{'fraction_non_alphanumeric': 0.03657161803713528, 'fraction_numerical': 0.01694297082228117, 'mean_word_length': 4.9914580850218515, 'pattern_counts': {'":': 0, '<': 4, '<?xml version=': 0, '>': 0, 'https://': 0, 'lorem ipsum': 0, 'www.': 0, 'xml': 0}, 'pii_count': 0, 'substrings_counts': 0, 'word_list_counts': {'cursed_substrings.json': 0, 'profanity_word_list.json': 0, 'sexual_word_list.json': 0, 'zh_pornsignals.json': 0}}
Structural and functional analysis of cell adhesion and nuclear envelope nano-topography in cell death

The cell death mechanisms of necrosis and apoptosis generate biochemical and morphological changes in different manners. However, the changes that occur in cell adhesion and nuclear envelope (NE) topography, during necrosis and apoptosis, are not yet fully understood. Here, we show the different alterations in cell adhesion function, as well as the topographical changes occurring to the NE, during the necrotic and apoptotic cell death process, using the xCELLigence system and atomic force microscopy (AFM). Studies using xCELLigence technology and AFM have shown that necrotic cell death induced the expansion of the cell adhesion area, but did not affect the speed of cell adhesion. Necrotic nuclei showed a round shape and presence of nuclear pore complexes (NPCs). Moreover, we found that the process of necrosis in combination with apoptosis (termed nepoptosis here) resulted in the reduction of the cell adhesion area and cell adhesion speed through the activation of caspases. Our findings showed, for the first time, a successful characterization of NE topography and cell adhesion during necrosis and apoptosis, which may be of importance for the understanding of cell death and might aid the design of future drug delivery methods for anti-cancer therapies.


DOX completely induced necrosis, while ETO induced necrosis in combination with apoptosis.
We investigated apoptotic and necrotic cell death, caused by DOX and ETO, by evaluating cell viability, caspase-3/7 activity, and Annexin V/propidium iodide (PI) staining. For cell viability analysis, DOX (1 μ M) and ETO (50 μ M) treatment were used to induce cytotoxicity in a time-dependent manner (half maximal inhibitory concentration; IC50 induced at 72 h) ( Supplementary Fig. S1A). ETO-treated cells exhibited significantly higher caspase-3/7 activity and percentage of cells positively stained with PI (indicating necrosis) and Annexin V (indicating apoptosis) simultaneously, than control cells. On the other hand, DOX-treated cells exhibited a higher percentage of PI-positive cells than the control, and this increase was almost independent of caspase-3/7 activity and Annexin V-positive cells ( Supplementary  Fig. S1B, C). These findings suggest that DOX-treatment definitively induced necrosis, but ETO-treatment simultaneously induced necrosis and apoptosis (referred to as nepoptosis).
Necrosis and nepoptosis generated cell swelling, but yield different effects on cell adhesion. In our experiments, we observed that DOX-and ETO-treatment of cells commonly resulted in an expanded cell adhesion area on culture dishes. Moreover, almost all DOX-treated cells attached to the dish, but a large number of ETO-treated cells remained in the culture supernatant, indicating a loss of cell adhesion capability (data not shown). Thus, we determined that necrosis and nepoptosis induced different alterations in the cells' ability to adhere in culture. The measurement of alterations in cell size, adhesion, and morphology caused by necrosis and nepoptosis were confirmed by the use of a hemocytometer, fluorescence activated cell sorting (FACS), xCELLigence technology and CNT/AFM probe methods (Fig. 1A).
