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18 pages, 3984 KiB  
Article
Susceptibility of HPV-18 Cancer Cells to HIV Protease Inhibitors
by Lilian Makgoo, Salerwe Mosebi and Zukile Mbita
Viruses 2024, 16(10), 1622; https://doi.org/10.3390/v16101622 (registering DOI) - 17 Oct 2024
Viewed by 137
Abstract
Cervical cancer cases continue to rise despite all the advanced screening and preventative measures put in place, which include human papillomavirus (HPV) vaccination. These soaring numbers can be attributed to the lack of effective anticancer drugs against cervical cancer; thus, repurposing the human [...] Read more.
Cervical cancer cases continue to rise despite all the advanced screening and preventative measures put in place, which include human papillomavirus (HPV) vaccination. These soaring numbers can be attributed to the lack of effective anticancer drugs against cervical cancer; thus, repurposing the human immunodeficiency virus protease inhibitors is an attractive innovation. Therefore, this work was aimed at evaluating the potential anticancer activities of HIV-PIs against cervical cancer cells. The MTT viability assay was used to evaluate the effect of HIV protease inhibitors on the viability of cervical cancer cells (HeLa) and non-cancerous cells (HEK-293). Further confirmation of the MTT assay was performed by confirming the IC50s of these HIV protease inhibitors on cervical cancer cells and non-cancerous cells using the Muse™ Count and Viability assay. To confirm the mode of death induced by HIV protease inhibitors in the HPV-associated cervical cancer cell line, apoptosis was performed using Annexin V assay. In addition, the Muse™ Cell Cycle assay was used to check whether the HIV protease inhibitors promote or halt cell cycle progression in cervical cancer cells. HIV protease inhibitors did not affect the viability of non-cancerous cells (HEK-293), but they decreased the viability of HeLa cervical cancer cells in a dose-dependent manner. HIV protease inhibitors induced apoptosis in HPV-related cervical cancer cells. Furthermore, they also induced cell cycle arrest, thus halting cell cycle progression. Therefore, the use of HIV drugs, particularly HIV-1 protease inhibitors, as potential cancer therapeutics represents a promising strategy. This is supported by our study demonstrating their anticancer properties, notably in HPV-associated cervical cancer cell line. Full article
(This article belongs to the Special Issue Chronic Infection by Oncogenic Viruses)
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Figure 1

Figure 1
<p>Effect of lopinavir HIV protease inhibitor on the viability of HeLa cells after 24 h treatment (<b>A</b>), 48 h treatment (<b>B</b>), and 72 h treatment (<b>C</b>). Lopinavir significantly (<span class="html-italic">p</span> ≤ 0.01 **/<span class="html-italic">p</span> ≤ 0.001 ***) reduced the viability of HeLa cells in a time and concentration-dependent manner. “ns” indicates a non-significant effect.</p>
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<p>Effect of Aluvia HIV protease inhibitor tablets on the viability of HeLa cells after 24 h treatment (<b>A</b>) and 48 h treatment (<b>B</b>). Aluvia tablets significantly (<span class="html-italic">p</span> ≤ 0.05 */<span class="html-italic">p</span> ≤ 0.001 ***) reduced the viability of HeLa cells in a time and concentration-dependent manner. “ns” indicates a non-significant effect.</p>
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<p>Effect of atazanavir HIV protease inhibitor on the viability of HeLa cells after 24 h treatment (<b>A</b>) and 48 h treatment (<b>B</b>). Atazanavir significantly (<span class="html-italic">p</span> ≤ 0.05 */<span class="html-italic">p</span> ≤ 0.01 **/<span class="html-italic">p</span> ≤ 0.001 ***) reduced the viability of HeLa cells in a time and concentration-dependent manner. “ns” indicates a non-significant effect.</p>
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<p>Effect of Ritoataz HIV protease inhibitor tablets on the viability of HeLa cells after 24 h treatment (<b>A</b>) and 48 h treatment (<b>B</b>). Ritoataz tablets significantly (<span class="html-italic">p</span> ≤ 0.05 */<span class="html-italic">p</span> ≤ 0.01 **/<span class="html-italic">p</span> ≤ 0.001 ***) reduced the viability of HeLa cells in a time and concentration-dependent manner. “ns” indicates a non-significant effect.</p>
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<p>An analysis of the cytotoxicity of pure HIV protease inhibitors and HIV protease inhibitor tablets against HEK-293 human embryonic kidney cells. Statistically, there was no significant difference (ns) between the treated and untreated control groups.</p>
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<p>Confirmation of percentage viability of HeLa cells in response to treatment with lopinavir (<b>A</b>), Aluvia tablets (<b>B</b>), atazanavir (<b>C</b>), and Ritoataz tablets (<b>D</b>). All the HIV drugs significantly (<span class="html-italic">p</span> ≤ 0.001 ***) inhibited the viability of HeLa cells in vitro when compared to the untreated control. “ns” indicates a non-significant effect.</p>
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<p>Confirmation of percentage viability of HEK-293 cells in response to treatment with pure HIV protease inhibitors and HIV protease inhibitor tablets. Compared to the untreated control, the difference was found to be statistically non-significant (ns).</p>
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<p>Average % apoptosis in lopinavir (<b>A</b>), Aluvia (<b>B</b>), atazanavir (<b>C</b>), and Ritoataz (<b>D</b>)-treated HeLa cells. HIV protease inhibitors and curcumin (40 μM) significantly (<span class="html-italic">p</span> ≤ 0.01 **/<span class="html-italic">p</span> ≤ 0.001 ***) induced apoptosis cell death in HeLa cells relative to the untreated control. “ns” indicates a non-significant effect.</p>
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<p>The effect of lopinavir (<b>A</b>), Aluvia (<b>B</b>), atazanavir (<b>C</b>) and Ritoataz (<b>D</b>) on cell cycle progression. The solvent controls had no significant effect (ns) on the progression of the cell cycle in HeLa cells when compared to the untreated control. All the HIV protease inhibitors significantly (<span class="html-italic">p</span> ≤ 0.01 <sup>##</sup>/<span class="html-italic">p</span> ≤ 0.001 <sup>###</sup>) halted the cell cycle progression at the G0/G1 phase.</p>
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<p>The effect of lopinavir (<b>A</b>), Aluvia (<b>B</b>), atazanavir (<b>C</b>) and Ritoataz (<b>D</b>) on cell cycle progression. The solvent controls had no significant effect (ns) on the progression of the cell cycle in HeLa cells when compared to the untreated control. All the HIV protease inhibitors significantly (<span class="html-italic">p</span> ≤ 0.01 <sup>##</sup>/<span class="html-italic">p</span> ≤ 0.001 <sup>###</sup>) halted the cell cycle progression at the G0/G1 phase.</p>
Full article ">Figure A1
<p>Effect of curcumin on the viability of HeLa cells after 24 h treatment. Curcumin significantly (<span class="html-italic">p</span> ≤ 0.01 **/<span class="html-italic">p</span> ≤ 0.001 ***) reduced the viability of HeLa cells in a time and concentration-dependent manner. “ns” indicates a non-significant effect.</p>
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17 pages, 1049 KiB  
Article
Synthesis of Tetrahydrocarbazole-Tethered Triazoles as Compounds Targeting Telomerase in Human Breast Cancer Cells
by Pradeep M. Uppar, Akshay Ravish, Zhang Xi, Keshav Kumar Harish, Arun M. Kumar, Lisha K. Poonacha, Toreshettahally R. Swaroop, Chaithanya Somu, Santosh L. Gaonkar, Mahendra Madegowda, Peter E. Lobie, Vijay Pandey and Basappa Basappa
Catalysts 2024, 14(10), 726; https://doi.org/10.3390/catal14100726 (registering DOI) - 16 Oct 2024
Viewed by 281
Abstract
Telomere shortening and the induction of senescence and/or cell death may result from inhibition of telomerase activity in cancer cells. Herein, the properties of carbazole–triazole compounds targeting telomerase in human breast cancer cells are explored. All derivatives were evaluated for loss of viability [...] Read more.
Telomere shortening and the induction of senescence and/or cell death may result from inhibition of telomerase activity in cancer cells. Herein, the properties of carbazole–triazole compounds targeting telomerase in human breast cancer cells are explored. All derivatives were evaluated for loss of viability in MCF-7 breast cancer cells, with compound 5g identified as the most potent within the examined series. Green synthesis was employed using water, a reusable nano-Fe2O3-catalyzed reaction, and an electrochemical method for the synthesis of tetrahydrocarbazole and triazoles. The crystal data of compound 4 is also reported. Furthermore, in silico analysis predicted that compound 5g may target human telomerase. Molecular docking analysis of compound 5g towards hTERT predicted a binding affinity of −6.74 kcal/mol. In flow cytometry assays, compound 5g promoted apoptosis and cell cycle arrest in the G2-M phase. Finally, compound 5g inhibited the enzymatic activity of telomerase in human breast cancer cells. In conclusion, a green synthesized series of carbazole–triazoles that target telomerase in cancer cells is reported. Full article
(This article belongs to the Section Catalysis in Organic and Polymer Chemistry)
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14 pages, 8177 KiB  
Article
The Protective Role of Intermedin in Contrast-Induced Acute Kidney Injury: Enhancing Peritubular Capillary Endothelial Cell Adhesion and Integrity Through the cAMP/Rac1 Pathway
by Tingting Gao, Ruiyuan Gu, Heng Wang, Lizheng Li, Bojin Zhang, Jie Hu, Qinqin Tian, Runze Chang, Ruijing Zhang, Guoping Zheng and Honglin Dong
Int. J. Mol. Sci. 2024, 25(20), 11110; https://doi.org/10.3390/ijms252011110 - 16 Oct 2024
Viewed by 239
Abstract
Contrast-induced acute kidney injury (CIAKI) is a common complication with limited treatments. Intermedin (IMD), a peptide belonging to the calcitonin gene-related peptide family, promotes vasodilation and endothelial stability, but its role in mitigating CIAKI remains unexplored. This study investigates the protective effects of [...] Read more.
Contrast-induced acute kidney injury (CIAKI) is a common complication with limited treatments. Intermedin (IMD), a peptide belonging to the calcitonin gene-related peptide family, promotes vasodilation and endothelial stability, but its role in mitigating CIAKI remains unexplored. This study investigates the protective effects of IMD in CIAKI, focusing on its mechanisms, particularly the cAMP/Rac1 signaling pathway. Human umbilical vein endothelial cells (HUVECs) were treated with iohexol to simulate kidney injury in vitro. The protective effects of IMD were assessed using CCK8 assay, flow cytometry, ELISA, and Western blotting. A CIAKI rat model was utilized to evaluate renal peritubular capillary endothelial cell injury and renal function through histopathology, immunohistochemistry, immunofluorescence, Western blotting, and transmission electron microscopy. In vitro, IMD significantly enhanced HUVEC viability and mitigated iohexol-induced toxicity by preserving intercellular adhesion junctions and activating the cAMP/Rac1 pathway, with Rac1 inhibition attenuating these protective effects. In vivo, CIAKI caused severe damage to peritubular capillary endothelial cell junctions, impairing renal function. IMD treatment markedly improved renal function, an effect negated by Rac1 inhibition. IMD protects against renal injury in CIAKI by activating the cAMP/Rac1 pathway, preserving peritubular capillary endothelial integrity and alleviating acute renal injury from contrast media. These findings suggest that IMD has therapeutic potential in CIAKI and highlight the cAMP/Rac1 pathway as a promising target for preventing contrast-induced acute kidney injury in at-risk patients, ultimately improving clinical outcomes. Full article
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Figure 1

Figure 1
<p>IMD can antagonize damage to HUVEC viability and apoptosis induced by iohexol. (<b>A</b>) HUVECs were preincubated with IMD (0, 1, 10, 100 nmol/L) for 30 min and then treated with iohexol (10, 20, 40, 80 mgI/mL) for 12 h. A CCK-8 kit was used to test cell viability. (<b>B</b>) HUVECs were preincubated with IMD (10 nmol/L) for 30 min, then were treated with iohexol (40 mgI/mL) or iohexol (40 mgI/mL) +NSC23766(50 μM) for 12 h. Apoptosis was detected using flow cytometry. (<b>C</b>) The apoptosis rate in each group was quantified (<span class="html-italic">n</span> = 6). The results were analyzed using ANOVA, followed by Tukey’s multiple comparisons test for subgroup analysis. All data are expressed as mean ± SD, * <span class="html-italic">p</span> &lt; 0.05 compared to the control group, # <span class="html-italic">p</span> &lt; 0.05. For comparison between groups, ns: not significant.</p>
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<p>IMD can protect the adherens junction of HUVECs by activating the cAMP/Rac1 pathway. (<b>A</b>) Intermedin induces cAMP production in HUVECs. The concentration of cAMP was measured using ELISA, <span class="html-italic">n</span> = 4. (<b>B</b>,<b>C</b>) HUVECs were treated with 10 nmol of IMD and/or 40 mgI/mL iohexol or 50 μM NSC23766 for 12 h. Whole-cell lysates of HUVECs were collected for immunoblotting analysis of Rac1, VE-cadherin, and GAPDH. The results were analyzed using ANOVA, with Tukey’s multiple comparisons test applied for subgroup analysis. All data are expressed as mean ± SD, * <span class="html-italic">p</span> &lt; 0.05 compared to the control group, <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05. For comparison between groups, ns: not significant.</p>
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<p>IMD attenuates renal injury in rat CIAKI models, and inhibition of Rac1 might abolish this protective effect. (<b>A</b>) Male SD rats were randomly divided into five groups (<span class="html-italic">n</span> = 6), and the rats in each group were treated as shown in the scheme. (<b>B</b>) Renal morphology (HE, magnification ×400) showed vacuolar degeneration of renal tubules (shown by red arrows) and dilatation of renal tubules (shown by green arrows). (<b>C</b>) Magnification of renal PAS staining (×400) showed the absence of brush edges of renal tubules, vacuolar degeneration, and dilatation of renal tubules. (<b>D</b>) Renal tubular injury score. (<b>E</b>) Serum creatinine levels for the indicated treatments. The results were analyzed using ANOVA, with Tukey’s multiple comparisons test applied for subgroup analysis. All data are expressed as mean ± SD, * <span class="html-italic">p</span> &lt; 0.05 compared to the control group, # <span class="html-italic">p</span> &lt; 0.05. For comparison between groups, ns: not significant.</p>
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<p>IMD activates the cAMP/Rac1 pathway and alleviates renal peritubular capillary injury in CIAKI rats. (<b>A</b>,<b>B</b>): The renal lysates of rats were collected and the expression of Rac1, VE-cadherin, VEGFR2, and GAPDH were detected using Western blotting. (<b>C</b>) Kidney sections were stained with CD34 immunohistochemical staining (magnification, ×400, scale 30 μm). (<b>D</b>) Kidney sections were stained with CD34 and ICAM1 immunofluorescence staining (magnification, ×200, scale 50 μm). (<b>E</b>) The average optical density of CD34 positive staining in each group was quantified (<span class="html-italic">n</span> = 6). (<b>E</b>) Quantification of the average optical density of ICAM1 immunofluorescence (<span class="html-italic">n</span> = 6). The data were analyzed using ANOVA, followed by Tukey’s multiple comparisons test for subgroup analysis. All data are expressed as mean ± SD, * <span class="html-italic">p</span> &lt; 0.05 compared to the control group, # <span class="html-italic">p</span> &lt; 0.05. For comparison between groups, ns: not significant.</p>
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<p>IMD protects the PTC endothelial barrier: a representative microphotograph of the ultrastructural changes of peritubular capillary endothelial cells, cell membrane discontinuity (shown by green arrows), and basement membrane fracture (shown by blue arrows) (original magnification, ×12,000; scale, 1 μm).</p>
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20 pages, 6974 KiB  
Article
Targeting Ferroptosis with Small Molecule Atranorin (ATR) as a Novel Therapeutic Strategy and Providing New Insight into the Treatment of Breast Cancer
by Mine Ensoy and Demet Cansaran-Duman
Pharmaceuticals 2024, 17(10), 1380; https://doi.org/10.3390/ph17101380 - 16 Oct 2024
Viewed by 253
Abstract
Background/Objectives: Ferroptosis results from the accumulation of iron-dependent lipid peroxides and reactive oxygen species (ROS). Previous research has determined the effect of atranorin (ATR) on other cell death mechanisms, but its potential for a ferroptotic effect depending on ROS levels is unclear. This [...] Read more.
