAC1 O/E, a full-length human HDAC1 coding sequence (GenBank Accession No. NM_004964) was amplified and cloned into the pCDH-GFP-puro lentiviral vector (Generay Biotech Co., Ltd.), with the empty pCDH vector serving as the corresponding control. For β-catenin O/E, a full-length human CTNNB1 coding sequence (GenBank Accession No. NM_001904) was cloned into the same vector to generate pCDH-β-catenin, again with the empty pCDH vector as the control. The identity of each insert was verified by Sanger sequencing prior to virus production. Recombinant lentiviruses were produced using a second-generation packaging system. Briefly, 1 µg pMD2.G and 3 µg psPAX2 were cotransfected with 4 µg pCDH-HDAC1, pLKO.1-HDAC1 or pCDH-β-catenin, or with 4 µg of the corresponding empty vector (pCDH or pLKO.1), into 293T cells (cat. no. NCL-10003; NCM Biotech) using Nano293T Transfection Reagent (cat. no. C500T-1; NCM Biotech) at 37°C for 12 h according to the manufacturer's instructions. The culture medium was replaced 12 h following transfection and virus-containing supernatants were harvested at 24, 48 and 72 h post-transfection. The supernatants were filtered through 0.45-µm polyvinylidene fluoride membranes to remove cell debris, pooled and stored at −80°C until use. Target cells were seeded in six-well plates and transduced with the harvested lentivirus in the presence of 8 µg/ml polybrene (MilliporeSigma). The multiplicity of infection in A549, H1299 and H1975 cells was 20, 40 and 20 for pCDH-HDAC1; 40, 40 and 20 for pLKO.1-HDAC1; and 20, 40 and 10 for pCDH-β-catenin. After 24 h, the medium was replaced with fresh complete medium. Transduced cells were selected with 6 µg/ml puromycin (Beijing Solarbio Science & Technology Co., Ltd.) over 3 weeks, with the medium refreshed every 2–3 days. The resulting stable lines were designated A549-HDAC1 K/D, H1299-HDAC1 K/D and H1975-HDAC1 K/D for the knockdown groups, and A549-HDAC1 O/E, H1299-HDAC1 O/E and H1975-HDAC1 O/E for the overexpression groups, alongside their respective empty vector controls. All subsequent experiments were performed using the stable cell lines obtained after this 3-week selection period. Knockdown and overexpression efficiencies were validated by western blotting. All shRNA, overexpression and control sequences used in the present study are listed in Table SI.

To investigate whether the oncogenic effects of HDAC1 in NSCLC are mediated through β-catenin signaling, rescue experiments were performed by overexpressing β-catenin in HDAC1 K/D stable cell lines. Lentivirus production, transduction and puromycin selection were performed as described above. The resulting double-modified stable lines were designated A549-HDAC1 K/D + β-catenin O/E, H1299-HDAC1 K/D + β-catenin O/E and H1975-HDAC1 K/D + β-catenin O/E. In parallel, β-catenin was overexpressed in parental cells to generate β-catenin O/E-alone groups (A549-β-catenin O/E, H1299-β-catenin O/E and H1975-β-catenin O/E), which served as the reference groups for the rescue experiments. β-catenin overexpression efficiency was confirmed by western blotting.

Cell proliferation was measured using Cell Counting Kit-8 (CCK-8; Vazyme Biotech Co., Ltd.) and colony formation assays. For the CCK-8 assay, cells (1.5×10³ cells/well) were seeded into 96-well plates in 200 µl culture medium. After incubation for 0, 24, 48, 72 and 96 h at 37°C with 5% CO₂, 10 µl of CCK-8 reagent was added to each well. The absorbance was then measured at 450 nm. Cell proliferation was calculated as the optical density (OD) of the experimental well/OD of control well (Day 1). For the colony assay, cells (1×10⁴ cells/well) were plated in 6-well plates in 2 ml complete medium and cultured for 2 weeks. The cells were fixed with 4% paraformaldehyde (MilliporeSigma) at room temperature for 30 min and stained with 0.05% crystal violet (MilliporeSigma) at room temperature for 30 min. Subsequently, colonies >1 mm in diameter were manually counted. Colony formation efficiency was calculated as the number of colonies divided by the number of cells seeded initially and the results were normalized to the control conditions. Each condition in both assays was tested in triplicate and each experiment was independently repeated at least three times.