We demonstrated that cell size of DOX-and ETO-treated cells was visually increased, via analysis on a hemocytometer, when compared to the control cells (Fig. 1B). Using forward scatter light (FSC, indicative of cell size) analysis in FACS, we determined the induced FSC levels from 1 × 10 4 cells (Fig. 1C). ETO (green color)-treated cells were harvested after 72 h and cell adhesion was measured, including cell index (CI) and saturation times (ST) detected using xCELLigence for 10 s intervals and monitored for 3 h. Black closed diagram indicate saturation (± 1.0%) times. (E,F) Table shows the cell size (FSC), adhesion area (CI), and saturation times (ST; h) at which cell adhesion was examined using computational analysis (described in results) compared to the control. The histogram shows cell adhesion levels calculated using the derived formula (described in results) for DOX-and ETO-treated cells for 72 h. (G) DOX-and ETOtreated cells were seeded in culture dishes for 3 h. Expression level of F-actin (red) was measured using immunofluorescence staining and detected by confocal microscopy. Hoechst 33258 (blue) was used for nuclear staining and scale bar represents 50 μ m. All histograms represent statistical analysis (P-value of *P < 0.05, **P < 0.01). Subsequently, we investigated alterations in cell adhesion ability caused by DOX and ETO, using the xCELLigence system. The xCELLigence system is used for cell-based measurements for analysis, including cell viability, invasion, and migration, by electrical impedance (cell index [CI]) in real-time using a gold microelectrode pattern device 31,32 , but alterations in cell adhesion are not usually determined during cell death. Here, we suggested a new application of this method to measure changes in cell adhesion, and established a formula to calculate cell adhesion alteration caused by necrosis and nepoptosis. We harvested DOX-and ETO-treated cells at 72 h (only attached cells on culture dish; supernatant cells were removed), seeded them (1 × 10 5 /well) into devices, and used the xCELLigence system to measure CI and saturation time (ST) at 10 s intervals, for 3 h. Results from the xCELLigence system showed that the CI level was significantly higher (3.2-fold increase) in DOX-treated cells than in control cells, but the ST was not affected. The CI for ETO-treated cells was also higher (2.1-fold increase) than the control cells, and the ST was delayed (90 min to 150 min) (Fig. 1D). The CI value is indicative of the cell adhesion area, and the CI ratio represents the increase from the cell size (FSC) ratio. Accordingly, when the CI ratio value is divided by the FSC ratio value, a value greater than 1.0 indicates the extension of the cell adhesion area, and generation of cell swelling, but values less than 1.0 indicate a loss of cell adhesion. Acceleration or deceleration of ST indicates the increase or decrease in cell adhesion speed, respectively. Based on these assumptions, we established the following formula to calculate cell adhesion (with each value described as shown in Fig. 1E (Fig. 1F). Similarly, DOX-and ETO-treated cell size was initially increased from the size of the control cells, at 0 min, and progressive cell swelling was observed, using real-time live-cell imaging analysis (Supplementary Fig. S2A and Movie S1). DOX-treated cells were almost fully attached to the culture dish, but attachment of ETO-treated cells was less than DOX-treated cells ( Supplementary Fig. S2B and Movie S1). Analysis of diameter to adhesion cells using phase contrast microscopy showed that control and DMSO-treated cells appeared approximately 20 ~ 30 nm in diameter, but DOX-and ETO-treated cells appeared approximately 60 ~ 100 nm in diameter at 3 h ( Supplementary  Fig. S2B). When measured using confocal microscopy, control and DMSO-treated cells indicated lower cell spreading area (averagely control: 243.7 μ m 2 and DMSO: 230.9 μ m 2 ), and DOX-and ETO-treated cells indicated higher cell spreading area (averagely DOX: 8188.7 μ m 2 and ETO: 7269.6 μ m 2 ). The expression of cytoskeleton protein such as F-actin in the cytoplasm was observed in control, DMSO and DOX groups. However, ETO-treated cells showed significantly decreased expression levels when compared with DOX group (Fig. 1G). Thus, necrotic cell death induced the expansion of the cell adhesion area, but did not affect the speed of cell adhesion, while apoptotic cell death reduced the cell adhesion area and cell adhesion speed.

