The shRNAs were synthesized as DNA oligos and then subcloned into the pLKO.1 expression plasmid (8453, Addgene). The lentivirus particles were generated by the co-transfection of pLKO.1 and two helper (pVSVG and pMD2.G) plasmids into human embryonic kidney 293T (HEK293T) cells (CRL-11268, ATCC) using the polyethylenimine reagent (101000029, Polyplus). The sequence of shRNAs is provided in Supplementary Table 1 .

The HEK293T cells were cultured in DMEM (11965-065, Thermo Fisher Scientific) containing 10% fetal bovine serum (FBS) (10099141C, Thermo Fisher Scientific) and 1% penicillin/streptomycin (15140122, Thermo Fisher Scientific). The colorectal carcinoma CT26 cells (CRL-2638, ATCC) were maintained in RPMI-1640 medium (11875093, Thermo Fisher Scientific) containing 10% FBS and 1% penicillin/streptomycin. For stable cell generation, lentivirus particles containing the control or lamin shRNA were incubated with the cells for 72 hours, which were then selected by 5 μg/mL puromycin (A1113803, Thermo Fisher Scientific) for one week. The desired cells were expanded later and cultured in 5 μg/mL puromycin. Murine macrophage cell line RAW264.7 (TIB-71, ATCC) and mammary cancer cell 4T1 (CRL-2539, ATCC) were also cultured in RPMI-1640 medium containing 10% FBS and 1% penicillin/streptomycin. All cells were cultured at 37°C with 5% CO2. To explore the downstream pathway underlying lamin knockdown, control and lamin-deficient cells were seeded into 12-well culture plates at a density of 1.5 × 10⁵ per well, and then treated with the small-molecule inhibitor of selective pathways. These inhibitors were listed as below: BAY872243 (SC1193, Beyotime), FR180204 (HY-12275, MedChemExpress), GSK8612 (HY-111941, MedChemExpress), Sorafenib (HY-10201, MedChemExpress), and/or Rosiglitazone (HY-17386, MedChemExpress).

The animal research conducted in this study was in accordance with the Scientific and Ethical Committee of the Shanghai First Maternity and Infant Hospital, School of Medicine, Tongji University. Four-week-old BALB/c and BALB/cNj-Foxn1nu/Gpt mice were purchased from GemPharmatech (Nanjing, China) and then housed in a constant temperature (24-26°C) and humidity (40-60%) room with a 12-hour light-dark cycle. The cancer cells without or with lamin knockdown were subcutaneously injected into the left side of 5-week-old mice at a dosage of 5 × 10⁵ cells/mice. For measuring the responsiveness of lamin-deficient tumor to clinically anticancer drugs, 10 mg/kg sorafenib was infused daily into the mice via intragastric administration from posttransplant day 8. The tumor volumes were periodically determined with a manual caliper and calculated using the below formula: ½ × longitudinal diameter (length) × the greatest transverse diameter (width)². When their volume reached 2000 mm³, the tumors were harvested for further analysis.

The cells without or with lamin were plated in a 6-well culture plate at a density of 1 × 10³/well and then cultured at 37°C, 5% CO₂ for 2 weeks. These cells were stained with crystal violet and then visualized using an Olympus CK30-F200 microscope.

PerkinElmer 96-well culture plate was coated with the laminin at a concentration of 1μg/cm² for 24 hours. Single cell population was seeded into the above 96-well plate at a density of 1 × 10⁴/well and then maintained at 37°C for 120 hours. Cell behaviors were scanned and imaged every 4 hours using the Livecyte^® Kinetic Cytomete (phasefocus) equipped with a sCMOS camera and 20×Plan Phase objective (Olympus). The cell index was calculated with Livecyte software after normalization to the initial image.

This EdU proliferation assay was measured using BeyoClick™ EdU-555 kit (C0075S, Beyotime). EdU is a thymidine analog and labels the dividing cell by incorporating into nascent DNA during the S-phase. The EdU solution was added into the culture medium at a final concentration of 10 μM following the manufacturer’s instruction. Two hours later, the cells were washed with PBS (C0221A, Beyotime) and then fixed in 4% paraformaldehyde (P0099, Beyotime) for 15 minutes (mins). After their permeabilization with 0.3% Triton X-100, the cells were incubated in Click Additive Solution for 30 mins at room temperature (RT). In the end, DAPI was added and treated for 10 mins. The cells were imaged using an Olympus BX63F microscope.