Cell migration was assessed using a Transwell assay. Transwell inserts with 8-µm pore membranes (Corning Inc.) were placed in 24-well culture plates. Cells (5×104) suspended in 200 µl of serum-free culture medium were added to the upper chamber, whereas the lower chamber was filled with 600 µl of medium supplemented with 10% FBS as a chemoattractant. After 24 h of incubation at 37°C, the non-migrated cells in the chamber were removed with a cotton swab. The migrated cells were fixed with 20% methanol for 30 min and stained with 0.1% crystal violet for 30 min (both at room temperature). Migratory cells in four randomly selected fields per insert were counted under a light microscope (magnification, ×200; Olympus BX53; Olympus Corporation). Each experiment was performed in triplicate.

Cell invasion was evaluated using a Matrigel-coated Transwell chamber. Transwell inserts with 8-µm pores were pre-coated with Matrigel (3 mg/ml) and the gel was allowed to solidify at 37°C for 30 min. Then, 1×10⁵ cells suspended in 200 µl of serum-free medium were added to the upper chamber and 600 µl of medium containing 10% FBS was added to the lower chamber. After 24 h of incubation at 37°C, non-invading cells in the upper chamber were removed with a cotton swab. The invading cells were fixed with 4% paraformaldehyde for 30 min and stained with 0.1% crystal violet for 30 min (both at room temperature). Invading cells were counted under a light microscope (magnification, ×200; Olympus BX53; Olympus Corporation) across four randomly chosen fields per insert. Each experiment was performed in triplicate.

Nuclear and cytosolic proteins were extracted using a nucleocytoplasmic separation and extraction kit (Thermo Fisher Scientific Inc.) according to the manufacturer's instructions. The supernatants were collected and analyzed by western blotting. To verify the effectiveness of the fractionation, western blotting was performed using Lamin A as a nuclear marker and GAPDH as a cytoplasmic marker.

Cells were lysed in radioimmunoprecipitation assay buffer (Beyotime Biotechnology) containing protease inhibitors for 15 min. Lysates were clarified by centrifugation at 16,000 × g for 15 min at 4°C. Protein concentrations were quantified using the Bradford assay (Bio-Rad Laboratories, Inc.). Samples were mixed with 5X Laemmli sample buffer and boiled. Proteins (30–40 µg/well) were separated by 8% sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and then transferred onto nitrocellulose membranes (Whatman; Cytiva). The membranes were blocked with 5% non-fat milk in Tris-buffered saline containing 0.1% Tween-20 for 1 h at room temperature, and then incubated overnight at 4°C with primary antibodies against HDAC1 (dilution, 1:1,000), phosphorylated (p-)ERK1/2 (dilution, 1:2,000), total ERK1/2 (dilution, 1:2,000), p-AKT (T308) (dilution, 1:2,000), p-AKT (S473) (dilution, 1:2,000), total AKT (dilution, 1:2,000), c-Myc (dilution, 1:2,000), cyclin D1 (dilution, 1:2,000), E-cadherin (dilution, 1:2,000), vimentin (dilution, 1:2,000), β-catenin (dilution, 1:2,000), GAPDH (dilution, 1:3,000) and Lamin A (dilution, 1:1,000), all obtained from Cell Signaling Technology, Inc. The membranes were subsequently incubated with HRP-conjugated goat anti-rabbit IgG (cat. no. L3012; dilution, 1:5,000; SAB Biotherapeutics, Inc.) or HRP-conjugated goat anti-mouse IgG (cat. no. L3032; dilution, 1:5,000; SAB Biotherapeutics, Inc.) for 1 h at room temperature. Signals were detected using an enhanced chemiluminescence substrate (Thermo Fisher Scientific, Inc.) and images were captured using an Amersham Imager 600 (GE Healthcare). Band intensities were quantified using ImageJ software (version 1.8.0; National Institutes of Health).