Preservation of necrotic morphological changes during necrosis and nepoptosis. Cell swelling
is an important indicator of necrosis; another characteristic of necrosis induction is the rupturing of the plasma membrane. In our study, DOX-and ETO-treated cells also illustrated cell swelling and expansion of cell adhesion area compared with control and DMSO groups ( Fig. 2A). Therefore, we examined plasma membrane topography of adherent DOX-and ETO-treated cells, using CNT/AFM probes. AFM has been identified as a reliable, convenient technique for high-resolution investigation of nanostructures and biological materials at a nanometer scale, and AFM image resolution is dependent on the tip sharpness. In a previous report, we demonstrated that the CNT/AFM probes fabricated by the Langmuir-Blodgett technique had high-resolution imaging capability, for visualizing nanostructures and biological materials, such as nanoporous alumina membranes and plasmid DNA, respectively, to accurately identify their morphological traits 33 . In our study, AFM images showed that HK-2 and DMSO-treated cells appeared to have a low number of ruptures in their plasma membrane, when collected with trypsin-EDTA, showing a only small amount of damage was caused by the collection technique. On the other hand, DOX-and ETO-treated cells were shown to definitively induce necrotic morphological changes as evidenced by cell swelling and the significant plasma membrane rupturing observed at the nanometer scale ( Fig. 2 and Supplementary Fig. S3). In the roughness analysis on the plasma membrane at 2 μ m scale, control and DMSO-treated cells appeared 37.5 nm and 31.2 nm roughness, respectively. However, DOX-and ETO-treated cells significantly increased roughness to 53. Necrosis and nepoptosis generated nucleus swelling and altered nucleus morphology. In our experiments, DOX-and ETO-treated cells exhibited an increase in nucleus size, which is a typical morphological characteristic observed during necrotic cell death. Therefore, we used nucleus targeting dyes, Hoechst 33258 and PI, to assess nucleus area, and quantitatively measured the sizes of the nuclei using a Cellomics ArrayScan HCS reader system and analyzing at least 200 cells per well. This computerized system automatically provides various analyses, including cell viability, protein expression, and morphological changes, based on the acquired nucleus area and fluorescence intensity parameters. When compared to the control cells, the area of the nuclei of DOX-and ETO-treated cells, as well as the amount of blue staining from the Hoechst 33258 dye (visual representation of nucleus swelling by confocal microscopy analysis) was dramatically increased ( Supplementary Fig. S4A,D). In addition, Hoechst 33258 and PI fluorescence intensities were higher in the nuclei of DOX-and ETO-treated cells than control cells, but relatively, the intensity was lower in the nuclei of ETO-treated cells when compared to that of the DOX-treated cells ( Supplementary Fig. S4B,C). Based on these observations, we hypothesized that necrosis and nepoptosis regulate nucleus morphology and NE topography in different manners, and we used three measurement methods; hemocytometer analysis, confocal microscopy and CNT/AFM probes, to directly evaluate this (Fig. 3A). Preferentially, we used NE-PER Nuclear and Cytoplasmic Extraction Reagents Kit for accurate and secure nucleus extraction. The extraction kit contained Cytoplasmic Extraction Reagent I and II, and Nuclear Extraction Reagent. Cytoplasmic Extraction Reagent I causes cell membrane disruption but not disruption of the NE, while Cytoplasmic Extraction Reagent II inhibits the activity of Cytoplasmic Extraction Reagent I. The nuclear extracts were seeded onto a hemocytometer to visualize nucleus swelling that was definitively shown to result in increased nucleus size, when compared to the control on an equal scale (Fig. 3B). Interestingly, we observed that when extracting the nuclei of ETO-treated cells a lot of debris and leaked DNA were present in the supernatant from the nuclear extraction, when compared to the supernatant from DOX-treated cell nuclear extraction, in which neither debris nor leaked DNA were observed. Moreover, we observed that extracted nuclei attached easily to the coverslip and cell