Background/Objectives: Ferroptosis results from the accumulation of iron-dependent lipid peroxides and reactive oxygen species (ROS). Previous research has determined the effect of atranorin (ATR) on other cell death mechanisms, but its potential for a ferroptotic effect depending on ROS levels is unclear. This study details the therapeutic role of small-molecule ATR through ferroptosis by suppressing MDA-MB-231, MCF-7, BT-474, and SK-BR-3 breast cancer cells. Methods: The anti-proliferative effect of ATR on cells was evaluated by xCELLigence analysis, and ferroptotic activity was evaluated by enzymatic assay kits. The changes in gene and protein expression levels of ATR were investigated by the qRT-PCR and western blot. In addition, mitochondrial changes were examined by transmission electron microscopy. Results: ATR was found to reduce cell viability in cancer cells in a dose- and time-dependent manner without showing cytotoxic effects on normal breast cells. In BT-474 and MDA-MB-231 cells, ATR, which had a higher anti-proliferative effect, increased iron, lipid peroxidation, and ROS levels in cells and decreased the T-GSH/GSSG ratio. The results revealed for the first time that small-molecule ATR exhibited anti-cancer activity by inducing the glutathione pathway and ferroptosis. Conclusions: This study highlights the potential of ATR as a drug candidate molecule that can be used in the development of new therapeutic strategies for the treatment of triple-negative and luminal-B breast cancer subtypes. Full article
(This article belongs to the Section Biopharmaceuticals)
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Graphical abstract

Graphical abstract
Full article ">Figure 1
<p>Dose- and time-dependent anti-proliferative effects of ATR on (<b>A</b>) MCF-12A, (<b>B</b>) BT-474, (<b>C</b>) MDA-MB-231, (<b>D</b>) MCF-7, and (<b>E</b>) SK-BR-3 cells using MTT assays; (<b>F</b>) cell viability (%) after 48 h in cells treated with different concentrations of ATR compared to control. Data are represented as the mean ± SD error of three biological replicates. * <span class="html-italic">p</span> &lt; 0.05; ** <span class="html-italic">p</span> &lt; 0.01; *** <span class="html-italic">p</span> &lt; 0.001 (compared to control).</p>
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<p>Dose- and time-dependent anti-proliferative effects of ATR on (<b>A</b>) BT-474, (<b>B</b>) MDA-MB-231, and (<b>C</b>) MCF-12A cells using the xCELLigence real-time cell analyzer. (** <span class="html-italic">p</span> &lt; 0.01; *** <span class="html-italic">p</span> &lt; 0.001).</p>
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<p>Dose- and time-dependent anti-proliferative effects of (<b>A</b>) erastin and (<b>B</b>) ferrostatin-1 on BT-474 cells and (<b>C</b>) erastin and (<b>D</b>) ferrostatin-1 on MDA-MB-231 cells using the xCELLigence real-time cell analyzer (** <span class="html-italic">p</span> &lt; 0.01).</p>
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<p>The effect of different combinations of ATR, erastin, and ferrostatin-1 molecules on cell viability and proliferation in (<b>A</b>) BT-474 and (<b>B</b>) MDA-MB-231 cell lines (*** <span class="html-italic">p</span> &lt; 0.001).</p>
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<p>(<b>A</b>) Iron ion level, (<b>B</b>) T-GSH/GSSG ratio, (<b>C</b>) MDA level, (<b>D</b>) ROS level in BT-474 cells, and (<b>E</b>) iron ion level, (<b>F</b>) T-GSH/GSSG ratio, (<b>G</b>) MDA level, (<b>H</b>) ROS level in MDA-MB-231 cells with and without ATR (IC<sub>50</sub> concentration); ferroptosis-related gene expression levels in (<b>I</b>) BT-474 and (<b>J</b>) MDA-MB-231 cells treated and untreated with ATR. <span class="html-italic">Gapdh</span> was used as the housekeeping gene. Data are represented as the mean ± SD error of three biological replicates; (<b>K</b>,<b>L</b>) ferroptosis-related protein levels in BT-474 and MDA-MB-231 cells with and without ATR. Results are normalized to β-Actin. The dash lines are used to show the change according to the control. (* <span class="html-italic">p</span> &lt; 0.05; ** <span class="html-italic">p</span> &lt; 0.01; *** <span class="html-italic">p</span> &lt; 0.001; **** <span class="html-italic">p</span> &lt; 0.0001).</p>
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<p>Images of mitochondrial morphology under transmission electron microscopy in BT-474 cells of treated and non-treated ATR. Images were obtained with a magnification scale of 8000× and 15,000×. The red arrows indicate mitochondria.</p>
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<p>Images of mitochondrial morphology under transmission electron microscopy in MDA-MB-231 cells treated and non-treated ATR. Images were obtained with a magnification scale of 8000× and 15,000×. The red arrows indicate mitochondria.</p>
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14 pages, 2928 KiB  
Article
PEP-1–PIN1 Promotes Hippocampal Neuronal Cell Survival by Inhibiting Cellular ROS and MAPK Phosphorylation
by Jung Hwan Park, Min Jea Shin, Gi Soo Youn, Hyeon Ji Yeo, Eun Ji Yeo, Hyun Jung Kwon, Lee Re Lee, Na Yeon Kim, Su Yeon Kwon, Su Min Kim, Yong-Jun Cho, Sung Ho Lee, Hyo Young Jung, Dae Won Kim, Won Sik Eum and Soo Young Choi
Biomedicines 2024, 12(10), 2352; https://doi.org/10.3390/biomedicines12102352 (registering DOI) - 15 Oct 2024
Viewed by 435
Abstract
Background: The peptidyl-prolyl isomerase (PIN1) plays a vital role in cellular processes, including intracellular signaling and apoptosis. While oxidative stress is considered one of the primary mechanisms of pathogenesis in brain ischemic injury, the precise function of PIN1 in this disease remains [...] Read more.
Background: The peptidyl-prolyl isomerase (PIN1) plays a vital role in cellular processes, including intracellular signaling and apoptosis. While oxidative stress is considered one of the primary mechanisms of pathogenesis in brain ischemic injury, the precise function of PIN1 in this disease remains to be elucidated. Objective: We constructed a cell-permeable PEP-1–PIN1 fusion protein and investigated PIN1’s function in HT-22 hippocampal cells as well as in a brain ischemic injury gerbil model. Methods: Transduction of PEP-1–PIN1 into HT-22 cells and signaling pathways were determined by Western blot analysis. Intracellular reactive oxygen species (ROS) production and DNA damage was confirmed by DCF-DA and TUNEL staining. Cell viability was determined by MTT assay. Protective effects of PEP-1-PIN1 against ischemic injury were examined using immunohistochemistry. Results: PEP-1–PIN1, when transduced into HT-22 hippocampal cells, inhibited cell death in H2O2-treated cells and markedly reduced DNA fragmentation and ROS production. This fusion protein also reduced phosphorylation of mitogen-activated protein kinase (MAPK) and modulated expression levels of apoptosis-signaling proteins in HT-22 cells. Furthermore, PEP-1–PIN1 was distributed in gerbil hippocampus neuronal cells after passing through the blood–brain barrier (BBB) and significantly protected against neuronal cell death and also decreased activation of microglia and astrocytes in an ischemic injury gerbil model. Conclusions: These results indicate that PEP-1–PIN1 can inhibit ischemic brain injury by reducing cellular ROS levels and regulating MAPK and apoptosis-signaling pathways, suggesting that PIN1 plays a protective role in H2O2-treated HT-22 cells and ischemic injury gerbil model. Full article
(This article belongs to the Section Cell Biology and Pathology)
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Figure 1

Figure 1
<p>Construction of PEP-1–PIN1 and control PIN1 protein. Constructed map of PEP-1–PIN1 based on the pET-15b vector. PEP-1–PIN1 was designed to contain histidine, PEP-1-PTD and PIN1 (<b>A</b>). Purified PEP-1–PIN1 and control PIN1 were confirmed by Coomassie brilliant blue staining and Western blot analysis using anti-histidine antibody (<b>B</b>).</p>
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<p>Delivery of PEP-1–PIN1 into HT-22 cells. HT-22 cells were treated with PEP-1–PIN1 (0.5–5 μM) for 3 h (<b>A</b>) or PEP-1–PIN1 (5 μM) for different time periods (30–180 min) (<b>B</b>). The intracellular stability of delivered PEP-1–PIN1 into the cells. HT-22 cells were treated with PEP-1–PIN1 for 3 h and washed. The cells were then further incubated for 1 to 60 h (<b>C</b>) and delivered PEP-1–PIN1 was assessed by Western blotting. The intensity of the bands was measured by a densitometer. Data are represented as mean ± SEM (n = 3).</p>
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<p>Effects of delivered PEP-1–PIN1 against H<sub>2</sub>O<sub>2</sub>-induced cell death. HT-22 cells were treated with PEP-1–PIN1 (5 μM) for 3 h. The localization of delivered PEP-1–PIN1 was confirmed by fluorescence microscopy (<b>A</b>). Scale bar = 20 μm. Effect of delivered PEP-1–PIN1 against H<sub>2</sub>O<sub>2</sub>-induced cell viability. The cells were pretreated with PEP-1–PIN1 (0.5–5 μM) for 3 h and exposed to H<sub>2</sub>O<sub>2</sub> (1 mM) for 2 h. Cell viability was assessed by MTT assay (<b>B</b>). Data are represented as mean ± SEM (n = 3). * <span class="html-italic">p</span> &lt; 0.05 and ** <span class="html-italic">p</span> &lt; 0.01 compared with H<sub>2</sub>O<sub>2</sub>-treated cells.</p>
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<p>Effects of delivered PEP-1–PIN1 against H<sub>2</sub>O<sub>2</sub>-induced ROS production and DNA damage. HT-22 cells were treated with PEP-1–PIN1 (5 μM) for 3 h before treatment with 1 mM H<sub>2</sub>O<sub>2</sub> for 1 h or 6 h. Intracellular ROS levels (<b>A</b>) and DNA damage (<b>B</b>) were determined by DCF-DA and TUNEL staining. Fluorescence intensity was quantified using an ELISA plate reader. Scale bar = 50 μm. Data are represented as mean ± SEM (n = 3). ** <span class="html-italic">p</span> &lt; 0.01 compared with H<sub>2</sub>O<sub>2</sub>-treated cells.</p>
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<p>Effects of delivered PEP-1–PIN1 against H<sub>2</sub>O<sub>2</sub>-induced MAPK and NF-κB expression in HT-22 cells. The cells were treated with PEP-1–PIN1 (5 μM) for 3 h before being exposed to H<sub>2</sub>O<sub>2</sub> (1 mM) for 60 min or 30 min, respectively. The expression levels of MAPKs (<b>A</b>) and NF-κB (<b>B</b>) were analyzed by Western blotting. The intensity of the bands was measured by a densitometer. Data are represented as mean ± SEM (n = 3). * <span class="html-italic">p</span> &lt; 0.05 and ** <span class="html-italic">p</span> &lt; 0.01 compared with H<sub>2</sub>O<sub>2</sub> treated cells.</p>
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<p>Effects of delivered PEP-1–PIN1 against H<sub>2</sub>O<sub>2</sub>-induced Bax, Bcl-2, and p53 protein expression in HT-22 cells. Three-hour pretreatment of HT-22 cells with PEP-1–PIN1 (5 μM) was followed by treatments with H<sub>2</sub>O<sub>2</sub> (1 mM) for 120 min (Bcl-2), 240 min (Bax), and 360 min (p53). The expression levels of Bcl-2 and Bax (<b>A</b>) and p53 (<b>B</b>) were determined by Western blot analysis. The intensity of the bands was measured by a densitometer. Data are represented as mean ± SEM (n = 3). * <span class="html-italic">p</span> &lt; 0.05 and ** <span class="html-italic">p</span> &lt; 0.01 compared with H<sub>2</sub>O<sub>2</sub> treated cells.</p>
Full article ">Figure 7
<p>The neuroprotective effects of delivered PEP-1–PIN1 against ischemic damage. Gerbils were treated with a single injection of PEP-1–PIN1 (2 mg/kg) and sacrificed after 7 days. Delivery of PEP-1–PIN1 into the CA1 region of the hippocampus was determined by anti-histidine immunohistochemistry. Scale bar = 400 μm. The hippocampus was stained with NeuN, CV, GFAP, Iba-1 and FJB in sham-, vehicle-, PEP-1–PIN1-, PIN1-, and PEP-1-treated gerbils after ischemic injury. The graphic shows the relative numerical analyses of CV, GFAP, Iba-1 and FJB positive neurons in the CA1 region. Scale bar = 400 and 50 μm. Each bar represents the mean ± SEM of ten mice. ** <span class="html-italic">p</span> &lt; 0.01, significant difference from the vehicle group.</p>
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18 pages, 3455 KiB  
Article
Isolation and Characterization of Canine Adipose-Derived Mesenchymal Stromal Cells: Considerations in Translation from Laboratory to Clinic
by Michael A. Rivera Orsini, Emine Berfu Ozmen, Alyssa Miles, Steven D. Newby, Nora Springer, Darryl Millis and Madhu Dhar
Animals 2024, 14(20), 2974; https://doi.org/10.3390/ani14202974 (registering DOI) - 15 Oct 2024
Viewed by 290
Abstract
In allogeneic MSC implantation, the cells are isolated from a donor different from the recipient. When tested, allogeneic MSCs have several advantages over autologous ones: faster cell growth, sufficient cell concentration, and readily available cells for clinics. To ensure the safe and efficient [...] Read more.