Cancer cells were digested into single cell and then resuspended in fresh culture medium. Hoechst33342 (C1022, Beyotime) was added into the medium at a final concentration of 10 μg/ml and then incubated at 37°C for 45 mins. After three washes, the cells were run on the BD Fortessa machine. Apoptosis assay was performed following the manufacturer’s protocol (#C1062S, Boomtime). The data were analyzed using the FlowJo.

For surface marker analysis, cancer or RAW264.7 cells were suspended in cell staining buffer (FXP005, 4abio) and stained with specific fluorochrome-conjugated antibodies at appropriate concentrations as previous reports (26, 27). For the engrafted tumor, a portion of tumors were digested into single cells, filtrated through 70 μm cell strainers, and then resuspended in cell staining buffer. Zombie NIR Fixable kit (423105, Biolegend) was used to determine live or dead cells. Tumor cells were incubated with fluorochrome-conjugated antibodies at RT for 1 hour. These antibodies were listed below: APC anti-mouse CD3 (100236, Biolegend); FITC anti-mouse CD4 (100405, Biolegend); Brilliant Violet 421 anti-mouse CD8 (100737, Biolegend); PE anti-mouse CD69 (104507, Biolegend); APC anti-mouse PD1 (135210, Biolegend); FITC anti-mouse CD44 (156008, Biolegend); PE anti-mouse CD133 (141203, Biolegend); Brilliant Violet 421 anti-mouse CD47 (127527, Biolegend); PE anti-mouse H-2Kb (116508, Biolegend); APC anti-mouse PD-L1 (124311, Biolegend); FITC anti-mouse CD80 (104705, Biolegend); PE/Cy7 anti-mouse CD86 (105014, Biolegend). Following three washes, the stained cells were analyzed on BD Fortessa machine (BD Biosciences), and the data were processed with the FlowJo software.

The tumors were washed with PBS and then fixed in 4% paraformaldehyde overnight at 4°C. The samples were embedded in the paraffin tissue blocks, cut into 10-μm sections, and then transferred onto Bond-Rite™ slides. After being heated at 60°C for 60 mins, the sections were deparaffinized in fresh xylene and then rehydrated with descending grades of alcohol. The sections were soaked in blocking buffer (P0260, Beyotime) for 20 mins and then stained with primary antibodies at 4°C overnight. These primary antibodies were listed below: Rabbit anti-mouse CD3 (78588, Cell Signaling Technology); Rabbit anti-mouse CD8 (98941, Cell Signaling Technology); Rabbit anti-mouse STING (13647, Cell Signaling Technology); Rabbit anti-mouse Ki67 (12202, Cell Signaling Technology). After three washes, the sections were successively incubated with secondary antibody and DAPI for 3 hours. The coverslips were then applied with permanent synthetic mounting media. All pictures were viewed and taken under an inverted microscope.