culture dish. For that reason, the extracted nuclei were seeded on coverslips and cell culture dishes for a short time (15 minutes), in order to remove the debris, then washed with PBS for at least 3 times, for a complete removal of debris. Nuclear extracts were put into culture dishes and stained with nucleus targeting dyes (Hoechst 33258 and PI) for confirmation of the nucleus. The extracts exhibited nuclear swelling and co-localization (pink color) of the Hoechst 33258 (blue color) and PI (red color) dyes (Fig. 3C). The nuclei of DOX-treated cells were round in shape with a distinct nuclear boundary, while the nuclei of ETO-treated cells were an irregular shape, and DNA leakage from the nuclei was observed. Nepoptosis induced NE rupturing and DNA leakage from the nuclei. From our morphological analyses during necrosis and nepoptosis, we identified changes in nucleus morphology and NE topography, using CNT/AFM probes at a nanometer scale, in DOX-and ETO-treated cells, at 72 h post-treatment. Based on the analysis of AFM images, we determined that the nucleus area of HK-2 and DMSO-treated cells was 140.0 ± 20.3 μ m 2 , and the volume was 18.7 ± 2.9 μ m 3 for 10 evaluated nuclei. DOX-treatment induced nucleus swelling, with a mean nucleus area of 492.1 ± 54.5 μ m 2 and volume of 87.6 ± 18.0 μ m 3 , and the cells typically exhibited nuclei with a round shape and a distinct nuclear boundary. The nuclei of ETO-treated cells were also shown to undergo nuclear swelling, with a nucleus area of 436.1 ± 67.0 μ m 2 and a volume of 88.8 ± 20.7 μ m 3 , and nuclei has a visibly irregular shape with indistinct nuclear boundaries (Fig. 4A,C,D and Supplementary Fig. S5). ETO-treatment caused significant NE rupturing, with the size of ruptures showing a mean height of 94.2 ± 28.1 nm and width of 644.3 ± 189.0 nm, for 20 evaluated ruptures. A large number of linked-fiber chromatin was released from nuclei, and each single chromatin exhibited a diameter of 70-100 nm (Fig. 4B). Additionally, the NE changes in the ETO-treated HK-2 cells were analyzed after 24 h, using CNT/AFM probes. Interestingly, the results showed characteristic features of early apoptotic cell death, including NE ruptures and DNA release (Supplementary Fig. S6). However, the changes observed for 24 h ETO-treated cells were less significant than those reported in cells treated for 72 h. Therefore, in the early apoptotic cell death stage, small NE ruptures and release of small amounts of DNA were reported, while in late apoptotic cell death stages, severe NE ruptures as well as an excessive release of DNAs were observed.

Cell adhesion disruption and NE rupturing are induced by caspase during nepoptosis.
Nepoptotic cell death generated cell adhesion disruption and NE rupturing, while necrotic cell death did not have any effect on cell adhesion or NE topography. Based on these results, we predicted, from a number of previous studies, that these outcomes were likely attributable to caspase activation during apoptosis 15,25 . Moreover, we demonstrated that caspases regulate changes in cell adhesion and NE topography using the pharmacological pan-caspase inhibitor, z-VAD, which almost completely inhibited ETO-induced caspase-3/7 activity (83.3% inhibition) at 72 h post-treatment (Fig. 5A). In these conditions, we demonstrated an alteration in cell adhesion using the xCELLigence system, and showed that inhibition of pan-caspase increased the CI from 2.2 ± 0.1-fold to 3.2 ± 0.3-fold and accelerated the ST from 175 min to 160 min when compared to ETO-treatment (Fig. 5B,C). In addition, we determined that cell nuclei of HK-2 and z-VAD-treated cells were round in shape with a distinct boundary, and also exhibited a large number of NPCs which were observed by using CNT/AFM probes ( Fig. 6A and Supplementary Fig. S7B). Other studies showed similar results to ours 34,35 . The nuclei of ETO-treated cells clearly exhibited NE ruptures and DNA leakages, which were significantly suppressed by the inhibition of pan-caspase ( Fig. 6A and Supplementary Fig. S7A). However, the inhibition of pan-caspase did not significantly alter nucleus area or volume of ETO-treated cells, based on 10 evaluated nuclei ( Supplementary Fig. S7C).

ENDOG is translocated in the nucleus through caspase-induced NE rupturing during nepoptosis.