In allogeneic MSC implantation, the cells are isolated from a donor different from the recipient. When tested, allogeneic MSCs have several advantages over autologous ones: faster cell growth, sufficient cell concentration, and readily available cells for clinics. To ensure the safe and efficient use of allogeneic MSCs in clinics, the MSCs need to be first tested in vitro. With this study, we paved the way by addressing the in vitro aspects of canine adipose-derived MSCs, considering the limited studies on the clinical use of canine cells. We isolated cAD-MSCs from canine falciform ligament fat and evaluated their viability and proliferation using an MTS assay. Then, we characterized the MSC-specific antigens using immunophenotyping and immunofluorescence and demonstrated their potential for in vitro differentiation. Moreover, we established shipping and cryobanking procedures to lead the study to become an off-the-shelf therapy. During expansion, the cells demonstrated a linear increase in cell numbers, confirming their proliferation quantitatively. The cells showed viability before and after cryopreservation, demonstrating that cell viability can be preserved. From a clinical perspective, the established shipping conditions demonstrated that the cells retain their viability for up to 48 h. This study lays the groundwork for the potential use of allogeneic cAD-MSCs in clinical applications. Full article
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<p>The proliferation of cAD-MSCs at passage 2 was assessed by MTS proliferation assay. Data were normalized using cell growth media alone as the control. Note the linear trend (R<sup>2</sup> = 0.9477) in proliferation with time.</p>
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<p>MSC surface marker expressions. Representative images of immunofluorescence show the expression of CD29 (<b>A</b>), CD44 (<b>B</b>), and CD90 (<b>C</b>). Cells were fixed and stained at 24 h post-seeding.</p>
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<p>In vitro differentiation of cAD-MSCs. Representative images demonstrating the in vitro potential of cAD-MSCs to undergo differentiation. (<b>A</b>) Adipogenic differentiation was examined using Oil Red O staining. Note that the cell morphology and the Oil Red O-stained cells appear at around day 9 and continue to progress with time. (<b>B</b>) Osteogenic differentiation was examined using alizarin red staining. Note that the cell morphology and the alizarin red-stained cells appear as nodules around day 17 and progress with time. Nodules that are rich in calcium are the hallmark features of osteogenic differentiation. Insets show the corresponding undifferentiated control cAD-MSCs. These cells were maintained in normal growth media without any inducers.</p>
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<p>Percentage of live to dead cells over a period of 75 h in shipping conditions (ambient temperature of transport vehicle). The viability of cAD-MSCs was maintained in shipping conditions for over 48 h. Viability decreases at 75 h post-preparation.</p>
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<p>Extracellular matrix (ECM) protein expressions by immunofluorescence. (<b>A</b>) Collagen Type I. (<b>B</b>) Collagen Type II. (<b>C</b>) Fibronectin. (<b>D</b>) Vimentin. (<b>E</b>) Vinculin. (<b>F</b>) F-actin. Even though the ECM expressions are not the same between all these images from different proteins, they demonstrate the expression of all the ECM proteins.</p>
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21 pages, 16172 KiB  
Article
Taurine Protects against Silica Nanoparticle-Induced Apoptosis and Inflammatory Response via Inhibition of Oxidative Stress in Porcine Ovarian Granulosa Cells
by Fenglei Chen, Jiarong Sun, Rongrong Ye, Tuba Latif Virk, Qi Liu, Yuguo Yuan and Xianyu Xu
Animals 2024, 14(20), 2959; https://doi.org/10.3390/ani14202959 - 14 Oct 2024
Viewed by 245
Abstract
Silica nanoparticles (SNPs) induce reproductive toxicity through ROS production, which significantly limits their application. The protective effects of taurine (Tau) against SNP-induced reproductive toxicity remain unexplored. So this study aims to investigate the impact of Tau on SNP-induced porcine ovarian granulosa cell toxicity. [...] Read more.
Silica nanoparticles (SNPs) induce reproductive toxicity through ROS production, which significantly limits their application. The protective effects of taurine (Tau) against SNP-induced reproductive toxicity remain unexplored. So this study aims to investigate the impact of Tau on SNP-induced porcine ovarian granulosa cell toxicity. In vitro, granulosa cells were exposed to SNPs combined with Tau. The localization of SNPs was determined by TEM. Cell viability was examined by CCK-8 assay. ROS levels were measured by CLSM and FCM. SOD and CAT levels were evaluated using ELISA and qPCR. Cell apoptosis was detected by FCM, and pro-inflammatory cytokine transcription levels were measured by qPCR. The results showed that SNPs significantly decreased cell viability, while increased cell apoptosis and ROS levels. Moreover, SOD and CAT were decreased, while IFN-α, IFN-β, IL-1β, and IL-6 were increased after SNP exposures. Tau significantly decreased intracellular ROS, while it increased SOD and CAT compared to SNPs alone. Additionally, Tau exhibited anti-inflammatory effects and inhibited cell apoptosis. On the whole, these findings suggest that Tau mitigates SNP-induced cytotoxicity by reducing oxidative stress, inflammatory response, and cell apoptosis. Tau may be an effective strategy to alleviate SNP-induced toxicity and holds promising application prospects in the animal husbandry and veterinary industry. Full article
(This article belongs to the Special Issue Developmental and Reproductive Toxicity of Nanoparticles in Animals)
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<p>Characteristics and cytotoxicity of SNPs: (<b>A</b>) Representative TEM image of SNPs. (<b>B</b>) Size distribution of SNPs. (<b>C</b>) CCK-8 assay. (<b>D</b>) LDH leakage assay. ** <span class="html-italic">p</span> &lt; 0.01, and **** <span class="html-italic">p</span> &lt; 0.0001 vs. Control.</p>
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<p>Cellular uptake and distribution of SNPs in porcine ovarian granulosa cells: (<b>A</b>) Representative TEM image in the control group. The cells were not exposed to SNPs. (<b>B</b>) Zoomed-in image of the white box in (<b>A</b>). (<b>C</b>) Zoomed-in image of the black box in (<b>B</b>). (<b>D</b>) Representative TEM image in the SNP-exposed group. The cells were exposed to 400 μg/mL SNPs for 48 h. (<b>E</b>) Zoomed-in image of the white box in (<b>D</b>). (<b>F</b>) Zoomed-in image of the black box in (<b>E</b>). Black arrows indicate vesicles in the cytoplasm and white arrows indicate SNPs. Scale bar, 5 μm (<b>A</b>,<b>D</b>), 2 μm (<b>B</b>,<b>E</b>), and 1 μm (<b>C</b>,<b>F</b>).</p>
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<p>SNP-induced oxidative stress in porcine ovarian granulosa cells. (<b>A</b>) Representative images of ROS staining by CLSM. Ovarian granulosa cells were exposed to control (<b>a</b>), 200 (<b>b</b>), 400 (<b>c</b>), and 800 (<b>d</b>) μg/mL SNPs for 48 h. Green indicates fluorescence of ROS and blue indicates the nucleus of ovarian granulosa cells. Scale bar, 30 μm. (<b>B</b>) Quantitative analysis of the intracellular ROS levels by FCM. (<b>C</b>) Corresponding analysis of fluorescence intensity in Figure (<b>B</b>). (<b>D</b>) Quantitative analysis of CAT mRNA levels by qPCR. (<b>E</b>) Quantitative analysis of SOD mRNA levels by qPCR. (<b>F</b>) Quantitative analysis of CAT enzyme activity by ELISA. (<b>G</b>) Quantitative analysis of SOD enzyme activity by ELISA. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control.</p>
Full article ">Figure 3 Cont.
<p>SNP-induced oxidative stress in porcine ovarian granulosa cells. (<b>A</b>) Representative images of ROS staining by CLSM. Ovarian granulosa cells were exposed to control (<b>a</b>), 200 (<b>b</b>), 400 (<b>c</b>), and 800 (<b>d</b>) μg/mL SNPs for 48 h. Green indicates fluorescence of ROS and blue indicates the nucleus of ovarian granulosa cells. Scale bar, 30 μm. (<b>B</b>) Quantitative analysis of the intracellular ROS levels by FCM. (<b>C</b>) Corresponding analysis of fluorescence intensity in Figure (<b>B</b>). (<b>D</b>) Quantitative analysis of CAT mRNA levels by qPCR. (<b>E</b>) Quantitative analysis of SOD mRNA levels by qPCR. (<b>F</b>) Quantitative analysis of CAT enzyme activity by ELISA. (<b>G</b>) Quantitative analysis of SOD enzyme activity by ELISA. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control.</p>
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<p>SNP-activated inflammatory response in porcine ovarian granulosa cells: (<b>A</b>–<b>D</b>) Quantitative analysis of the mRNA levels for IFN-α (<b>A</b>), IFN-β (<b>B</b>), IL-1β (<b>C</b>), and IL-6 (<b>D</b>) by qPCR. * <span class="html-italic">p</span> &lt; 0.05 and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control.</p>
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<p>Cell apoptosis was activated in porcine ovarian granulosa cells after SNP exposures: (<b>A</b>) The apoptotic rate was determined by FCM. Q1-UL quadrant represents cell death caused by mechanical damage or necrotic cells, Q1-UR quadrant represents late apoptotic cells, Q1-LL quadrant represents the normal cells, and Q1-LR quadrant represents early apoptotic cells. (<b>B</b>) Quantification of the apoptotic rate. (<b>C</b>–<b>F</b>) Quantitative analysis of the mRNA levels for BCL-2 (<b>C</b>), BAX (<b>D</b>), Caspase-3 (<b>E</b>), and PARP (<b>F</b>) by qPCR. (<b>G</b>) Detection of BCL-2, BAX, cleaved Caspase-3, and cleaved PARP expressions by Western Blot. (<b>H</b>) Quantitative analysis of the band intensity for BCL-2, BAX, cleaved Caspase-3, and cleaved PARP. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control.</p>
Full article ">Figure 5 Cont.
<p>Cell apoptosis was activated in porcine ovarian granulosa cells after SNP exposures: (<b>A</b>) The apoptotic rate was determined by FCM. Q1-UL quadrant represents cell death caused by mechanical damage or necrotic cells, Q1-UR quadrant represents late apoptotic cells, Q1-LL quadrant represents the normal cells, and Q1-LR quadrant represents early apoptotic cells. (<b>B</b>) Quantification of the apoptotic rate. (<b>C</b>–<b>F</b>) Quantitative analysis of the mRNA levels for BCL-2 (<b>C</b>), BAX (<b>D</b>), Caspase-3 (<b>E</b>), and PARP (<b>F</b>) by qPCR. (<b>G</b>) Detection of BCL-2, BAX, cleaved Caspase-3, and cleaved PARP expressions by Western Blot. (<b>H</b>) Quantitative analysis of the band intensity for BCL-2, BAX, cleaved Caspase-3, and cleaved PARP. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control.</p>
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<p>Tau inhibited SNP-induced oxidative stress in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>) Representative images of ROS staining by CLSM. Ovarian granulosa cells were exposed to control (<b>a</b>), 200 μg/mL SNP group (<b>b</b>), 400 μg/mL SNP (<b>c</b>), 800 μg/mL SNP (<b>d</b>), 10 mM Tau (<b>e</b>), 200 μg/mL SNP combined with 10 mM Tau (<b>f</b>), 400 μg/mL SNP combined with 10 mM Tau (<b>g</b>), and 800 μg/mL SNP combined with 10 mM Tau (<b>h</b>). Green indicates fluorescence of ROS and blue indicates the nucleus of ovarian granulosa cells. Scale bar, 30 μm. (<b>B</b>) Quantitative analysis of the intracellular ROS levels by FACS. (<b>C</b>) Corresponding analysis of the fluorescence intensity in Figure (<b>B</b>). (<b>D</b>) Quantitative analysis of CAT mRNA levels by qPCR. (<b>E</b>) Quantitative analysis of SOD mRNA levels by qPCR. (<b>F</b>) Quantitative analysis of CAT enzyme activity by ELISA. (<b>G</b>) Quantitative analysis of SOD enzyme activity by ELISA. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05 and <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01 vs. SNP-exposed group.</p>
Full article ">Figure 6 Cont.
<p>Tau inhibited SNP-induced oxidative stress in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>) Representative images of ROS staining by CLSM. Ovarian granulosa cells were exposed to control (<b>a</b>), 200 μg/mL SNP group (<b>b</b>), 400 μg/mL SNP (<b>c</b>), 800 μg/mL SNP (<b>d</b>), 10 mM Tau (<b>e</b>), 200 μg/mL SNP combined with 10 mM Tau (<b>f</b>), 400 μg/mL SNP combined with 10 mM Tau (<b>g</b>), and 800 μg/mL SNP combined with 10 mM Tau (<b>h</b>). Green indicates fluorescence of ROS and blue indicates the nucleus of ovarian granulosa cells. Scale bar, 30 μm. (<b>B</b>) Quantitative analysis of the intracellular ROS levels by FACS. (<b>C</b>) Corresponding analysis of the fluorescence intensity in Figure (<b>B</b>). (<b>D</b>) Quantitative analysis of CAT mRNA levels by qPCR. (<b>E</b>) Quantitative analysis of SOD mRNA levels by qPCR. (<b>F</b>) Quantitative analysis of CAT enzyme activity by ELISA. (<b>G</b>) Quantitative analysis of SOD enzyme activity by ELISA. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05 and <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01 vs. SNP-exposed group.</p>
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<p>Tau inhibited SNP-activated inflammatory response in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>–<b>D</b>) Quantitative analysis of the mRNA levels for IFN-α (<b>A</b>), IFN-β (<b>B</b>), IL-1β (<b>C</b>), and IL-6 (<b>D</b>) by qPCR. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05, <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01, and <sup>###</sup> <span class="html-italic">p</span> &lt; 0.001 vs. SNP-exposed group.</p>
Full article ">Figure 7 Cont.