All animal experiments were approved by the Institutional Animal Care and Use Committee of Hainan Medical University (approval no. 2024.331) and were performed at the Medical Research Center, Hainan Cancer Hospital. A total of 20 six-week-old female BALB/c nude mice (nu/nu; body weight, 15–20 g; SJA Laboratory Animal Center) were maintained under specific pathogen-free conditions at a temperature of 22±2°C and a relative humidity of 50±10%, with a 12-h light/dark cycle and ad libitum access to food and water. Prior to tumor cell inoculation, mice were allowed to acclimate for one week. A549 cells were selected as a representative lung adenocarcinoma model for the in vivo rescue experiments, as the in vitro effects of HDAC1 knockdown and β-catenin re-expression were consistent across all three cell lines (A549, H1299 and H1975). Mice were randomly divided into four groups and inoculated with stable A549 cells: empty-vector control, HDAC1 K/D, β-catenin O/E, or HDAC1 K/D + β-catenin O/E. For each mouse, 5×10⁵ cells were resuspended in 100 µl of a 1:1 mixture of PBS and Matrigel and injected subcutaneously into the right flank. Tumor diameter was measured every three days and tumor volume was calculated as (length × width²)/2. Mice were sacrificed when any individual tumor reached ~2,000 mm³ or when an animal lost 20% of its initial body weight, in accordance with humane endpoint guidelines. No mouse met these criteria during the study. At the experimental endpoint, four weeks after inoculation, all mice were sacrificed by carbon dioxide (CO₂) inhalation in accordance with institutional guidelines. CO₂ was introduced using a gradual-fill method at a rate displacing 30–70% of the chamber volume per minute. Tumor tissues were snap-frozen or lysed for western blotting analysis.

Data are expressed as mean ± standard deviation (SD) of at least three independent experiments. Associations between HDAC1 expression and clinicopathological characteristics were evaluated using the χ² test or Fisher's exact test, as appropriate. Comparisons between two groups were performed using a two-tailed Student's t-test (paired for matched tumor and adjacent tissue specimens; unpaired for independent cell-line experiments), whereas multigroup comparisons were analyzed using one-way analysis of variance followed by Tukey's post hoc test. All analyses were performed using SPSS for Windows (version 22.0; IBM Corp.). P<0.05 was considered to indicate a statistically significant difference.

To evaluate the expression of HDAC1 in LUAD, IHC staining was performed using tissue microarrays comprising 157 paired tumor and adjacent non-neoplastic lung tissues. As shown in Fig. 1A, HDAC1 immunoreactivity was weak or absent in adjacent non-neoplastic tissues, whereas LUAD tissues exhibited strong, diffuse nuclear staining. Consistent with these observations, the mean IRS was significantly higher in LUAD tissues compared with that in the matched adjacent tissues (P<0.001; Fig. 1B). When cases were dichotomized by IRS, high HDAC1 expression was detected in 143 of 157 (91.1%) LUAD specimens, compared with only 37 of 157 (23.6%) adjacent non-neoplastic tissues (χ²=146.273, P<0.001; Table I). These results demonstrated that HDAC1 is frequently overexpressed in LUAD.

The association between HDAC1 expression and the clinicopathological characteristics was further analyzed (Table II). High HDAC1 expression was significantly associated with lymph node metastasis (P=0.027) and the histological differentiation grade (P=0.006). Notably, all 46 patients with lymph node metastasis (100%) exhibited high HDAC1 expression compared with 97 of 111 (87.4%) patients without lymph node involvement. Regarding tumor differentiation, the proportion of high HDAC1 expression was greatest in poorly differentiated tumors (64/65, 98.5%), followed by well to moderately differentiated tumors (79/92, 85.9%), suggesting that HDAC1 upregulation is accompanied by a less differentiated and more aggressive phenotype. By contrast, no significant association was observed between HDAC1 expression and patient age, sex, maximum tumor diameter, T stage, vascular invasion, perineural invasion, necrotic components, or Ki-67 proliferation index (all P>0.05). Collectively, these findings indicate that elevated HDAC1 expression is closely linked to lymph node metastasis and poor tumor differentiation in LUAD.

To examine the functional role of HDAC1 in NSCLC, the present study established stable HDAC1-K/D cell lines using two independent shRNA constructs (K/D-1 and K/D-2) in A549, H1299 and H1975 cells. CCK-8 assay revealed that HDAC1 knockdown significantly reduced the proliferative capacity of all three cell lines over a 96-h observation period compared with the empty vector controls (P<0.001; Fig. 2A-C). Colony formation assays further confirmed these findings; HDAC1-K/D cells exhibited significantly reduced clonogenic ability compared with control cells (P<0.001; Fig. 2D and E). Next, it was examined whether HDAC1 depletion affects the migratory and invasive potential of NSCLC cells using Transwell assays. HDAC1 knockdown led to a significant reduction in both the migration (Fig. 2F and G) and invasiveness (Fig. 2H and I) of all three cell lines relative to their respective controls (P<0.001). Collectively, these data indicated that HDAC1 is required for the proliferative, migratory and invasive capacities of NSCLC cells.