Translocation of endonuclease G (ENDOG) to the nucleus has been shown to trigger DNA fragmentation in the caspase-independent apoptosis pathway 36 ; however, the mechanism for translocation is still unknown. Based on our results, we hypothesized that ENDOG translocation is mediated by caspase-induced NE rupturing, during ETO-induced nepoptosis. Thus, we measured translocation of ENDOG levels in pan-caspase inhibited cells. We confirmed the translocation of ENDOG (green color) to the nucleus of ETO-treated cells (indicated by the arrow) through disruption of NE using a nucleus-targeting dye (Hoechst 33258, blue color) and confocal microscopy (Fig. 6B, Supplementary  Fig. S8A,B). In the pan-caspase-inhibited nuclei, ENDOG translocation and NE disruption were not apparent, as there were no significant changes in nucleus size from that of the uninhibited cells ( Fig. 6B and Supplementary Fig. S8C). Similarly, results from the Cellomics ArrayScan HCS reader analysis showed that ETO-treatment increased ENDOG translocation in nuclei, by 3.1-fold from the control levels. Inhibition of pan-caspase resulted in a decrease in ENDOG translocation (33.7%), from levels of translocation for the ETO control cells, but did not significantly change the nucleus area-measured for at least 200 cells per well ( Fig. 6C and Supplementary Fig. S7C). Furthermore, ETO treatment induced the expression of cleaved-caspase-3 and cleaved-lamin A/C (NE protein). However, the expression of these proteins and the translocation of ENDOG from cytosol to nucleus were significantly suppressed by the inhibition of pan-caspase (Fig. 6D,E).

Discussion
The main purpose of the present study was to characterize cell adhesion, using xCELLigence analysis and NE topography obtained by AFM analysis, in necrotic and apoptotic cell death (Fig. 6F).
Cell adhesion plays an important role in cell migration, growth, differentiation, and morphology 9 , thus, it is associated with various human diseases, such as ischemia, asthma, diabetes, bacterial infections, and others 37,38 . In particular, the alteration of cell adhesion molecules has been shown to modulate invasion, metastasis and morphology in multistage carcinogenesis 39 . Currently, various cell adhesion-targeting drugs are being tested in clinical trials, in order to prevent the occurrence of carcinogenesis [40][41][42] . Amongst them, VS-6063 (formerly PF-04554878), has been reported to increase the Scientific RepoRts | 5:15623 | DOi: 10.1038/srep15623 apoptosis induced by paclitaxel, in ovarian cancer cells resistant to taxane, through the inhibition of the FAK-mediated chemoresistance signaling pathway 43 . This favored the application of these drugs in multiple clinical trials, such as trials for solid cancers, mesothelioma cancer, and ovarian cancer. In studies addressing cell death, it has been reported that apoptosis involved the disruption of key proteins for focal adhesion, including FAK and CAS, and of cytoskeletal proteins, including α -tubulin and phalloidin important for cell behavior and morphology, by the activation of caspases 13,16 . Generally, cell adhesion ability is measured by fibronectin and extracellular matrix protein-coated fluorometric or colorimetric detection assay kits, or by the enzyme-linked immunosorbent assay (ELISA), in studies of cancer and cell death [44][45][46] . However, these methods are limited by the indirect detection of cell adhesion ability and their detection only of the end-points. Here, we suggested the use of the xCELLigence system to directly measure and characterize the changes in cell adhesion during apoptosis and necrosis, in real-time. The xCELLigence system is used for cell-based measurements of cell viability, invasion, and migration, in real-time, which could not be detected by fluorometric or colorimetric methods 31,32 . Recently, some researchers used the xCELLigence system to study the migration of venous endothelial cells in chronic venous disease patients, or to study human neuronal cell networks 47,48 . In this study, we demonstrated, using xCELLigence system, that necrotic cell death generated cell swelling, while apoptotic cell death rapidly triggered loss of cell adhesion, and a decrease in cell-adhesion speed. The inhibition of pan-caspases limited destruction of the cell adhesion area during apoptosis to a level similar to that usually observed in necrosis. Overall, we determined that cell adhesion could be characterized in apoptosis and necrosis, based on the following properties: apoptosis induces loss of cell adhesion, through caspase involvement, but necrosis does not have any significant effect on cell adhesion. The calculation of a cell adhesion score presents a key indicator for both necrosis and apoptosis. During necrosis, an increase in the cell adhesion score is observed, while this score decreases during apoptosis. The xCELLigence method presented here, might represent a powerful tool for the measurement of cell adhesion, as it provides information regarding cell adhesion ability during necrosis and apoptosis, which serves to better characterize cell death. This method might also be a valuable aid to studies that address cell behavior in a variety of human diseases. In addition, the novel measurement ability realized by this method, and its concepts, will help in the design and production of new biosensors.