<p>Tau inhibited SNP-activated inflammatory response in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>–<b>D</b>) Quantitative analysis of the mRNA levels for IFN-α (<b>A</b>), IFN-β (<b>B</b>), IL-1β (<b>C</b>), and IL-6 (<b>D</b>) by qPCR. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05, <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01, and <sup>###</sup> <span class="html-italic">p</span> &lt; 0.001 vs. SNP-exposed group.</p>
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<p>Tau inhibited SNP-induced cell apoptosis in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>) The apoptotic rate was determined by FCM. Q1-UL quadrant represents cell death caused by mechanical damage or necrotic cells, Q1-UR quadrant represents late apoptotic cells, Q1-LL quadrant represents the normal cells, and Q1-LR quadrant represents early apoptotic cells. (<b>B</b>) Quantification of the apoptotic rate. (<b>B</b>) Quantification of the apoptotic rate. (<b>C</b>–<b>F</b>) Quantitative analysis of the mRNA levels for BCL-2 (<b>C</b>), BAX (<b>D</b>), Caspase-3 (<b>E</b>), and PARP (F) by qPCR. (<b>G</b>) Detection of BCL-2, BAX, cleaved Caspase-3, and cleaved PARP expressions by Western Blot. (<b>H</b>) Quantitative analysis of the band intensity for BCL-2, BAX, cleaved Caspase-3, and cleaved PARP. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05, <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01, and <sup>###</sup> <span class="html-italic">p</span> &lt; 0.001 vs. SNP-exposed group.</p>
Full article ">Figure 8 Cont.
<p>Tau inhibited SNP-induced cell apoptosis in porcine ovarian granulosa cells. Ovarian granulosa cells were exposed to SNPs in the absence or presence of 10 mM Tau for 48 h: (<b>A</b>) The apoptotic rate was determined by FCM. Q1-UL quadrant represents cell death caused by mechanical damage or necrotic cells, Q1-UR quadrant represents late apoptotic cells, Q1-LL quadrant represents the normal cells, and Q1-LR quadrant represents early apoptotic cells. (<b>B</b>) Quantification of the apoptotic rate. (<b>B</b>) Quantification of the apoptotic rate. (<b>C</b>–<b>F</b>) Quantitative analysis of the mRNA levels for BCL-2 (<b>C</b>), BAX (<b>D</b>), Caspase-3 (<b>E</b>), and PARP (F) by qPCR. (<b>G</b>) Detection of BCL-2, BAX, cleaved Caspase-3, and cleaved PARP expressions by Western Blot. (<b>H</b>) Quantitative analysis of the band intensity for BCL-2, BAX, cleaved Caspase-3, and cleaved PARP. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.01, and *** <span class="html-italic">p</span> &lt; 0.001 vs. Control. <sup>#</sup> <span class="html-italic">p</span> &lt; 0.05, <sup>##</sup> <span class="html-italic">p</span> &lt; 0.01, and <sup>###</sup> <span class="html-italic">p</span> &lt; 0.001 vs. SNP-exposed group.</p>
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10 pages, 1591 KiB  
Article
Luteolin (LUT) Induces Apoptosis and Regulates Mitochondrial Membrane Potential to Inhibit Cell Growth in Human Cervical Epidermoid Carcinoma Cells (Ca Ski)
by Sung-Nan Pei, Kuan-Ting Lee, Kun-Ming Rau, Tsung-Ying Lin, Tai-Hsin Tsai and Yi-Chiang Hsu
Biomedicines 2024, 12(10), 2330; https://doi.org/10.3390/biomedicines12102330 - 14 Oct 2024
Viewed by 325
Abstract
Background/Objectives: Luteolin (LUT) is a natural flavonoid with known anti-inflammatory, antioxidant, and anti-cancer properties. Cervical cancer, particularly prevalent in certain regions, remains a significant health challenge due to its high recurrence and poor response to treatment. This study aimed to investigate the anti-tumor [...] Read more.
Background/Objectives: Luteolin (LUT) is a natural flavonoid with known anti-inflammatory, antioxidant, and anti-cancer properties. Cervical cancer, particularly prevalent in certain regions, remains a significant health challenge due to its high recurrence and poor response to treatment. This study aimed to investigate the anti-tumor effects of LUT on human cervical epidermoid carcinoma cells (Ca Ski), focusing on cell growth inhibition, apoptosis induction, and regulation of mitochondrial membrane potential. Methods: Ca Ski cells were treated with varying concentrations of LUT (0, 25, 50, 100 µM) for different time periods (24, 48, 72 hours). Cell viability was measured using the MTT assay, apoptosis was assessed by flow cytometry with annexin V-FITC/PI staining, and changes in mitochondrial membrane potential were evaluated using JC-1 staining. Caspase-3 activation was examined by flow cytometry, and expression of apoptosis-related proteins (caspase-3, -8, -9, AIF) was analyzed via Western blotting. Results: LUT significantly inhibited the growth of Ca Ski cells in a dose- and time-dependent manner, with the most pronounced effects observed at 100 µM over 72 hours. Flow cytometry confirmed that LUT induced apoptosis without causing necrosis. Mitochondrial membrane potential was reduced after LUT treatment, coinciding with increased caspase-3 activation. Western blot analysis revealed the upregulation of pro-apoptotic proteins caspase-3, -8, -9, and AIF, indicating that LUT induces apoptosis through the intrinsic mitochondrial pathway. Conclusions: Luteolin effectively inhibits cervical cancer cell proliferation and induces apoptosis by disrupting mitochondrial membrane potential and activating caspases. These findings suggest that LUT holds potential as a therapeutic agent for cervical cancer, with further studies needed to explore its in vivo efficacy and broader clinical applications. Full article
(This article belongs to the Section Cell Biology and Pathology)
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<p>LUT affects the survival of cervical cancer cells (Ca Ski) and inhibits their proliferation. Ca Ski cells were treated with varying concentrations of LUT (0, 25, 50, and 100 µM) for 24 to 72 h, and cell viability was measured using the MTT assay. Results are presented as a percentage relative to the control, which is set at 100%. Data are expressed as the mean ± SEM from at least three experiments. Statistical significance was determined using a <span class="html-italic">t</span>-test, with * <span class="html-italic">p</span> &lt; 0.05 indicating a significant difference from the control group, # <span class="html-italic">p</span> &lt; 0.05 indicating a significant difference from the 24 h group, and &amp; <span class="html-italic">p</span> &lt; 0.05 indicating a significant difference from the 48 hrs group.</p>
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<p>The role of LUT in inducing cell apoptosis in cervical cancer cells (Ca Ski). (<b>A</b>) The extent of total apoptosis and necrosis in Ca Ski cells after 4 h of incubation with LUT. (<b>B</b>) Results are presented as a percentage relative to the control group, including necrosis and the total number of apoptotic cells (both early and late apoptosis). Statistical significance was assessed using a <span class="html-italic">t</span>-test, with * <span class="html-italic">p</span> &lt; 0.05 indicating a significant difference compared to the control group.</p>
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<p>Influence of LUT on cell cycle progression/distribution in cervical cancer cells. (<b>A</b>) Cell cycle analysis of Ca Ski cells after 24 h of treatment with LUT. (<b>B</b>) LUT treatment led to an increase in the percentage of cells in the sub-G1 phase. Results are presented as the mean ± SD from three experiments. The asterisk (*) in each bar group indicates that the difference compared to the 0 µM LUT treatment is statistically significant at <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of LUT on caspase 3 activation and mitochondrial membrane potential. (<b>A</b>) LUT-induced changes in mitochondrial membrane potential (ΔΨm) were assessed in Ca Ski cells. ΔΨm levels were determined using JC-1 staining and flow cytometry. (<b>B</b>) Cells were treated with varying concentrations of LUT for 24 h. Statistical significance is indicated by * <span class="html-italic">p</span> &lt; 0.05. (<b>C</b>) Following 24 h treatment with LUT, cells were harvested and labeled with FITC-conjugated anti-active caspase 3 antibody. (<b>D</b>) Caspase 3 activation was quantified by flow cytometry. All data are expressed as mean ± SD from three independent experiments, with statistical significance denoted by * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The expression of LUT-regulated caspase family in Ca Ski cells. (<b>A</b>) The expression of pro-caspase 3, 8, 9, and AIF were assessed by Western blot analysis in Ca Ski cells. (<b>B</b>) The quantification of pro-caspase 3, 8, 9, and AIF is shown in the bar graph. Data were normalized to GAPDH as a control and expressed as a percentage relative to the control group, which was set at 100%. All results are presented as the mean (±SEM) of at least three independent experiments. Statistical significance was determined using a <span class="html-italic">t</span>-test, with * <span class="html-italic">p</span> &lt; 0.05 indicating significant differences compared to the control group.</p>
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32 pages, 11462 KiB  
Article
Selective Inhibition of Deamidated Triosephosphate Isomerase by Disulfiram, Curcumin, and Sodium Dichloroacetate: Synergistic Therapeutic Strategies for T-Cell Acute Lymphoblastic Leukemia in Jurkat Cells
by Luis A. Flores-López, Ignacio De la Mora-De la Mora, Claudia M. Malagón-Reyes, Itzhel García-Torres, Yoalli Martínez-Pérez, Gabriela López-Herrera, Gloria Hernández-Alcántara, Gloria León-Avila, Gabriel López-Velázquez, Alberto Olaya-Vargas, Saúl Gómez-Manzo and Sergio Enríquez-Flores
Biomolecules 2024, 14(10), 1295; https://doi.org/10.3390/biom14101295 - 13 Oct 2024
Viewed by 436
Abstract
T-cell acute lymphoblastic leukemia (T-ALL) is a challenging childhood cancer to treat, with limited therapeutic options and high relapse rates. This study explores deamidated triosephosphate isomerase (dTPI) as a novel therapeutic target. We hypothesized that selectively inhibiting dTPI could reduce T-ALL cell viability [...] Read more.