Previous studies have demonstrated that HDAC1 expression is associated with activation of the PI3K/Akt, MAPK/ERK and Wnt/β-catenin signaling pathways in lung cancer (13–15). To determine which of these pathways mediated the observed phenotypes, the present study examined the expression of key signaling molecules by western blotting and nuclear-cytoplasmic fractionation (Fig. 2J-M). HDAC1 K/D significantly decreased c-Myc and cyclin D1 protein levels in all three cell lines (P<0.05), indicating impaired cell cycle progression following HDAC1 depletion. With respect to EMT markers, HDAC1 silencing led to a marked increase in the epithelial marker E-cadherin (P<0.01) and a concomitant decrease in the mesenchymal marker vimentin (P<0.01), indicating a reversal of the mesenchymal phenotype. Subcellular fractionation analysis further revealed that nuclear β-catenin levels were substantially reduced in HDAC1-K/D cells (P<0.001), whereas cytoplasmic β-catenin remained largely unchanged (P>0.05), suggesting that HDAC1 promoted the nuclear accumulation of β-catenin. By contrast, the phosphorylation of AKT (at both Thr308 and Ser473) and ERK as well as the total AKT and ERK protein levels showed no significant alterations following HDAC1 knockdown (P>0.05), indicating that the PI3K/AKT and MAPK/ERK pathways are not the primary downstream effectors of HDAC1 in this context. Thus, these results demonstrated that HDAC1 knockdown suppresses the proliferation, migration and invasion of NSCLC cells, accompanied by downregulation of c-Myc and cyclin D1, reversal of EMT and reduced nuclear accumulation of β-catenin.

The present study performed reciprocal gain-of-function experiments by stably overexpressing HDAC1 in A549, H1299 and H1975 cells. CCK-8 assays demonstrated that HDAC1 O/E significantly enhanced the proliferative capacity of all three cell lines over a 96-h period compared with the empty-vector controls (P<0.001; Fig. 3A-C). Consistent with these results, colony formation assays revealed that HDAC1-O/E cells exhibited significantly greater clonogenic ability than the control cells (P<0.001; Fig. 3D and E). Transwell assays further showed that HDAC1 O/E augmented both the migratory (Fig. 3F and G) and invasive (Fig. 3H and I) abilities of all three cell lines (P<0.001 for all comparisons). These gain-of-function data mirror the knockdown results and collectively established that HDAC1 promotes the proliferation, migration and invasion of NSCLC cells.

Next, the present study assessed the effect of HDAC1 O/E on downstream signaling molecules using western blotting and nuclear-cytoplasmic fractionation (Fig. 3J-M). Consistent with the knockdown data, HDAC1 O/E led to significant upregulation of c-Myc (P<0.01) and cyclin D1 (P<0.01) protein levels across all three cell lines, further supporting the role of HDAC1 in driving cell cycle progression. Regarding EMT-related markers, HDAC1 O/E resulted in a marked decrease in E-cadherin expression (P<0.01) and a corresponding increase in vimentin expression (P<0.01), indicating a shift toward a mesenchymal phenotype. Subcellular fractionation analysis demonstrated that nuclear β-catenin levels were substantially elevated in HDAC1-O/E cells (P<0.001), whereas cytoplasmic β-catenin remained largely unchanged (P>0.05), reinforcing the idea that HDAC1 facilitates nuclear accumulation of β-catenin. Consistent with the knockdown data, neither the phosphorylation of AKT (Thr308 and Ser473) nor ERK or total AKT and ERK protein levels were significantly affected by HDAC1 O/E (P>0.05 for all comparisons). Collectively, the complementary knockdown and overexpression results demonstrated that HDAC1 drives NSCLC cell proliferation, migration and invasion, at least in part through upregulation of c-Myc and cyclin D1, induction of EMT and promotion of nuclear β-catenin accumulation, independent of the PI3K/AKT and MAPK/ERK pathways.