The NE separates the genetic information within a cell, from the cytoplasm, and controls the bi-directional transport of diverse molecules between the nucleus and cytoplasm. Moreover, changes in the NE have been reported to be associated with the development of multistage disease and carcinogenesis 49,50 . For example, both basal and squamous carcinoma cells present increased lamin A/C protein expression levels 51 . The NPC, composed of NUP88, has been found to be strongly expressed in multiple tumor cells, including adenocarcinoma cells, cervical carcinoma cells, and breast cancer cells 52 . In a study of stem cells, mouse and human ESCs both expressed B-type lamin proteins, but did not express A/C-type lamin proteins. However, lamin A/C proteins have been found to be expressed during the differentiation of both mouse and human ESCs 53 . Moreover, lamin B proteins are required for proper organogenesis, but do not have any effect on mouse ESCs 54 . Structural analysis of ESCs showed the presence of irregular and wide-shaped NEs in the intermembrane space of ESCs, in comparison to (A) Cells treated with ETO and cells co-treated with z-VAD were harvested. Nuclear extracts were seeded on culture dishes for 15 min and the nuclei were fixed. We measured nuclear envelope topography using a CNT/AFM probes system. Images shown are representative of 3D topography at 30-, 10-, and 2-μ m scales. See also Supplementary Fig. S7. (B) Cells treated with ETO and co-treated with z-VAD regulated endonuclease G (green color) translocation levels, indicated by immunofluorescence staining and Hoechst 33258 (blue color) that was used for nuclear staining and were measured using confocal microscopy (scale bar represents 10 μ m) at 72 h. Arrow indicates nuclear envelope ruptures (collapse Hoechst 33258 intensity) and/or endonuclease G translocation which also is also shown in Supplementary Fig. S8. (C) Translocation endonuclease G levels stained by immunofluorescence and Hoechst 33258 for nuclei that were measured using the Cellomics ArrayScan HCS Reader for at least 200 cells. Endonuclease G intensity was dependent on Hoechst 33258-positive area that is shown in the histogram compared to the control (statistical analysis P-value of *P < 0.05, **P < 0.01). (D) Cells treated with ETO and co-treated with z-VAD regulated expression of cleaved-caspase 3 and cleaved-lamin A/C protein in whole-extracted proteins evaluated using western blot analysis with β -actin as the loading control. (E) Cells treated with ETO and cotreated with z-VAD regulated expression of endonuclease G in the cytoplasm and nuclei as determined by western blot analysis with β -actin and HDAC1 as cytoplasm and nucleus loading controls, respectively. (F) The summarized cell adhesion, nuclear envelope shape, and nuclear envelope topography observed during necrosis and apoptosis.
Scientific RepoRts | 5:15623 | DOi: 10.1038/srep15623 differentiated cells, through use of fluorescence staining and electron microscopy 55 . Further, apoptotic cell death showed disassembling of NE proteins, and structural changes in nucleus, such as shrinkage, NE rupture, irregular nucleus shape, and DNA leakage, which were measured by fluorescence staining and electron microscopy 29,56,57 . However, the study of structural and topographical changes of the NE using morphometric analysis is not ideal, due to the indirect detection of change by current methods, and the detection of only partial changes. Furthermore, necrosis-regulated NE alteration is not fully understood, and the methods for measuring these changes are limited. Here, we used a direct method of measurement, and determined that the nuclei of cells undergoing necrotic cell death exhibited a distinct rounded shape and clear nuclear boundary; NPCs were present in the NE, with a NE topography similar to that usually observed in normal nuclei. On the other hand, the activation of caspase triggered the formation of irregular-shaped nuclei and the disappearance of NPCs, in addition to the formation of significant ruptures and swelling in the NE, during apoptotic cell death. Therefore, this measurement method using CNT/AFM probes is a powerful tool, as it provides information regarding topographical alterations of the plasma membrane and the NE during necrosis and apoptosis, leading to better characterization of cell death features. Moreover, this method can be used for structural and functional analyses of the NE in studies of stem cells, and other cells in human multistage diseases.