T-cell acute lymphoblastic leukemia (T-ALL) is a challenging childhood cancer to treat, with limited therapeutic options and high relapse rates. This study explores deamidated triosephosphate isomerase (dTPI) as a novel therapeutic target. We hypothesized that selectively inhibiting dTPI could reduce T-ALL cell viability without affecting normal T lymphocytes. Computational modeling and recombinant enzyme assays revealed that disulfiram (DS) and curcumin (CU) selectively bind and inhibit dTPI activity without affecting the non-deamidated enzyme. At the cellular level, treatment with DS and CU significantly reduced Jurkat T-ALL cell viability and endogenous TPI enzymatic activity, with no effect on normal T lymphocytes, whereas the combination of sodium dichloroacetate (DCA) with DS or CU showed synergistic effects. Furthermore, we demonstrated that dTPI was present and accumulated only in Jurkat cells, confirming our hypothesis. Finally, flow cytometry confirmed apoptosis in Jurkat cells after treatment with DS and CU or their combination with DCA. These findings strongly suggest that targeting dTPI represents a promising and selective target for T-ALL therapy. Full article
(This article belongs to the Section Enzymology)
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Figure 1

Figure 1
<p>Molecular docking analysis of n-dTPI and dTPI crystallographic structures. Conformational representations of DS docked at the interface of n-dTPI (<b>a</b>) and dTPI (<b>b</b>) are shown. The proximity of Cys residues near the interface is highlighted, suggesting potential interaction sites for DS. Conformational representations of CU docked at the interface of n-dTPI (<b>c</b>) and dTPI (<b>d</b>) are shown; the hydrophobic surfaces of proteins are highlighted in red. A deeper penetration of both DS and CU into the interface of the deamidated enzyme (dTPI) compared to the non-deamidated form (n-dTPI) is observed. Figures modeled with PyMOL version 2.5.0 (Schrödinger Inc., New York, NY, USA).</p>
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<p>Inactivation assays of recombinant n-dTPI and dTPI enzymes. Both enzymes (0.2 mg/mL) were incubated for 2 h at 37 °C with gradually increasing concentrations of the respective compounds: (<b>a</b>) DS (0 to 1000 μM) and (<b>b</b>) CU (0 to 1500 μM). Following incubation, aliquots were taken for enzyme activity measurement as described in the Methods section. The enzymatic activity was normalized, and 100% corresponds to the assays in the absence of the compound. Filled black squares represent n-dTPI activity, while filled red squares represent dTPI activity. The results represent the average of three independent experiments, with error bars indicating the variation observed in the experiments.</p>
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<p>Extrinsic fluorescence spectra of TPIs after incubation with DS and CU. Enzymes (0.2 mg/mL) were incubated without or with 250 μM DS and 1500 μM CU for 2 h at 37 °C. Following incubation, excess compounds were removed, and extrinsic fluorescence was measured in the presence of 100 μM ANS with excitation at 395 nm. (<b>a</b>) shows the spectra in the absence of any compounds (control). (<b>b</b>,<b>c</b>) show the spectra after incubation with DS and CU, respectively. n-dTPI (black line) and dTPI (red line). The results represent the mean of three independent experiments.</p>
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<p>Effects of DS and CU on cell viability and TPI activity in normal T-cell lymphocytes and Jurkat cells. Cells (1 × 10<sup>5</sup> per well) were incubated with increasing concentrations of DS and CU. Following incubation, cell viability was assessed using MTT assays and endogenous TPI activity was determined by enzymatic activity assays. (<b>a</b>,<b>c</b>) normal T lymphocytes, (<b>b</b>,<b>d</b>) Jurkat cells. Results are expressed as percentages relative to the untreated control set to 100%. The results represent the average of three independent experiments, with error bars indicating the variation observed in the experiments. Statistical differences were analyzed using one-way ANOVA with Tukey’s post-hoc test, with a significance level set at <span class="html-italic">p</span> = 0.01 (**).</p>
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<p>Effects of combined DCA and CU treatment on cell viability and TPI activity in normal T lymphocytes and Jurkat cells. Cells (1 × 10<sup>5</sup> per well) were pre-treated with 12 mM DCA for 24 h at 37 °C, followed by exposure to increasing concentrations of DS or CU for an additional 24 h. MTT and enzyme activity assays were then performed to assess cell viability and endogenous TPI activity, respectively. (<b>a</b>,<b>c</b>) normal T lymphocytes and (<b>b</b>,<b>d</b>) Jurkat cells. Results are expressed as percentages relative to the untreated control group (set to 100%). The results represent the average of three independent experiments, with error bars indicating the variation observed in the experiments. Statistical differences were analyzed using one-way ANOVA with Tukey’s post-hoc test, with a significance level set at <span class="html-italic">p</span> = 0.01 (**).</p>
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<p>Western blot analysis of TPI isoforms in normal T lymphocytes and Jurkat cells. In (<b>a</b>) and (<b>b</b>), lanes 1, 2, and 3, each loaded with 1 μg of protein, serve as migration standards for recombinant n-dTPI, dTPI, and ddTPI, respectively. Lanes 4–6 in (<b>a</b>) contain total protein extracts from normal T lymphocytes, while lanes 7–9 in (<b>a</b>) and lanes 4–10 in (<b>b</b>) correspond to protein extracts from Jurkat cells, with 100 μg of protein loaded per lane. For normal T lymphocytes, lane 4 is the control (untreated), lane 5 is treated with 250 μM DS, and lane 6 is treated with 1500 μM CU. In Jurkat cells, lanes 7, 8, and 9 in (<b>a</b>) represent the control (untreated), treatment with 250 μM DS, and treatment with 1500 μM CU, respectively. In (<b>b</b>), lane 4 is the control condition (untreated), while lanes 6 and 7 were treated with 100 and 250 μM DS, respectively. Finally, lanes 8, 9, and 10 were treated with 500, 1000, and 1500 μM of CU, respectively. The positive and negative poles of the gel are indicated on the left side of each panel. Full-length (uncropped) blots for panels (<b>a</b>,<b>b</b>) are presented in <a href="#app1-biomolecules-14-01295" class="html-app">Supplementary Figure S5</a>.</p>
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<p>Western blot analysis of TPI isoforms in normal T-cell lymphocytes and Jurkat cells with combined DCA and CU treatment. In both panels, lanes 1–3 represent recombinant n-dTPI, dTPI, and ddTPI enzymes loaded at 1 μg protein per lane, serving as migration standards. In both panels, lanes 4–9 with 100 μg of protein loaded per lane. In a and b, lane 4 shows control Jurkat cells (untreated); lane 5 shows 12 mM DCA. In (<b>a</b>) lane 6 and 7 are Jurkat cells pretreated with 12 mM DCA followed by incubation with 100 and 250 μM DS, respectively. In (<b>b</b>) lane 6 and 7 are Jurkat cells pretreated with 12 mM DCA followed by incubation with 1000 μM and 1500 μM CU, respectively. Finally, in both, a and b, lanes 8–9 show normal T lymphocytes pretreated with 12 mM DCA followed by incubation with 250 μM DS (<b>a</b>) or 1500 μM CU (<b>b</b>). The polarity of the gel is indicated on the left side of the panel. Full-length blots (uncropped blots) are shown in <a href="#app1-biomolecules-14-01295" class="html-app">Supplementary Figure S6</a>.</p>
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<p>MGO and AGEs levels in Jurkat cells following DCA, DS, and CU treatment. Quantification of MGO (<b>a</b>,<b>c</b>) and AGEs (<b>b</b>,<b>d</b>) in Jurkat cells following treatment with DCA, DS, CU, or their combination (*). As observed, increasing concentrations of DS and CU lead to a dose-dependent rise in both MGO and AGEs. Notably, pretreatment with DCA before DS or CU administration results in a further significant increase in MGO and AGEs production. The results represent the average of three independent experiments, with error bars indicating the variation observed in the experiments.</p>
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<p>Western blot analysis of apoptosis-related proteins in Jurkat cells. (<b>a</b>) Expression of ERK1/2 and its phosphorylated form following treatment with DCA, CU, or their combination. The corresponding bar graphs quantify the decrease in total and phosphorylated ERK1/2 levels relative to the control. (<b>b</b>) Expression of Bcl-2 and Bax proteins under treatment with DCA, CU, or their combination. The bar graphs represent the relative expression levels of these proteins. (<b>c</b>) Procaspase-7 and cleaved caspase-7 levels following treatment with DCA, CU, or their combination. The bar graphs quantify the relative abundance of procaspase-7 and cleaved caspase-7 compared to the control. β-Actin was used as a loading control for all Western blots. Each lane was loaded with 100 μg of total protein extract. Statistical analysis was performed using a one-way ANOVA followed by the Tukey-Kramer test. Significance levels are indicated as follows: * <span class="html-italic">p</span> ≤ 0.01 compared to the control, ¥ <span class="html-italic">p</span> ≤ 0.01 compared to 12 mM DCA treatment, &amp; <span class="html-italic">p</span> ≤ 0.01 compared to 0.5 mM CU treatment, <span>$</span> <span class="html-italic">p</span> ≤ 0.01 compared to 12 mM DCA + 0.5 mM CU treatment. Full-length blots (uncropped blots) are shown in <a href="#app1-biomolecules-14-01295" class="html-app">Supplementary Figure S7b–d</a> for Total ERK 1/2, pERK, and β-Actin, respectively; <a href="#app1-biomolecules-14-01295" class="html-app">Supplementary Figure S8b–d</a> for Bcl2, Bax, and β-Actin, respectively, and <a href="#app1-biomolecules-14-01295" class="html-app">Supplementary Figure S9b,c</a> for Procaspase-7, cleaved caspase-7, and β-Actin, respectively.</p>
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<p>Detection of cell death in normal T lymphocytes and Jurkat cells after treatment with DCA, DS, CU, or their combinations (DCA + DS or DCA + CU). Cells were incubated with 12 mM DCA, 100 µM DS, or 250 µM CU for 24 h at 37 °C. For combination treatments, cells were first exposed to 12 mM DCA for 24 h, followed by an additional 24-h incubation with 100 µM DS or 250 µM CU. At the end of the incubation period, cells were washed, resuspended at a density of 1 × 10<sup>6</sup>/mL, and stained with Annexin V and Propidium Iodide, for flow cytometry analysis. (<b>a</b>) shows untreated control cells. (<b>b</b>–<b>d</b>) represent cells treated with DCA, DS, and CU, respectively. (<b>e</b>,<b>f</b>) display cells pretreated with DCA followed by DS and CU, respectively. Quadrants indicate distinct cell populations: Q1 (necrotic cells), Q2 (late apoptotic cells), and Q4 (early apoptotic cells). The data presented in the figure are representative of 100,000 cells analyzed across two independent experiments.</p>
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<p>Detection of cell death in normal T lymphocytes and Jurkat cells after treatment with DCA, DS, CU, or their combinations (DCA + DS or DCA + CU). Cells were incubated with 12 mM DCA, 100 µM DS, or 250 µM CU for 24 h at 37 °C. For combination treatments, cells were first exposed to 12 mM DCA for 24 h, followed by an additional 24-h incubation with 100 µM DS or 250 µM CU. At the end of the incubation period, cells were washed, resuspended at a density of 1 × 10<sup>6</sup>/mL, and stained with Annexin V and Propidium Iodide, for flow cytometry analysis. (<b>a</b>) shows untreated control cells. (<b>b</b>–<b>d</b>) represent cells treated with DCA, DS, and CU, respectively. (<b>e</b>,<b>f</b>) display cells pretreated with DCA followed by DS and CU, respectively. Quadrants indicate distinct cell populations: Q1 (necrotic cells), Q2 (late apoptotic cells), and Q4 (early apoptotic cells). The data presented in the figure are representative of 100,000 cells analyzed across two independent experiments.</p>
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17 pages, 8054 KiB  
Article
Incorporation of Superparamagnetic Magnetic–Fluorescent Iron Oxide Nanoparticles Increases Proliferation of Human Mesenchymal Stem Cells
by Willian Pinheiro Becker, Juliana Barbosa Torreão Dáu, Wanderson de Souza, Rosalia Mendez-Otero, Rosana Bizon Vieira Carias and Jasmin
Magnetochemistry 2024, 10(10), 77; https://doi.org/10.3390/magnetochemistry10100077 - 12 Oct 2024
Viewed by 379
Abstract
Mesenchymal stem cells (MSCs) have significant therapeutic potential and their use requires in-depth studies to better understand their effects. Labeling cells with superparamagnetic iron oxide nanoparticles allows real-time monitoring of their location, migration, and fate post-transplantation. This study aimed to investigate the efficacy [...] Read more.
Mesenchymal stem cells (MSCs) have significant therapeutic potential and their use requires in-depth studies to better understand their effects. Labeling cells with superparamagnetic iron oxide nanoparticles allows real-time monitoring of their location, migration, and fate post-transplantation. This study aimed to investigate the efficacy and cytotoxicity of magnetic–fluorescent nanoparticles in human adipose tissue-derived mesenchymal stem cells (hADSCs). The efficacy of Molday ION rhodamine B (MIRB) labeling in hADSCs was evaluated and their biocompatibility was assessed using various techniques and differentiation assays. Prussian blue and fluorescence staining confirmed that 100% of the cells were labeled with MIRB and this labeling persisted for at least 3 days. Transmission electron microscopy revealed the internalization and clustering of the nanoparticles on the outer surface of the cell membrane. The viability assay showed increased cell viability 3 days after nanoparticle exposure. Cell counts were higher in the MIRB-treated group compared to the control group at 3 and 5 days and an increased cell proliferation rate was observed at 3 days post-exposure. Adipogenic, osteogenic, and chondrogenic differentiation was successfully achieved in all groups, with MIRB-treated cells showing an enhanced differentiation rate into adipocytes and osteocytes. MIRB was efficiently internalized by hADSCs but induced changes in cellular behavior due to the increased cell proliferation rate. Full article
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Figure 1
<p>Characterization of MIRB NPs: (<b>A</b>) Transmission electron microscopy (TEM) image of MIRB in water. Images kindly provided by MAGTech Brazil; (<b>B</b>) hydrodynamic diameter (HD, nm) and polydispersity index (PdI) of MIRB NPs in water and cell culture medium determined by dynamic light scattering (DLS). (<b>C</b>) Surface charge analysis of the zeta potential of MIRB (ζ (mV)) in water and culture media. A statistically significant difference was found in the diameter (<span class="html-italic">p</span> = 0.004), PdI (<span class="html-italic">p</span> &lt; 0.0001) and zeta potential (* <span class="html-italic">p</span> = 0.009) between the groups.</p>
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<p>Labeling of hADSCs with MIRB for 18 h. (<b>A</b>) Control cells. (<b>B</b>) MIRB-treated cells were stained with Prussian blue. (<b>C</b>) Fluorescence optical microscopy image demonstrating the nuclei stained with DAPI (blue) and the nanoparticles (red). Scale bar = 100 μm (<b>A</b>–<b>C</b>). (<b>D</b>) Quantification of MIRB-positive hADSCs. (<b>E</b>) Transmission electron micrograph of control and (<b>F</b>) MIRB-interacting cells. Arrows show clustering of MIRB on the cell membrane, and arrowheads indicate intracellular endosomal vesicles with nanoparticles. (<b>G</b>) Endosomal vesicle at high magnification demonstrating the internalization of MIRB. Scale bar = 2 μm (<b>E</b>,<b>F</b>); 300 nm (<b>G</b>).</p>
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<p>Colorimetric cell viability assay after treatment with MIRB. The graph represents the cell viability analyzed 3 days after the start of exposure. *** <span class="html-italic">p</span> &lt; 0.01.</p>
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<p>Quantification of hADSCs treated with MIRB for different durations and stained with Prussian blue and acid fuchsin. (<b>A</b>) Control cells on day 1. (<b>B</b>) MIRB-treated cells on day 1. (<b>C</b>) Control cells on day 2. (<b>D</b>) MIRB-treated cells on day 2. (<b>E</b>) Control cells on day 3. (<b>F</b>) MIRB-treated cells on day 3. Scale bars = 100 μm. (<b>G</b>) hADSCs were treated and quantified daily for up to 5 days after initial exposure. The black bar represents control hADSCs, and the gray bar represents hADSCs treated with MIRB. * <span class="html-italic">p</span> &lt; 0.05, *** <span class="html-italic">p</span> &lt; 0.0001.</p>
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<p>Immunofluorescence of hADSCs treated or not treated with MIRB. (<b>A</b>) Control cells stained with DAPI (blue) and (<b>A’</b>) anti-Ki67 antibody (green) and (<b>A”</b>) overlay of images. (<b>B</b>) Representative images of cells treated with MIRB and stained with DAPI (blue) and (<b>B’</b>) anti-Ki67 antibody (green). (<b>B”</b>) Overlay of images B and B’. (<b>B’”</b>) Overlay of images B and B’ together with rhodamine B (red) detection. Scale bar = 100 μm. (<b>C</b>) Quantification of Ki67-positive hADSCs on day 1 and day 3 after treatment. * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>Analysis of the adipogenic differentiation potential of MIRB-treated hADSCs revealed by Oil Red O staining. (<b>A</b>) Control cells not induced or (<b>B</b>) induced to adipogenesis. (<b>C</b>) MIRB-induced hADSCs. (<b>D</b>) MIRB-hADSCs were not induced to differentiate but were also stained with Prussian blue and acid fuchsin 1%; note the MIRB-positive cells 21 days after culture. (<b>E</b>) Differentiated MIRB cells stained with Prussian blue and (<b>E’</b>) higher magnification of the marked square in image E demonstrating lipid vacuoles and the presence of MIRB in the same cell after 21 days. (<b>F</b>) Percentage of hADSCs positive for oil red O dye. (<b>G</b>) Quantification of the number of control and MIRB cells. * <span class="html-italic">p</span> &lt; 0.05, ** <span class="html-italic">p</span> &lt; 0.001. Scale bar = 100 μm.</p>
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<p>Osteogenic differentiation of hADSCs after 18 days of staining with Alizarin Red. (<b>A</b>) Quantification of Alizarin Red staining in control and induced cells. (<b>B</b>) Uninduced MIRB-treated cells. (<b>C</b>) Differentiated control hADSCs. (<b>D</b>) MIRB-treated hADSCs were induced to differentiate. ** <span class="html-italic">p</span> &lt; 0.01. Scale bar = 100 μm.</p>
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<p>Histological micromass sections of hADSCs induced into chondrocytes over 14 days. (<b>A</b>) Histological section of a micromass from induced control cells. (<b>B</b>) Histological section of micromass from induced MIRB-treated cells. (<b>C</b>) Higher magnification of a micromass from the MIRB group. (<b>D</b>) Histological section of micromass from hADSCs treated with MIRB stained with hematoxylin and eosin. Scale bars: 50 μm (<b>A</b>,<b>B</b>,<b>D</b>) and 20 μm (<b>C</b>).</p>
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27 pages, 9656 KiB  
Article
Assessment of the Antioxidant and Photoprotective Properties of Cornus mas L. Extracts on HDF, HaCaT and A375 Cells Exposed to UVA Radiation
by Martyna Zagórska-Dziok, Agnieszka Mokrzyńska, Aleksandra Ziemlewska, Zofia Nizioł-Łukaszewska, Ireneusz Sowa, Marcin Feldo and Magdalena Wójciak
Int. J. Mol. Sci. 2024, 25(20), 10993; https://doi.org/10.3390/ijms252010993 - 12 Oct 2024
Viewed by 431
Abstract
The influence of UV radiation on skin discoloration, skin aging and the development of skin cancer is widely known. As a part of this study, the effect of extracts from three varieties of Cornus mas L. (C. mas L.) on skin cells [...] Read more.