Cells were dissociated in RIPA lysis buffer (P0013B, Beyotime) containing proteinase inhibitor cocktail. After protein concentration was determined, total proteins were separated on SDS-PAGE gels, separated at 200V for 1 hour, and transferred onto the PVDF membranes. The membranes were blocked in 5% non-fat milk/TBST buffer and then incubated overnight with primary antibodies as previous reports (28, 29). Primary antibodies were listed below: NANOG (A22625, ABclonal), ALDH1A1 (A0157, ABclonal), DDB1 (A2896, ABclonal), Ku70 (A0883, ABclonal), Ku80 (A5862, ABclonal), RAD50 (A3078, ABclonal), RAD51 (A2829, ABclonal), BRCA1 (A11034, ABclonal), NBS1 (7703, ABclonal), DDB1 (A2896, ABclonal), DDB2 (A1848, ABclonal), Tubulin (11224-1-AP, Proteintech), INO80 (ab105451, Abcam), MRE11 (A2559, ABclonal), DDB2 (A1848, ABclonal), Rad23B (A1034, ABclonal), XPC (A8354, ABclonal), Phospho-STING (50907, Cell Signaling Technology), SDHA (11998, Cell Signaling Technology), PHB1 (2426, Cell Signaling Technology), VDAC (4661, Cell Signaling Technology), HSP60 (12165, Cell Signaling Technology), COX IV (4850, Cell Signaling Technology), Vinculin (4650, Cell Signaling Technology), ERK1/2 (4695, Cell Signaling Technology), Phospho-ERK1/2 (4370, Cell Signaling Technology), IL-1β (A16288, ABclonal), NLRP3 (15101, Cell Signaling Technology), TBK1 (38066, Cell Signaling Technology), Phospho-TBK1 (5483, Cell Signaling Technology), cGAS (15102, Cell Signaling Technology), Phospho-STING (50907, Cell Signaling Technology), STING (19851-1-AP, Proteintech), Phospho-STAT1 (8826, Cell Signaling Technology), Phospho-STAT3 (9145, Cell Signaling Technology), PPAR-γ (2443, Cell Signaling Technology), Phospho-P62 (13121, Cell Signaling Technology), LMNA (AF7350, Beyotime), LMNB1 (AF1408, Beyotime), LMNB2 (AF0219, Beyotime). Following three washes, the membranes were probed with HRP-conjugated secondary antibodies at RT for 2 hours. The target proteins were viewed with an ECL Plus Kit (Abcam). ImageJ software was used to view protein intensity.

Total mRNAs were purified from the cells with Trizol reagent kit (Invitrogen, USA) according to the manufacturer’s instructions. They were reversely transcribed into cDNAs using the HiScript II Q Select RT SuperMix kit (R232-01, Vazyme). The reaction of quantitative PCR was performed with SYBR Green PCR Master Mix (Q111-02, Vazyme). The 2^-ΔΔCt method was used to calculate the relative expression level of target genes after normalization to the internal gene GAPDH. The sequences of RT-PCR primers are listed in the Supplementary Table 2 .

Total RNA quality was determined using Agilent 2100 Bioanalyzer (Agilent Technologies, USA). Eukaryotic mRNA was enriched by Oligo(dT) beads, fragmented into short fragments, and reversely transcribed into cDNA using the Next Ultra RNA Library Prep Kit for Illumina (7530, NEB). These double-stranded cDNA fragments were added with A-base, ligated to sequencing adapters, and then purified with the AMPure XP Beads. The cDNA library was sequenced using Illumina Novaseq 6000 by Gene Denovo Biotechnology Co. (Guangzhou, China).

Cancer cells without or with lamin knockdown were washed two times in PBS and then changed with fresh culture medium containing 10 μg/ml Hoechst33342. One hour later, the cells were washed with culture medium and viewed under the Leica TCS/SP8 microscope.

Quantification was derived from at least three independent experiments and shown as mean ± SEM. The statistical significance was determined by Student’s t-test or by one-way analysis of variance (ANOVA). P-value of less than 0.05 was thought to be significant.