In the mitochondrial or intrinsic signaling pathways, during apoptosis, the expression of BCL-2-associated X protein (BAX) or the BCL-2 antagonist/killer protein (BAK) cause the permeabilization of the mitochondrial outer membrane, and then production of apoptosis-inducing proteins, including cytochrome c (CYTC), caspase-activated DNase (CAD), apoptosis inducing factor (AIF), and ENDOG, which are released from the mitochondria to the cytoplasm 4,58 . CYTC forms an apoptosome by complex formation with apoptotic protease activating factor 1 (APAF1), and brings about DNA fragmentation and diverse protein destruction, through the activation of caspases [58][59][60] . DNA fragmentation is induced by nucleases, according to one of two mechanisms: caspase-dependent activation of CAD or caspase-independent release of AIF and ENDOG proteins from the mitochondria 1,4,61 . CAD released from the mitochondria is translocated into the nucleus, leading to DNA fragmentation, through the activation of caspase-3 in nucleus 62 . On the other hand, AIF and ENDOG are caspases that independently induce DNA fragmentation by translocating from the nucleus to the cytoplasm during apoptosis 36,63 . However, the mechanism for caspase and nuclease translocation in the nucleus is fully understood. In our study, we showed that apoptotic cell death was induced by ENDOG translocation and the leakage of DNA in the nucleus, through caspase-triggered NE ruptures. We also showed that caspases induced NE ruptures, and ultimately caused fragmentation of DNA and nuclear proteins, and leakage of DNA, in early-stage apoptosis. Translocation of caspases induced intermediate filament protein disassembly, triggering collapse of the NPCs and other proteins, by causing membrane infirmness in the NE. The progression of serious intermediate filament damage was triggered by pyknosis and/or karyorrhexis, through the collapse of structures due to external pressures during late-stage apoptosis.
The swelling of ETO-treated cells gradually caused apoptotic morphological changes, including cell shrinkage and apoptotic bodies (Supplementary Fig. S1 and Movie S1). On the other hand, necrotic cells stopped proliferating and maintained cell swelling when they were re-seeded in fresh media, but apoptosis continued to induce cell death and cell shrinkage. Hence, in DNA damage-induced cell death progressing from apoptosis to necrosis, as well as apoptosis alone, cells could not recover from the damage caused by destruction of the nucleus with DNA fragmentation. Taken together, our studies provide the structural and functional analyses of cell adhesion and nuclear envelope nano-topography in cell death.
Analysis of cell swelling measured using hemocytometer and FACS systems. HK-2 cells (4 × 10 5 cells/6-cm culture dish) were grown overnight and treated with DOX and ETO for the specified times. After treatment, cells were collected and washed with 3 mL of phosphate buffered saline (PBS) and centrifuged at 200 × g for 5 min. For measurement, the harvested cells were first seeded in a hemocytometer (Paul Marienfeld GmbH & Co., KG, Bad Mergentheim, Germany); cell swelling was measured by phase contrast microscopy (E-scope i304, Macrotech Corporation, Goyang, South Korea) and analyzed using Scopephoto software. Next, forward scattered light (FSC) units indicating cell size were measured using FACS Aria III with Diva software (BD Biosciences., San Diego, CA, USA) for at least 1 × 10 5 cells.
Analysis of cell adhesion alteration using an xCELLigence system for real-time measurements. HK-2 cells (4 × 10 5 cells/6-cm culture dish) were grown overnight and treated at the specified conditions for 72 h. Cells were then collected, washed with PBS (3 mL), and centrifuged at 200 × g for 5 min.