The influence of UV radiation on skin discoloration, skin aging and the development of skin cancer is widely known. As a part of this study, the effect of extracts from three varieties of Cornus mas L. (C. mas L.) on skin cells exposed to UVA radiation was assessed. The analyses were performed on both normal and cancer skin cells. For this purpose, the potential photoprotective effects of the obtained extracts (aqueous and ethanolic) was assessed by performing two cytotoxicity tests (Alamar blue and Neutral red). Additionally, the antioxidant capacity was compared using three different assays. The 2′,7′-dichlorodihydrofluorescein diacetate (H2DCFDA) probe was used to evaluate the intracellular level of free radicals in cells exposed to the simultaneous action of UVA radiation and dogwood extracts. Additionally, the ability to inhibit excessive pigmentation was determined by assessing the inhibition of melanin formation and tyrosinase activity. The obtained results confirmed the strong antioxidant properties of dogwood extracts and their photoprotective effect on normal skin cells. The ability to inhibit the viability of melanoma cells was also observed. Additionally, a reduction in oxidative stress in skin cells exposed to UVA radiation and a strong inhibition of melanin formation and tyrosinase activity have been demonstrated. This study shows that dogwood extract could be a valuable cosmetic raw material that can play both a photoprotective and antihyperpigmentation role in cosmetic preparations. Full article
(This article belongs to the Special Issue Bioactive Compounds of Natural Origin)
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark ruby-red fruits) on ABTS radical scavenging. Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark ruby-red fruits) on DPPH radical scavenging. Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001.</p>
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<p>Antioxidant capacity in FRAP assay of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark ruby-red fruits). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The results were expressed in Trolox equivalents (µmol TE/L). Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the intracellular level of reactive oxygen species in fibroblasts (HDF) exposed to UVA radiation (5 J/cm<sup>2</sup> for 1 h). The positive control (PC) are HDF cells treated with UVA radiation (without the addition of extracts), and the negative control (NC) are cells exposed to neither UVA radiation nor extracts. Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the intracellular level of reactive oxygen species in keratinocytes (HaCaT) exposed to UVA radiation (5 J/cm<sup>2</sup> for 1 h). The positive control (PC) are HaCaT cells treated with UVA radiation (without the addition of extracts), and the negative control (NC) are cells exposed to neither UVA radiation nor extracts. Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the intracellular level of reactive oxygen species in melanoma cells (A375) exposed to UVA radiation (5 J/cm<sup>2</sup> for 1 h). The positive control (PC) are A375 cells treated with UVA radiation (without the addition of extracts), and the negative control (NC) are cells exposed to neither UVA radiation nor extracts. Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the reduction of resazurin in fibroblasts after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were fibroblasts treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments, in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the neutral red dye uptake in fibroblasts after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were fibroblasts treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the reduction of resazurin in keratinocytes after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were keratinocytes treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
Full article ">Figure 10
<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the neutral red dye uptake in keratinocytes after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span> at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were keratinocytes treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments, in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the reduction of resazurin in melanoma cells after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20, and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were melanoma cells treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> &lt; 0.001, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>The effect of extracts from three varieties of <span class="html-italic">Cornus mas</span> L. (with yellow, red and dark red fruits) on the neutral red dye uptake in melanoma cells after UVA radiation (5 J/cm<sup>2</sup> for 1 h). Analyses were performed for water and two water–ethanolic extracts (30 and 70% (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) at a dilution of 1:100, 1:20 and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). The exposure time to the extracts was 24 h. The positive control (+UV) were melanoma cells treated with UVA radiation (without the addition of extracts), and the negative control (−UV) were cells exposed to neither UVA radiation nor extracts, for which the viability was assumed to be 100%. Data represent mean ± SD of three independent experiments in which each sample was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, *** <span class="html-italic">p</span> = 0.0005, ** <span class="html-italic">p</span> &lt; 0.01, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>Assessment of inhibition of melanin formation by water, ethanol–water (30:70 (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) and ethanol–water (70:30 (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) extracts from three varieties of <span class="html-italic">C. mas</span> L. Extracts were obtained from varieties with yellow, red and dark ruby-red fruit. Analyses were performed for extracts at dilution of 1:100, 1:20 and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). Data represent mean ± SD of three independent experiments in which each dilution of individual dogwood extracts was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001, ** <span class="html-italic">p</span> = 0.0029, * <span class="html-italic">p</span> &lt; 0.05.</p>
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<p>Assessment of tyrosinase level in B16F10 melanoma cells after exposure to water, ethanol–water (30:70 (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) and ethanol–water (70:30 (<span class="html-italic">v</span>/<span class="html-italic">v</span>)) extracts from three varieties of <span class="html-italic">C. mas</span> L. Extracts were obtained from varieties with yellow, red and dark ruby-red fruit. Analyses were performed for extracts at dilutions of 1:100, 1:20 and 1:10 (<span class="html-italic">v</span>/<span class="html-italic">v</span>). The research used yellow fruit extracts (water (YW), water–ethanol 30:70 (YE30), water–ethanol 70:30 (YE70)), red fruit extracts (water (RW), water–ethanol 30:70 (RE30), water–ethanol 70:30 (RE70)) and dark ruby-red extracts (water (DRW), water–ethanol 30:70 (DRE30), water–ethanol 70:30 (DRE70)). Data represent mean ± SD of three independent experiments in which each dilution of individual extracts was tested in triplicate. **** <span class="html-italic">p</span> &lt; 0.0001.</p>
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12 pages, 2901 KiB  
Article
Development of a Bioactive Titanium Surface via Alkalinization and Naringenin Coating for Peri-Implant Repair: In Vitro Study
by Isabela Massaro Ribeiro, Lais Medeiros Cardoso, Taisa Nogueira Pansani, Ana Carolina Chagas, Carlos Alberto de Souza Costa and Fernanda Gonçalves Basso
Coatings 2024, 14(10), 1303; https://doi.org/10.3390/coatings14101303 - 12 Oct 2024
Viewed by 292
Abstract
This study assessed the effects of titanium (Ti) surface modification with sodium hydroxide (NaOH) associated or not with Naringenin (NA) citrus flavonoid-coating on osteoblastic-like cells (Ob) metabolism. Ti discs were submitted to alkalinization by NaOH solution (5 M, 60 °C) for 24 h; [...] Read more.
This study assessed the effects of titanium (Ti) surface modification with sodium hydroxide (NaOH) associated or not with Naringenin (NA) citrus flavonoid-coating on osteoblastic-like cells (Ob) metabolism. Ti discs were submitted to alkalinization by NaOH solution (5 M, 60 °C) for 24 h; then, the discs were impregnated or not with 100 µg/mL of NA and dried for 1 h at room temperature. The chemical composition, surface topography, and NA release were evaluated. For the biological assays, the discs were placed on 24-well cell culture plates and Ob (Saos-2; ATCC HTB-85) was seeded onto the discs. After different periods, cell adhesion and viability, alkaline phosphatase activity (ALP), and mineralized nodules deposition (MND) were assessed. In addition, cells stimulated with tumor necrosis factor-alpha (TNF-α) were submitted to matrix metalloproteinase (MMP)-2 synthesis and ALP gene expression assessment. Since data presented normal distribution and homogeneity (Shapiro-Wilk; Levene), Student’s t-test or one-way ANOVA/post-hoc tests were selected for data analysis (α = 0.05). Higher roughness was observed on Ti discs submitted to NaOH treatment, while the chemical and NA release evaluations indicated the successful adsorption of NA to alkali-treated Ti surface. Higher cell adhesion, cell viability (after 7 days of culture), ALP activity, and MND were observed on Ti NaOH coated with NA compared to the control group (Ti NaOH) (p < 0.05). Moreover, NA coating also promoted decreased MMP-2 synthesis and increased ALP gene expression in the presence of the inflammatory stimulus TNF-α (p < 0.05). The modification of Ti disks with NaOH associated with NA-coating enhanced bone cell metabolism, suggesting that this type of surface modification has a promising potential to accelerate bone repair and formation around dental implants. Full article
(This article belongs to the Special Issue Synthesis and Applications of Bioactive Coatings)
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Figure 1

Figure 1
<p>Qualitative analysis of surface topography of Ti and NaOH-treated Ti disks by scanning electron microscopy (SEM) and ImageJ software and quantitative analysis of surface roughness (Ra) for both surfaces.</p>
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<p>(<b>A</b>) Displays Fourier Transform Infrared (FTIR) spectra for Naringenin (NA) powder and Titanium (Ti) discs submitted to alkali treatment (NaOH) and coated or not with NA. (<b>B</b>) NA release from the alkalinized Ti-discs coated with NA over time after incubation at 37 °C in ultrapure water (<span class="html-italic">n</span> = 6). Data points are mean values and error bars denote 95% confidence intervals (α = 5%).</p>
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<p>(<b>A</b>) Cell viability (% of control Ti NaOH group) after 24 h, 48 h, and 7 days of cell culture on modified Ti surfaces (<span class="html-italic">n</span> = 6). Columns represent mean values and error bars represent standard deviations. Statistical notation is indicated by * (Student <span class="html-italic">t</span>-test, <span class="html-italic">p</span> &lt; 0.05). (<b>B</b>) Representative images of fluorescence microscopy for cell adhesion and spreading on modified Ti surfaces after 24 h and 48 h of culture (<span class="html-italic">n</span> = 4). Cell nuclei are stained in blue (Hoechst) and actin filaments are stained in red (ActinRed). Scale bar: 250 μm—10× magnification; 125 μm—20× magnification).</p>
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<p>(<b>A</b>) ALP activity and (<b>B</b>) mineralized nodule formation (% of control Ti NaOH group) after 7 days of cell culture on modified Ti surfaces (<span class="html-italic">n</span> = 6). Columns represent mean values and error bars represent standard deviations. Statistical notation is indicated by * (Student <span class="html-italic">t</span>-test, <span class="html-italic">p</span> &lt; 0.05).</p>
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<p>(<b>A</b>) Synthesis of MMP-2 (% of control Ti NaOH group) and (<b>B</b>) ALP gene expression (mRNA fold change) by osteoblasts exposed (+TNF) or not (−TNF) to tumor necrosis factor alpha (TNF-α) (<span class="html-italic">n</span> = 6). Columns represent mean values and error bars represent standard deviations. Groups identified by different symbols (*, ** or ***) are statistically different from each other (one-way ANOVA, followed by Tukey’s post-hoc test, <span class="html-italic">p</span> &lt; 0.05).</p>
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15 pages, 3373 KiB  
Article
Osteoblast Response to Widely Ranged Texturing Conditions Obtained through High Power Laser Beams on Ti Surfaces
by Federico Alessandro Ruffinatti, Tullio Genova, Ilaria Roato, Martina Perin, Giorgia Chinigò, Riccardo Pedraza, Olivio Della Bella, Francesca Motta, Elisa Aimo Boot, Domenico D’Angelo, Giorgio Gatti, Giorgia Scarpellino, Luca Munaron and Federico Mussano
J. Funct. Biomater. 2024, 15(10), 303; https://doi.org/10.3390/jfb15100303 - 12 Oct 2024
Viewed by 440
Abstract
Titanium and titanium alloys are the prevailing dental implant materials owing to their favorable mechanical properties and biocompatibility, but how roughness dictates the biological response is still a matter of debate. In this study, laser texturing was used to generate eight paradigmatic roughened [...] Read more.