To evaluate the pathological significance of lamin in tumor progression, we analyzed the expression profiles of lamin in various human cancers via the cBioPortal Cancer Genomics database. The expression levels of LMNA, LMNB1, and LMNB2 mRNAs were higher in most tumors than in normal adjacent tissues, such as bladder urothelial carcinoma (BLCA), colon adenocarcinoma (COAD), kidney renal clear cell carcinoma (KIRC), liver hepatocellular carcinoma (LIHC), pancreatic adenocarcinoma (PAAD), and thyroid carcinoma (THCA) ( Supplementary Figures 1A-C ). The transcriptome analysis showed that the COAD tumors had more lamin transcripts than normal tissue ( Figure 1A ), supporting the upregulation of LMNA, LMNB1, and LMNB2 expressions in tumor tissues. The COAD patients with low lamin expression likely presented better survival outcomes in comparison to the patients bearing high lamin level ( Supplementary Figure 1D ). To explore if lamin is essential in cancer cell plasticity, the CD44⁺ CD133⁺ cell population was sorted out from murine CT26 cancer cell and then applied in the below experiments. Both CD44 and CD133 are widely used surface markers for cancer stem-like cells (22, 30, 31). These cancer cells with stem-cell-like properties were engineered for the simultaneous knockdown of three lamin subtypes via the shRNA-mediated knockdown system. The knockdown efficiencies of LMNA, LMNB1, and LMNB2 were confirmed at both transcript and protein levels ( Figure 1B , Supplementary Figure 2A ). The expression of stemness marker CD133 was significantly decreased by lamin knockdown ( Figure 1C ). Consistently, lamin knockdown reduced the levels of pluripotent markers ALDH1 and NANOG ( Figure 1D ), indicating the stemness reduction after lamin knockdown. In line with this notion, lamin knockdown suppressed the colony formation ability of single cancer cell ( Figure 1E ). The CCK8 assay showed that lamin-deficient cell population exhibited a lower proliferation rate in comparison to the control cells ( Figure 1F ). Single-cell tracking assay revealed that lamin-deficient cell had a much weaker proliferative capacity than the control cell ( Figures 1G, H ), leading to the fewer cell number and the lower M-phase cell percentage at 96-hour time point ( Figures 1H, I ). Moreover, a single lamin-deficient cell needed more time to finish its double period, as demonstrated by the higher doubling time of lamin-deficient cell than the control cell ( Figure 1J ). Additionally, cell cycle analysis showed that lamin knockdown gave rise to the increase of the G1 phase and the reduction of S and G2/M phases ( Figures 1K, L ). Lamin knockdown resulted in the notable reduction of EdU⁺ cells ( Figures 1M, N ), suggesting the inhibition of lamin deficiency on cancer cell proliferation. However, lamin knockdown was found to decrease cell apoptosis, as evidenced by the lower percentages of Annexin-V⁺ or CASP3⁺ cells ( Figure 1O , Supplementary Figure 2B ). Thus, the inhibition of lamin knockdown on cancer cell proliferation and stemness was not due to the reduced apoptosis. Lamin-deficient cancer cell was shown to display anomalous nuclear shape in comparison to the round nuclear phenotype of control cell ( Supplementary Figure 2C ), indicating that all lamin knockdown might trigger nuclear anomaly. Collectively, these data suggest that lamin had higher expression levels in many cancers and its knockdown inhibited cancer cell stemness and proliferation.

To investigate how lamin deficiency affects cellular processes, we conducted transcriptome sequencing without or with lamin knockdown. Bioinformatic analysis showed that lamin knockdown changed the expression profiles of many functional genes, resulting in the downregulation or upregulation of 2667 or 1570 differently expressed genes (fold change ≥ 2) respectively ( Figures 2A, B ). A substantial number of the differently expressed genes were enriched in the processes, including but not limited to DNA binding, protein binding, transferase activity, phosphofructokinase activity, and biosynthetic and metabolic regulation ( Figure 2C , Supplementary Figure 2D ). Since DNA-binding proteins have critical roles in DNA repair pathways, we next tried to measure the influence of lamin knockdown on the genomic repair process. A sensitive fluorescent reporter assay, in which the I-SceI endonuclease creates genomic damage at the target locus (32, 33), was employed to determine the efficiency of homologous recombination (HR), non-homologous end joining (NHEJ), or nucleotide excision repair (NER). The successful repairment of genomic damage by the above DNA repair pathway led to the expression of the relative GFP reporter. Lamin knockdown was found to increase the repair efficiency of HR and NER pathways, as evidenced by the higher GFP⁺ cell proportion in lamin-deficient cells ( Figures 2D, E ). However, lamin deficiency did not affect the efficiency of the NHEJ repair pathway ( Figure 2F ). Western blotting analysis revealed that lamin knockdown led to the selective decrease in the levels of repair-related factors RAD50, RAD51, Ku80, NBS1, and RAD23B ( Figure 2G ). Conversely, other repair-associated proteins such as INO80 and MRE11 remained unchanged ( Figure 2H ). It is notable that lamin-deficient cancer cells might possess abnormal cellular components such as intracellular organelles ( Supplementary Figure 2E ). Given the importance of mitochondria in cellular metabolism and energy production, we then assessed the impact of lamin loss on mitochondrial function. GSEA analysis revealed that both mitochondrial gene expression and protein translation were suppressed by lamin knockdown ( Figure 2I ). Although having no affection on mitochondrial membrane potential ( Figure 2J ), lamin knockdown reduced the expression levels of functionally mitochondrial proteins SDHA, PHB1, VDAC, and HSP60 ( Figure 2K ). This observation suggests that mitochondrial activity in lamin-deficient cells was lower than in the control cells. Lamin-deficient cells were found to display less reactive oxygen species (ROS) than the control cell, as demonstrated by the lower fluorescent staining intensity in FACS assays ( Figures 2L ). Among the changed metabolic processes after lamin knockdown ( Supplementary Figure 2D ), the oxidative phosphorylation was likely decreased by lamin knockdown ( Figure 2M ). Nevertheless, the amount of oxidized lipids in the lamin-deficient cell was comparable to that in the control cell ( Supplementary Figure 2F ), suggesting that lamin knockdown had no affection on lipid oxidization. Therefore, these results indicate that lamin knockdown altered DNA repair choices and impaired mitochondrial activity.