To determine whether β-catenin signaling is functionally required for HDAC1-mediated oncogenic effects, the present study overexpressed β-catenin in HDAC1-K/D A549, H1299 and H1975 cells. Four experimental groups were established: Empty-vector control, HDAC1 K/D alone, β-catenin O/E alone and HDAC1 K/D + β-catenin O/E. As shown in Fig. 4A-C, CCK-8 assays revealed that HDAC1 K/D alone significantly suppressed the proliferative capacity of all three cell lines (P<0.001), whereas β-catenin O/E alone enhanced cell proliferation (P<0.001). Notably, co-expression of β-catenin in HDAC1-K/D cells restored proliferation to levels comparable to those of the β-catenin O/E group (P>0.05, HDAC1 K/D + β-catenin O/E vs. β-catenin O/E), indicating that β-catenin overexpression effectively abrogates the growth-inhibitory effect of HDAC1 silencing. Colony formation assays yielded concordant results; the reduced clonogenic ability caused by HDAC1 knockdown was effectively rescued by concomitant β-catenin overexpression (P>0.05, HDAC1 K/D + β-catenin O/E vs. β-catenin O/E; Fig. 4D and E). Similarly, Transwell assays demonstrated that β-catenin O/E largely reversed the inhibitory effects of HDAC1 K/D on both migration (P>0.05; Fig. 4F and G) and invasion (P>0.05; Fig. 4H and I) in all three cell lines. The double-modified group showed no significant difference compared with the β-catenin O/E-alone group for either endpoint. These results indicated that HDAC1 promotes NSCLC cell proliferation, migration and invasion, at least in part through β-catenin signaling.

Western blotting and nuclear-cytoplasmic fractionation further corroborated these functional observations at the molecular level (Fig. 4J-M). Consistent with earlier findings, HDAC1 K/D alone reduced c-Myc, cyclin D1 and vimentin protein levels while upregulating E-cadherin. Simultaneous overexpression of β-catenin in HDAC1-K/D cells effectively reversed these molecular changes: c-Myc, cyclin D1 and vimentin levels were restored and E-cadherin was suppressed to levels comparable with that of the β-catenin O/E-alone group (P>0.05, HDAC1 K/D + β-catenin O/E vs. β-catenin O/E). Subcellular fractionation revealed that nuclear β-catenin level, which was reduced following HDAC1 K/D (P<0.01), was substantially restored following β-catenin O/E (P>0.05), whereas cytoplasmic β-catenin levels remained relatively stable across all groups (P>0.05). Taken together, these results demonstrated that β-catenin functions as a critical downstream effector of HDAC1 in NSCLC cells. The tumor-promoting activities of HDAC1, including enhanced proliferation, migration, invasion and EMT, are largely dependent on β-catenin nuclear accumulation and its downstream transcriptional activity.

To validate the proposed molecular mechanism in vivo, the present study established a subcutaneous xenograft tumor model. A549 cells stably expressing empty vector (control), HDAC1 K/D, β-catenin O/E, or HDAC1 K/D + β-catenin O/E were injected subcutaneously into the flanks of nude mice (n=5 per group; Fig. 5A) and tumor growth was monitored over a 28-day period. HDAC1 K/D significantly suppressed xenograft tumor growth; both the final tumor volume and tumor weight in the HDAC1 K/D group were significantly reduced compared with those in the control group (P<0.001; Fig. 5B-D). Conversely, β-catenin O/E alone yielded significantly larger and heavier tumors than controls (P<0.001). Critically, simultaneous overexpression of β-catenin in HDAC1-K/D cells restored tumor volume and weight to levels comparable to those in the control group (P>0.05, HDAC1 K/D + β-catenin O/E vs. control; Fig. 5C-E), effectively abolishing the growth-inhibitory effect of HDAC1 silencing. Body weight remained stable in all four groups throughout the experimental period, indicating no overt systemic toxicity (Fig. 5F).

To determine whether the molecular changes observed in vitro are recapitulated in vivo, the present study performed western blotting of harvested tumor tissues (Fig. 5G and H). HDAC1 K/D reduced c-Myc, cyclin D1 and vimentin protein levels (P<0.01) and upregulated E-cadherin expression in xenograft tissues (P<0.01), which was consistent with the in vitro data. Importantly, concomitant β-catenin O/E in HDAC1-K/D cells effectively reversed these alterations: c-Myc, cyclin D1 and vimentin levels were restored and E-cadherin was suppressed to levels comparable to those in the β-catenin O/E-alone group (P>0.05, HDAC1 K/D + β-catenin O/E vs. β-catenin O/E). Subcellular fractionation of tumor lysates demonstrated that nuclear β-catenin, which was reduced following HDAC1 K/D (P<0.01), was substantially restored following β-catenin O/E (P>0.05, HDAC1 K/D + β-catenin O/E vs. β-catenin O/E), whereas cytoplasmic β-catenin levels remained relatively stable across all groups (P>0.05). Notably, β-catenin O/E did not alter HDAC1 protein levels (P>0.05), confirming that the observed rescue effect was mediated by β-catenin reactivation rather than by restoration of HDAC1 expression. In summary, these in vivo data are in full agreement with the in vitro observations and provide compelling evidence that HDAC1 promotes LUAD tumorigenesis through a β-catenin-dependent mechanism.