Titanium and titanium alloys are the prevailing dental implant materials owing to their favorable mechanical properties and biocompatibility, but how roughness dictates the biological response is still a matter of debate. In this study, laser texturing was used to generate eight paradigmatic roughened surfaces, with the aim of studying the early biological response elicited on MC3T3-E1 pre-osteoblasts. Prior to cell tests, the samples underwent SEM analysis, optical profilometry, protein adsorption assay, and optical contact angle measurement with water and diiodomethane to determine surface free energy. While all the specimens proved to be biocompatible, supporting similar cell viability at 1, 2, and 3 days, surface roughness could impact significantly on cell adhesion. Factorial analysis and linear regression showed, in a robust and unprecedented way, that an isotropic distribution of deep and closely spaced valleys provides the best condition for cell adhesion, to which both protein adsorption and surface free energy were highly correlated. Overall, here the authors provide, for the first time, a thorough investigation of the relationship between roughness parameters and osteoblast adhesion that may be applied to design and produce new tailored interfaces for implant materials. Full article
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Figure 1

Figure 1
<p>Scheme of the factorial experimental design followed for surface modification of titanium disks. Throughout this work, titanium samples are identified using an alphanumeric ID encoding these three features, in the following order: inter-pit distance (25 or 50 μm), pattern type (A for aligned or R for random), and pit depth (6 or 18 μm).</p>
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<p>SEM images showing the surface topography of the titanium samples at high magnification (1000×). Each surface is named after the alphanumeric ID defined above, which consists of the values of pit spacing (25 or 50 μm), pattern type (Aligned or Random), and pit depth (6 or 18 μm), respectively.</p>
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<p>Protein adsorption and cell adhesion measures. (<b>A</b>) Data from three independent protein adsorption assays are represented as mean ± SE of the amount of adsorbed protein per volume (µg/mL), for each different titanium surface. (<b>B</b>) Data from three independent cell adhesion assays are represented as mean ± SE of the number of counted cells per field of view, for each different titanium surface.</p>
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<p>Cell viability at 24 h and 72 h. Data from three independent experiments are presented as mean ± SE for each different titanium surface (RLU = relative light units).</p>
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<p>Factorial analysis. Main effects on cell adhesion of: (<b>A</b>) inter-pit distance (<span class="html-italic">p</span>-value = 2.55 × 10<sup>−2</sup>), (<b>B</b>) pattern type (<span class="html-italic">p</span>-value = 3.32 × 10<sup>−2</sup>), and (<b>C</b>) pit depth (<span class="html-italic">p</span>-value = 9.67 × 10<sup>−4</sup>). (<b>D</b>) A 3D representation of the global regression model. Light-blue and violet planes represent the linear model equation evaluated for aligned and random pattern, respectively.</p>
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<p>Graphical representation of the plane model. Two different views of the same plane given by Equation (1) linking cell adhesion data points (yellow dots) to the surface roughness properties of the titanium disks. Multiple linear regression analysis returned only two non-redundant and statistically significant coefficients out of the eleven initial roughness descriptors (<math display="inline"><semantics> <mrow> <msub> <mrow> <mi>β</mi> </mrow> <mrow> <mi>S</mi> <mi>t</mi> <mi>r</mi> </mrow> </msub> </mrow> </semantics></math> coefficient: <span class="html-italic">p</span>-value = 7.10 × 10<sup>−3</sup>; <math display="inline"><semantics> <mrow> <msub> <mrow> <mi>β</mi> </mrow> <mrow> <mi>S</mi> <mi>k</mi> <mi>u</mi> </mrow> </msub> </mrow> </semantics></math> coefficient: <span class="html-italic">p</span>-value = 3.57 × 10<sup>−2</sup>.</p>
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<p>Correlation analysis between cell adhesion, protein adsorption, and SFE. (<b>A</b>) Scatterplot and correlation analysis between protein adsorption and cell adhesion data. The numerical values of the correlation coefficients shown in the graph refer to the overall correlation analysis between the two datasets (Pearson correlation coefficient <math display="inline"><semantics> <mrow> <msub> <mrow> <mi>ρ</mi> </mrow> <mrow> <mi>P</mi> </mrow> </msub> <mo>=</mo> <mn>0.82</mn> </mrow> </semantics></math>, <math display="inline"><semantics> <mrow> <mi>p</mi> <mrow> <mtext>-</mtext> <mi>value</mi> </mrow> <mo>=</mo> <mn>0.012</mn> </mrow> </semantics></math>; Spearman correlation coefficient <math display="inline"><semantics> <mrow> <msub> <mrow> <mi>ρ</mi> </mrow> <mrow> <mi>S</mi> </mrow> </msub> <mo>=</mo> <mn>0.93</mn> </mrow> </semantics></math>, <math display="inline"><semantics> <mrow> <mi>p</mi> <mrow> <mtext>-</mtext> <mi>value</mi> </mrow> <mo>=</mo> <mn>0.002</mn> </mrow> </semantics></math>), while two independent loess curves were used to highlight the nearly deterministic relationship between protein adsorption and cell adhesion within the single subset of surfaces with 50 μm inter-pit distance (cyan curve) and 25 μm spacing (magenta curve). (<b>B</b>) Scatterplot and correlation analysis between SFE and cell adhesion data. The best fitting line (in blue) and the 95% confidence interval (shaded in gray) are superimposed on the data points (Pearson correlation coefficient <math display="inline"><semantics> <mrow> <msub> <mrow> <mi>ρ</mi> </mrow> <mrow> <mi>P</mi> </mrow> </msub> <mo>=</mo> <mn>0.88</mn> </mrow> </semantics></math>, <math display="inline"><semantics> <mrow> <mi>p</mi> <mrow> <mtext>-</mtext> <mi>value</mi> </mrow> <mo>=</mo> <mn>0.009</mn> </mrow> </semantics></math>; Spearman correlation coefficient <math display="inline"><semantics> <mrow> <msub> <mrow> <mi>ρ</mi> </mrow> <mrow> <mi>S</mi> </mrow> </msub> <mo>=</mo> <mn>0.86</mn> </mrow> </semantics></math>, <math display="inline"><semantics> <mrow> <mi>p</mi> <mrow> <mtext>-</mtext> <mi>value</mi> </mrow> <mo>=</mo> <mn>0.024</mn> </mrow> </semantics></math>).</p>
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18 pages, 9131 KiB  
Article
Protective Role of Eicosapentaenoic and Docosahexaenoic and Their N-Ethanolamide Derivatives in Olfactory Glial Cells Affected by Lipopolysaccharide-Induced Neuroinflammation
by Rosalia Pellitteri, Valentina La Cognata, Cristina Russo, Angela Patti and Claudia Sanfilippo
Molecules 2024, 29(20), 4821; https://doi.org/10.3390/molecules29204821 - 11 Oct 2024
Viewed by 317
Abstract
Neuroinflammation is a symptom of different neurodegenerative diseases, and growing interest is directed towards active drug development for the reduction of its negative effects. The anti-inflammatory activity of polyunsaturated fatty acids, eicosapentaenoic (EPA), docosahexaenoic (DHA), and their amide derivatives was largely investigated on [...] Read more.
Neuroinflammation is a symptom of different neurodegenerative diseases, and growing interest is directed towards active drug development for the reduction of its negative effects. The anti-inflammatory activity of polyunsaturated fatty acids, eicosapentaenoic (EPA), docosahexaenoic (DHA), and their amide derivatives was largely investigated on some neural cells. Herein, we aimed to elucidate the protective role of both EPA and DHA and the corresponding N-ethanolamides EPA-EA and DHA-EA on neonatal mouse Olfactory Ensheathing Cells (OECs) after exposition to lipopolysaccharide (LPS)-induced neuroinflammation. To verify their anti-inflammatory effect and cell morphological features on OECs, the expression of IL-10 cytokine, and cytoskeletal proteins (vimentin and GFAP) was evaluated by immunocytochemical procedures. In addition, MTT assays, TUNEL, and mitochondrial health tests were carried out to assess their protective effects on OEC viability. Our results highlight a reduction in GFAP and vimentin expression in OECs exposed to LPS and treated with EPA or DHA or EPA-EA or DHA-EA in comparison with OECs exposed to LPS alone. We observed a protective role of EPA and DHA on cell morphology, while the amides EPA-EA and DHA-EA mainly exerted a superior anti-inflammatory effect compared to free acids. Full article
(This article belongs to the Section Bioactive Lipids)
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Graphical abstract

Graphical abstract
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<p>Immunofluorescent staining for S-100 protein expression in primary mouse OECs. Magnification: 20×. Scale bar: 10 µm.</p>
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<p>Morphological analysis of OECs by phase contrast microscopy. Representative fields of OECs, without (<b>A</b>) and with (<b>B</b>) LPS exposure, were treated with EPA, DHA, EPA-EA, and DHA-EA at different concentrations (0.1 µM and 0.5 µM) for 24 h. Magnification: 10×. Scale bars: 50 μm.</p>
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<p>Percentage of viable OECs after treatment with EPA, DHA, EPA-EA, and DHA-EA. OECs without or with LPS stress were treated with EPA, DHA, EPA-EA, and DHA-EA at different concentrations (0.1 µM and 0.5 µM) for 24 h. One-way ANOVA experimental groups vs. CTR, F (18, 152) = 37.60. One-way ANOVA experimental groups vs. LPS, F (16, 136) = 28.58. Dunnett’s multiple comparison test: **** <span class="html-italic">p</span> &lt; 0.0001 vs. CTR; #### <span class="html-italic">p</span> &lt; 0.0001 vs. LPS.</p>
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<p>Dead cell detection by TUNEL assay. The test was carried out on OECs without (<b>A</b>) and with (<b>B</b>) LPS exposure and treated with EPA, DHA, EPA-EA, and DHA-EA. Photomicrographs are representative of randomly selected fields and were scanned by a Nikon Ti Eclipse inverted microscope. Scale bar 10 μm. (<b>C</b>) Data were taken from at least three independent experiments. TUNEL-positive cells were calculated as described in <a href="#sec4-molecules-29-04821" class="html-sec">Section 4</a> and are reported as mean ± SEM. One-way ANOVA experimental groups vs. CTRL condition F (10, 22) = 9.913; <span class="html-italic">p</span> &lt; 0.0001; One-way ANOVA experimental groups vs. LPS treatment F (8, 18) = 6.616; <span class="html-italic">p</span> = 0.0004; Dunnett’s multiple comparison tests: ** <span class="html-italic">p</span> &lt; 0.01 and *** <span class="html-italic">p</span> &lt; 0.001 vs. Ctrl; # <span class="html-italic">p</span> &lt; 0.05, ## <span class="html-italic">p</span> &lt; 0.01 and ### <span class="html-italic">p</span> &lt; 0.001 vs. LPS.</p>
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<p>Mitotoxicity and cytotoxicity levels in OECs exposed to different treatments. The test was carried out on OECs without (<b>A</b>) and with (<b>B</b>) LPS exposure and treated with EPA, DHA, EPA-EA, and DHA-EA. Representative images were taken from randomly selected fields of slides scanned by a Nikon Ti Eclipse inverted microscope (scale bar 10 μm). Mean Fluorescent Intensity (MFI) values (mean ± SEM) of the ratio MitoHealth (<b>C</b>) and DeadGreen (<b>D</b>) were calculated as described in the Material and Methods Section. Values are reported as mean ± SEM. For Mitohealth staining, One-way ANOVA experimental groups vs. CTRL condition F (10, 99) = 38.81; <span class="html-italic">p</span> &lt; 0.0001; One-way ANOVA experimental groups vs. LPS treatment F (8, 81) = 14.69; <span class="html-italic">p</span> &lt; 0.0001. For DeadGreen staining, One-way ANOVA experimental groups vs. CTRL condition F (10, 99) = 137.8; <span class="html-italic">p</span> &lt; 0.0001; One-way ANOVA experimental groups vs. LPS treatment F (8, 81) = 49.71; <span class="html-italic">p</span> &lt; 0.0001. Dunnett’s multiple comparison test: *** <span class="html-italic">p</span> &lt; 0.001 vs. CTRL # <span class="html-italic">p</span> &lt; 0.05, ## <span class="html-italic">p</span> &lt; 0.01, and ### <span class="html-italic">p</span> &lt; 0.001 vs. LPS.</p>
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<p>Immunocytochemistry for IL-10 in OECs exposed to different treatments. The test was carried out on OECs without (<b>A</b>) and with (<b>B</b>) LPS exposure and treated with EPA, DHA, EPA-EA, and DHA-EA. Immunostained samples were analyzed with a Zeiss fluorescence microscope, and images were captured with the Axiovision imaging system. (<b>C</b>) Fluorescence quantification data for IL-10 in each OEC cultured in different conditions. Bars represent CTCF mean value ± SD, obtained from at least three independent experiments. One-way ANOVA experimental groups vs. CTR, F (18,114) = 292.8. One-way ANOVA experimental groups vs. LPS, F (16, 102) = 278.5. Dunnett’s multiple comparison test: ** <span class="html-italic">p</span> &lt; 0.01; *** <span class="html-italic">p</span> &lt; 0.001; **** <span class="html-italic">p</span> &lt; 0.0001 vs. CTR; #### <span class="html-italic">p</span> &lt; 0.0001 vs. LPS. Scale bar 20 μm.</p>
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<p>Immunocytochemistry for GFAP in OECs exposed to different treatments. The test was carried out on OECs without (<b>A</b>) and with (<b>B</b>) LPS exposure and treated with EPA, DHA, EPA-EA, and DHA-EA. (<b>C</b>) Fluorescence quantification data for GFAP in each OEC treatment with EPA, DHA, EPA-EA, and DHA-EA at different concentrations (0.1 µm and 0.5 µm). Bars represent CTCF mean value ± SD, obtained from at least three independent experiments. One-way ANOVA experimental groups vs. CTR, F (18, 133) = 101.5. One-way ANOVA experimental groups vs. LPS, F (16, 119) = 116.5. Dunnett’s multiple comparison test: ** <span class="html-italic">p</span> &lt; 0.01; *** <span class="html-italic">p</span> &lt; 0.001; **** <span class="html-italic">p</span> &lt; 0.0001 vs. CTR; #### <span class="html-italic">p</span> &lt; 0.0001 vs. LPS. Scale bar 20 μm.</p>
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<p>Immunocytochemistry for vimentin in OECs exposed to different treatments. The test was carried out on OECs without (<b>A</b>) and with (<b>B</b>) LPS exposure and treated with EPA, DHA, EPA-EA, and DHA-EA. (<b>C</b>) Fluorescence quantification data for vimentin in each OEC treatment with EPA, DHA, EPA-EA, and DHA-EA at different concentrations (0.1 µm and 0.5 µm). Bars represent CTCF mean value ± SD, obtained from at least three independent experiments. One-way ANOVA experimental groups vs. CTR, F (18, 95) = 375.5. One-way ANOVA experimental groups vs. LPS, F (16, 85) = 396.1. Dunnett’s multiple comparison test: *** <span class="html-italic">p</span> &lt; 0.001; **** <span class="html-italic">p</span> &lt; 0.0001 vs. CTR; #### <span class="html-italic">p</span> &lt; 0.0001 vs. LPS. Scale bar 20 μm.</p>
Full article ">Scheme 1
<p>Chemical structures of the compounds tested in this study and their synthetic precursors.</p>
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25 pages, 9863 KiB  
Article
Targeting PARP-1 and DNA Damage Response Defects in Colorectal Cancer Chemotherapy with Established and Novel PARP Inhibitors
by Philipp Demuth, Lea Thibol, Anna Lemsch, Felix Potlitz, Lukas Schulig, Christoph Grathwol, Georg Manolikakes, Dennis Schade, Vassilis Roukos, Andreas Link and Jörg Fahrer
Cancers 2024, 16(20), 3441; https://doi.org/10.3390/cancers16203441 - 10 Oct 2024
Viewed by 512
Abstract
The DNA repair protein PARP-1 emerged as a valuable target in the treatment of tumor entities with deficiencies of BRCA1/2, such as breast cancer. More recently, the application of PARP inhibitors (PARPi) such as olaparib has been expanded to other cancer entities [...] Read more.