To explore whether lamin knockdown in cancer stem-like cell affects tumorigenesis in vivo, we performed the subcutaneous tumor experiment in Balb/c mice ( Figure 3A ). In brief, cancer cells without or with lamin knockdown were infused into the right subcutaneous flank of 5-week-old mice, and their relevant tumors were monitored periodically. Lamin knockdown was shown to inhibit dramatically the growth of xenograft tumors ( Figure 3B ), resulting in the smaller volume and lower weight of lamin-deficient tumors ( Figures 3C, D ). The mice bearing either lamin-deficient or control tumors had gradually increased body weights ( Figure 3E ), suggesting their normal diet. There was no significant alteration in mouse spleen weights ( Supplementary Figure 3A ). Immunohistochemistry staining revealed that the Ki67 proliferation index was slightly lower in the lamin-deficient tumor ( Figure 3F, G ), indicating that the suppression of lamin knockdown on tumorigenesis may not be due to reduced cancer cell proliferation. Since the immune score is positively linked with patient prognosis, cancer patients with LMNA mutation were found to present higher infiltration score of immune cells such as cytotoxic CD8⁺ T and dendritic cells ( Supplementary Figures 3B, C ). Indeed, lamin-deficient tumors were shown to exhibit more CD3⁺ and CD8⁺ T lymphocytes than the control xenografts ( Figures 3H–K ). FACS analysis confirmed that the proportions of CD3⁺ T, CD4⁺ CD3⁺ T, and CD8⁺ CD3⁺ T lymphocytes were significantly higher in the lamin-deficient tumors ( Figures 3L–O ). Additionally, lamin-deficient tumors presented a much higher frequency of CD69⁺ CD3⁺ T cells than the control xenografts ( Figures 3P, Q ). These results suggest that lamin knockdown in cancer cells enhanced the infiltration and activation of cytotoxic T lymphocytes in the lamin-deficient tumor. Moreover, the expression level of immune checkpoint PD-1 on tumor-infiltrating CD8⁺ T cells was much lower in the lamin-deficient tumors than in the control xenografts ( Figure 3R ), suggesting the increased T-cell functionality in the context of lamin knockdown.

To test whether lamin knockdown impedes tumor growth mainly by enhancing T cell infiltration and activation, we conducted the subcutaneous xenograft tumor experiment in immunodeficient nude mice lacking T cells ( Supplementary Figure 3D ). Unexpectedly, the growth and weight of lamin-deficient tumors were comparable to that of the control xenografts in nude mice ( Supplementary Figures 3E, F ). The body weights of nude mice carrying either lamin-deficient or control tumors increased gradually over time ( Supplementary Figure 3G ). Thus, lamin knockdown within cancer cells could not inhibit tumor progression in T-cell-deficient nude mice. Moreover, we employed the in vitro co-culture system to measure if lamin knockdown in cancer cell affects other immune cells such as macrophage. The conditional medium of cancer cells was incubated with macrophage Raw264.7 cells for at least 48 hours. When compared to the control medium, the conditional media of lamin-deficient cancer cells did not change NLRP3 inflammasome and STING/TBK1 pathways in macrophage cells ( Supplementary Figure 4A ). The expressions of MHC molecule H-2Kb and co-stimulator factors (CD80 and CD86) on the surface of macrophages were not affected by the co-culture of lamin-deficient cancer cells ( Supplementary Figures 4B–D ), suggesting that lamin knockdown in cancer cell might not influence macrophage activation. Collectively, these data indicate that lamin knockdown within cancer stem-like cell restrained the in vivo tumorigenesis via boosting the infiltration and activity of tumor-infiltrating T cells.

Given the promotion of lamin-deficient cancer cells on T-cell activation and infiltration, we next tried to assess whether lamin knockdown regulates immune checkpoint dynamic in cancer cell. The PD-L1 on cancer cell is a well-known immune checkpoint molecule impairing antitumor immunity via interaction with PD1 on T cell (34). The CD274 mRNA encoding PD-L1 protein was significantly reduced by lamin knockdown ( Figure 4A ). FACS analysis confirmed that lamin knockdown led to a notable reduction in PD-L1 protein expression on cancer cells ( Figure 4B ). Conversely, the expression levels of CD47 and CD86 proteins on cancer cells were not affected by lamin knockdown ( Figures 4C, D , Supplementary Figure 5A ). CD47 is a surface receptor on cancer cell and delivers the “don’t eat me” signal by interacting with SIRPα on myeloid cells (35), while costimulatory molecules CD80 and CD86 participate in the regulation of antigen presentation and T cell activation (36, 37). There was a slight alteration in CD80 and H-2Kb expression after lamin knockdown ( Figures 4E, F , Supplementary Figures 5B, C ), suggesting their minimal contribution to the anti-tumor effect of lamin knockdown. H-2Kb is a classical MHC-I molecule presenting tumor antigen peptides to cytotoxic T cell (38). Interestingly, lamin-deficient cancer cells were found to present higher expression levels of many inflammatory chemokines such as CCL2, CCL9, CCL22, CXCL9, and CXCL10 ( Figures 4G, H ). Lamin knockdown seemed not to affect the IFNGR2 and IFNAR1 levels ( Figure 4I ). IFNGR1/2 and IFNAR1, serving as the receptor for IFN-γ and IFN-α respectively, are critical in cancer cell fate and tumor microenvironment (39, 40). Given the importance of cGAS-STING pathway in immune checkpoint signaling (28, 41), we then examined how lamin knockdown impacts the cGAS-STING pathway. The transcriptional and protein levels of cGAS, STING, and TBK1 were not changed by lamin knockdown ( Figure 4J , Supplementary Figure 5D ). Immunohistochemistry analysis revealed that lamin-deficient tumor had a comparable STING intensity than the control xenograft ( Figures 4K, L ), indicating that the STING/TBK1 pathway may not be the downstream mechanism. Remarkably, bioinformatic analysis showed that lamin knockdown could regulate the expressions of many functional genes enriched in AMPK, MAPK, and HIF-1 signaling pathways ( Figure 4M , Supplementary Figure 5E ). Indeed, lamin knockdown gave rise to the downregulation of STAT1 phosphorylation and the upregulation of ERK phosphorylation and PPARγ ( Figure 4N ). Both HIF-1α and H3K36Me3 levels were also increased by the lamin knockdown ( Figure 4O ). To determine which pathway is the major one, we treated the control and lamin-deficient cancer cells with pathway inhibitor or activator. Both ERK inhibitor FR180204 and PPARγ agonist Rosiglitazone were found to generate a slight restoration on the lower levels of PD-L1 and CD133 caused by lamin knockdown ( Figures 4P–S ). However, neither ERK inhibitor nor PPARγ agonist could influence CD47 expression in lamin-deficient cancer cells ( Supplementary Figures 5F, G ). Unexpectedly, the selective TBK1 inhibitor GSK8612 reduced the PD-L1, CD133, and CD47 expressions in the absence or presence of lamin knockdown ( Supplementary Figures 5H–J ). Consistent with the promotion of lamin knockdown on HIF-1 signaling ( Figures 4O, T ), the HIF-1 inhibitor BAY872243 was shown to rescue the suppression of lamin knockdown on immune checkpoint PD-L1 and stemness marker CD133 ( Figures 4U, V ). The BAY872243 treatment generated an enhancement of CD47 level on lamin-deficient cell rather than the control cell ( Figure 4W ). The BAY872243 treatment led to the reduction of cellular apoptosis ( Figure 4X ), indicating the safety of its application dosage. Moreover, the BAY872243 was found to restore slightly the inhibition of lamin knockdown on cell cycle progression, as evidenced partially by the increase of G2/M phases in lamin-deficient cells with BAY872243 treatment ( Supplementary Figures 5K–M ). Overall, these data suggest that lamin knockdown modulated cancer cell stemness and inflammatory factors through the HIF-1 pathway.