The DNA repair protein PARP-1 emerged as a valuable target in the treatment of tumor entities with deficiencies of BRCA1/2, such as breast cancer. More recently, the application of PARP inhibitors (PARPi) such as olaparib has been expanded to other cancer entities including colorectal cancer (CRC). We previously demonstrated that PARP-1 is overexpressed in human CRC and promotes CRC progression in a mouse model. However, acquired resistance to PARPi and cytotoxicity-mediated adverse effects limit their clinical applicability. Here, we detailed the role of PARP-1 as a therapeutic target in CRC and studied the efficacy of novel PARPi compounds in wildtype (WT) and DNA repair-deficient CRC cell lines together with the chemotherapeutics irinotecan (IT), 5-fluorouracil (5-FU), and oxaliplatin (OXA). Based on the ComPlat molecule archive, we identified novel PARPi candidates by molecular docking experiments in silico, which were then confirmed by in vitro PARP activity measurements. Two promising candidates (X17613 and X17618) also showed potent PARP-1 inhibition in a CRC cell-based assay. In contrast to olaparib, the PARPi candidates caused no PARP-1 trapping and, consistently, were not or only weakly cytotoxic in WT CRC cells and their BRCA2- or ATR-deficient counterparts. Importantly, both PARPi candidates did not affect the viability of nonmalignant human colonic epithelial cells. While both olaparib and veliparib increased the sensitivity of WT CRC cells towards IT, no synergism was observed for X17613 and X17618. Finally, we provided evidence that all PARPi (olaparib > veliparib > X17613 > X17618) synergize with chemotherapeutic drugs (IT > OXA) in a BRCA2-dependent manner in CRC cells, whereas ATR deficiency had only a minor impact. Collectively, our study identified novel lead structures with potent PARP-1 inhibitory activity in CRC cells but low cytotoxicity due to the lack of PARP-1 trapping, which synergized with IT in homologous recombination deficiency. Full article
(This article belongs to the Special Issue Cancer Chemotherapy: Combination with Inhibitors (2nd Edition))
Show Figures

Figure 1

Figure 1
<p>(<b>A</b>) Binding modes of veliparib (<b>1</b>), PDB: 7AAC), olaparib ((<b>2</b>), PDB: 7AAD), and selected compounds from virtual screening ((<b>3</b>–<b>9</b>), PDB: 4PJT) to PARP-1. The binding to either G863 or S904, as also found for veliparib, was used as a constraint in docking. All active compounds are able to form this bond and adopt a similar binding mode. Through the indole NH, there is an interaction with E988 by a bridging water molecule. No preference between the binding modes of the S- (<b>4</b>–<b>6</b>) or R-enantiomers (<b>7</b>–<b>9</b>) is observed, while the scoring values also differ only slightly. (<b>B</b>) Chemical structure of the most active compounds X17613, X17618, X17620, and X17621, according to in vitro screening and the two established PARPi veliparib and olaparib.</p>
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<p>(<b>A</b>) Concentration–response curves of four potential PARP-1 inhibitors with the highest activity in the PARP-1 screening assay kit. All concentrations were tested in duplicates. IC<sub>50</sub> values were derived using a nonlinear regression model in GraphPad Prism 9 (<span class="html-italic">n</span> = 2). (<b>B</b>) Investigation of PARP inhibition by X17613, X17618, X17620, and X17621 in HCT116 cells. Cells were challenged with 1 mM H<sub>2</sub>O<sub>2</sub> for 5 min and pretreated or not with the indicated compounds for 2 h. PAR synthesis was identified by confocal IF microscopy using the PAR 10H antibody. The signal intensity of five images per concentration was evaluated by ImageJ (<span class="html-italic">n</span> ≥ 3). (<b>C</b>) Representative confocal microscopy images at 100× magnification after PAR staining in HCT116 cells treated with the indicated concentrations of X17613 for 2 h with or without subsequent PARP activation by H<sub>2</sub>O<sub>2</sub> treatment for 5 min. Scale bar: 100 µm. (<b>D</b>) Confocal microscopy images at 630× magnification after pan-PAR staining in HCT116 cells treated according to (<b>C</b>). Scale bar: 20 µm. Data are presented as mean +/− SEM. * <span class="html-italic">p</span> &lt; 0.01, ** <span class="html-italic">p</span> &lt; 0.01; <span class="html-italic">t</span>-test.</p>
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<p>(<b>A</b>,<b>B</b>) Analysis of PARP-1 trapping in HCT116 and Caco-2 cells. Immunoblot detection of PARP-1 after pre-treatment with X17613, X17618, and olaparib followed by MMS exposure for 1 h and chromatin isolation. The cytosolic marker Hsp90 and the chromatin marker Histone H3 served as respective loading controls. Representative Western blot images and densitometric evaluation are shown (<span class="html-italic">n</span> = 3). Data are shown as mean + SEM. (<b>C</b>) Cell viability determined by the resazurin reduction assay (RRA) in HCT116 PARP-1<sup>−/−</sup> and HCT116 PARP-1<sup>+/+</sup> cells after PARPi treatment for 72 h. A nonlinear regression curve fit was conducted using GraphPad Prism 9 (<span class="html-italic">n</span> ≥ 3). (<b>D</b>) Viability in Caco-2 cells after exposure to PARPi as indicated. (<span class="html-italic">n</span> = 3). All data are shown as mean +/− SEM.</p>
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<p>(<b>A</b>) Toxicity of PARPi in HCT116 cells depending on BRCA2 status. HCT116 WT and HCT116 BRCA2<sup>−/−</sup> cells were incubated with PARPi for 72 h and viability was assessed using the resazurin reduction assay (RRA). Nonlinear regression curve fit was conducted using GraphPad Prism 9 (<span class="html-italic">n</span> ≥ 3). (<b>B</b>) Toxicity of PARPi in DLD-1 cells depending on ATR status. DLD-1 WT and DLD-1 ATR<sup>s/s</sup> cells were incubated with PARPi for 72 h and viability was assessed using the RRA. Nonlinear regression curve fit was conducted using GraphPad Prism 9 (<span class="html-italic">n</span> ≥ 3). Data are depicted as mean +/− SEM.</p>
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<p>(<b>A</b>) Viability in HCT116 PARP-1<sup>−/−</sup> and HCT116 PARP-1<sup>+/+</sup> cells after treatment with PARPi olaparib or veliparib in combination with chemotherapeutic drugs irinotecan (IT, 0.5 µM), 5-fluorouracil (5-FU, 0.25 µM), and oxaliplatin (OXA, 0.5 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). (<b>B</b>) Viability in HCT116 PARP-1<sup>−/−</sup> and HCT116 PARP-1<sup>+/+</sup> cells after treatment with PARPi X17613 in combination with chemotherapeutic drugs for 72 h. Data (<span class="html-italic">n</span> ≥ 3) are given as mean +/− SEM. (<b>C</b>,<b>D</b>) Viability in Caco-2 cells after treatment with PARPi in combination with chemotherapeutic drugs irinotecan (IT, 10 µM), 5-fluorouracil (5-FU, 5 µM), and oxaliplatin (OXA, 1 µM). Data (<span class="html-italic">n</span> = 3) are shown as mean +/− SEM. ns: <span class="html-italic">p</span> &gt; 0.05, * <span class="html-italic">p</span> &lt; 0.01, ** <span class="html-italic">p</span> &lt; 0.01, *** <span class="html-italic">p</span> &lt; 0.001, **** <span class="html-italic">p</span> &lt; 0.0001; <span class="html-italic">t</span>-test.</p>
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<p>(<b>A</b>) Viability in HCT116 WT and HCT116 BRCA2<sup>−/−</sup> cells after treatment with PARPi olaparib or veliparib in combination with chemotherapeutic drugs (IT, 0.25 µM), 5-fluorouracil (5-FU, 0.1 µM), and oxaliplatin (OXA, 0.25 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). (<b>B</b>) Viability in HCT116 WT and HCT116 BRCA2<sup>−/−</sup> cells after treatment with PARPi X17613 in combination with chemotherapeutic drugs (IT, 0.25 µM), 5-fluorouracil (5-FU, 0.1 µM), and oxaliplatin (OXA, 0.25 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). (<b>C</b>) Representative brightfield microscopic images at 20X magnification of HCT116 WT and HCT116 BRCA2<sup>−/−</sup> cells after treatment with X17613 (50 µM), IT (0.25 µM), or a combination of both for 24 h. (<b>D</b>,<b>E</b>) γH2AX formation in HCT116 WT and BRCA2<sup>−/−</sup> cells after treatment as described in (<b>C</b>). Representative Western blot images and densitometric evaluation are shown (<span class="html-italic">n</span> = 4). Hsp90 served as loading control. All data are given as mean + SEM. ns: <span class="html-italic">p</span> &gt; 0.05, * <span class="html-italic">p</span> &lt; 0.01, ** <span class="html-italic">p</span> &lt; 0.01; <span class="html-italic">t</span>-test.</p>
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<p>(<b>A</b>) Binding modes of selected compounds from virtual screening ((<b>1</b>–<b>4</b>), PDB: 4PJT) to PARP-1. The binding to either G863 or S904 was used as a constraint in docking. All active compounds are able to form this bond and adopt a similar binding mode (compare <a href="#cancers-16-03441-f001" class="html-fig">Figure 1</a>A). The three compounds X17611, X17610, and X17608 (<b>1</b>–<b>3</b>) serve as a verification of the binding mode since the constraints can only be fulfilled with significant losses in the binding free energy due to unfavorable inter- and intramolecular interactions. X17616 (<b>4</b>) is similar to X17618 (<a href="#cancers-16-03441-f001" class="html-fig">Figure 1</a>A), but cannot form a potential hydrogen bond due to the lack of the amide function. (<b>B</b>) Chemical structure of the compounds X17611, X17610, X17608 and X17616.</p>
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<p>(<b>A</b>,<b>B</b>) Concentration response curve of established PARP inhibitors veliparib and olaparib (<b>A</b>) and eight potential PARP inhibitors (<b>B</b>) with low or no activity in the PARP-1 screening assay kit. All concentrations were tested in duplicates. IC<sub>50</sub> values were derived using a nonlinear regression model in GraphPad Prism 9 (<span class="html-italic">n</span> = 2).</p>
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<p>(<b>A</b>) Concentration-response curves of 4 potential PARP-1 inhibitors (X17613, X17618, X17620 and X17621) assessed in HCT116 cells as described in <a href="#cancers-16-03441-f002" class="html-fig">Figure 2</a>B (<span class="html-italic">n</span> = 3). IC<sub>50</sub> values were derived using a nonlinear regression model in GraphPad Prism 9. (<b>B</b>) Representative confocal microscopy images at 100× magnification after PAR staining in HCT116 cells treated with the indicated concentrations of X17618 for 2 h with or without subsequent PARP induction by H<sub>2</sub>O<sub>2</sub> treatment for 5 min. Scale bar: 100 µm. (<b>C</b>) Confocal microscopy images at 630× magnification after pan-PAR staining in HCT116 treated as described in B. Scale bar: 20 µm.</p>
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<p>(<b>A</b>) Western Blot analysis of PARP-1, BRCA2 and ATR expression in HCT116 WT, HCT116 BRCA2<sup>−/−</sup>, HCT116 PARP-1<sup>+/+</sup>, HCT116 PARP-1<sup>−/−</sup>, DLD-1 WT and DLD-1 ATR<sup>−/−</sup> cells. HSP90 served as loading control. (<b>B</b>) Toxicity of PARPi in human colonic epithelial cells (HCEC). Cells were incubated with PARPi for 72 h and viability was assessed using the RRA. Nonlinear regression curve fit was conducted using GraphPad Prism 9 (<span class="html-italic">n</span> ≥ 3).</p>
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<p>Cell viability of (<b>A</b>) HCT116 WT and HCT116 BRCA2<sup>−/−</sup>, (<b>B</b>) HCT116 PARP-1<sup>+/+</sup> and HCT116 PARP-1<sup>−/−</sup>, (<b>C</b>) DLD-1 WT and DLD-1 ATR<sup>s/s</sup>, (<b>D</b>) Caco-2 cells and (<b>E</b>) HCEC after monotreatment with cytostatic drugs IT, 5-FU and OXA for 72 h. Nonlinear regression curve fit was conducted using GraphPad Prism 9 (<span class="html-italic">n</span> ≥ 3).</p>
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<p>(<b>A</b>) Viability of HCT116 PARP-1<sup>−/−</sup> and HCT116 PARP-1<sup>+/+</sup> cells after treatment with PARPi X17618 in combination with chemotherapeutic drugs irinotecan (IT, 0.5 µM), 5-fluorouracil (5-FU, 0.25 µM) and oxaliplatin (OXA, 0.5 µM) for 72 h (<span class="html-italic">n</span> ≥ 3) (<b>B</b>) Viability of HCT116 WT and HCT116 BRCA2<sup>−/−</sup> cells after treatment with PARPi X17618 in combination with chemotherapeutic drugs irinotecan (IT, 0.25 µM), 5-fluorouracil (5-FU, 0.1 µM) and oxaliplatin (OXA, 0.25 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). (<b>C</b>) γH2AX formation in HCT116 WT and BRCA2<sup>−/−</sup> cells after treatment with X17618 (50 µM), IT (0.25 µM) or a combination of both for 24 h. Representative Western blot images and densitometric evaluation are shown (<span class="html-italic">n</span> = 4). All data are presented as mean +/− SEM. ns: <span class="html-italic">p</span> &gt; 0.05, * <span class="html-italic">p</span> &lt; 0.01, ** <span class="html-italic">p</span> &lt; 0.01, *** <span class="html-italic">p</span> &lt; 0.001, **** <span class="html-italic">p</span> &lt; 0.0001; <span class="html-italic">t</span>-test.</p>
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<p>(<b>A</b>) Viability of DLD-1 WT and DLD-1 ATR<sup>s/s</sup> cells after treatment with PARPi olaparib or veliparib in combination with chemotherapeutic drugs (IT, 2.5 µM), 5-fluorouracil (5-FU, 0.1 µM) and oxaliplatin (OXA, 5 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). (<b>B</b>) Viability of DLD-1 WT and DLD-1 ATR<sup>s/s</sup> cells after treatment with PARPi X17613 and X17618 in combination with chemotherapeutic drugs (IT, 2.5 µM), 5-fluorouracil (5-FU, 0.1 µM) and oxaliplatin (OXA, 5 µM) for 72 h (<span class="html-italic">n</span> ≥ 3). Data are shown as mean +/− SEM. * <span class="html-italic">p</span> &lt; 0.01, ** <span class="html-italic">p</span> &lt; 0.01, *** <span class="html-italic">p</span> &lt; 0.001, **** <span class="html-italic">p</span> &lt; 0.0001; <span class="html-italic">t</span>-test.</p>
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