To evaluate the responsivity of lamin-deficient cancer cells to anti-tumor drugs, we injected the immunocompetent mice with cancer cells and then administrated the chemotherapeutic agent sorafenib periodically ( Figure 5A ). Sorafenib, a multiple-target tyrosine kinase inhibitor, has been used clinically for the treatment of many solid tumors such as hepatic cancer through inhibiting cancer cell proliferation and inducing apoptosis (42, 43). The sorafenib inhibited xenograft tumor growth in comparison to the vehicle treatment ( Figure 5B ). Notably, the inhibition extent of lamin knockdown on tumor growth was much stronger than the sorafenib application ( Figure 5B , Supplementary Figure 5N ). The Balb/c mice receiving sorafenib treatment displayed slightly lower body weights at the experimental endpoint ( Figure 5C ). The sorafenib administration was found to generate smaller tumor volume and weight even in the presence of lamin knockdown ( Figures 5B–D ), presenting the enhanced sensitivity of lamin-deficient cancer cells to the chemotherapeutic agent. The spleen weights from different groups of mice were comparable to each other, except for the mice with lamin-deficient tumor and sorafenib treatment ( Figure 5E ). Although lamin-deficient cancer cells had lower PD-L1 and stemness markers ( Figures 1C , 4B ), the sorafenib did not alter PD-L1 and CD133 levels on cancer cells with or without lamin knockdown ( Figures 5F, G ). Unlike the lamin knockdown, the sorafenib led to a decrease in CD47 expression ( Figure 5H ). The level of CD44 protein in the control cell, rather than in lamin-deficient cell, was reduced by the sorafenib ( Figure 5I ). Moreover, the sorafenib increased the percentage of the G1 phase and decreased the proportion of the G2/M stages ( Figures 5J–L ), which is similar to the lamin knockdown. The combination of lamin knockdown and sorafenib could induce the highest proportion of G1 phase and generate the lowest percentage of G2/M phases ( Figures 5J–L ). These findings indicate that lamin knockdown in cancer cells enhanced their sensitivity to chemotherapy.

To determine whether all lamin knockdown changes PD-L1 expression and tumorigenesis in another cell model, we engineered the three lamin subtypes in murine breast cancer cell line using above shRNA-mediated knockdown system. The transcript levels of LMNA, LMNB1, and LMNB2 were dramatically reduced by the lamin knockdown ( Supplementary Figure 6A ). FACS analysis revealed that lamin-deficient breast cancer cells had much lower PD-L1 protein level and intracellular ROS amount ( Supplementary Figures 6B, C ). After subcutaneous injection into the immunocompetent Balb/c mice, lamin-deficient breast cancer cell was found to grow slowly ( Supplementary Figure 6D ). Consequently, the volume and weight of lamin-deficient breast tumors were much lower than the control xenografts ( Supplementary Figures 6E, F ). There existed no alteration in mouse spleen weights ( Supplementary Figure 6G ). Thus, these data suggest that lamin knockdown also impeded PD-L1 expression and subcutaneous tumor progression in second cancer cell model.