Figure 1. hMSC-to-MF Activation Reduces Clonogenicity and Lineage Differentiation Potential (A and B) hMSCs were cultured with and without TGF-β1 (2 ng/ml) for 4 days to assess MF activation by staining for α-SMA (red) and stress fibers (F-actin, green), for 10 days to assess CFU-F formation, and 14 days to assess adipogenesis (oil-red-O-positive lipids) and osteogenesis (Alizarin Red S-positive mineralized nodules) in respective induction media (+TGF-β1). (B) MF activation was quantified by assessing the percentage of α-SMA expressing hMSCs. The contraction was assessed by growing hMSC on deformable wrinkling substrates (white lines in immunofluorescence image overlay) and quantifying the total wrinkle area normalized by cell numbers. Clonogenicity was assessed by counting the number of colonies formed after 10 days. (C–G) hMSCs were cultured on FN-coated silicone substrates, mimicking fibrosis-rigid (65 kPa), wound-stiff (26 kPa), and normal-tissue-soft (3 kPa) environments. (D) After 5 days, cells were analyzed by western blotting for α-SMA and vimentin (loading control) and (E) immunofluorescence for α-SMA (red) and ED-A FN (green). The scale bar represents 50 μm. (F) MF activation was quantified as percentage of α-SMA stress fiber positive cells. (G) Clonogenicity was quantified in CFU-F assays on differently stiff substrates. All of the graphs show averages ± SD from at least five independent experiments (*p ≤ 0.05, **p ≤ 0.005, and ***p ≤ 0.0005 using ANOVA followed by a post hoc Tukey’s multiple comparison test). See also Figure S1.

MF Activation Results in Reduced Clonogenicity and Differentiation Potential of hMSCsIndependently of MSC origin, MF activation occurs spontaneously in standard cell culture on rigid tissue culture plastic in serum-containing media. Cultured hMSCs derived from adipose tissue, umbilical cord perivasculature, and bone marrow all developed MF characteristics, including α-SMA-positive stress fibers, extradomain-A fibronectin (ED-A FN) in the ECM, and high contractile activity, which were enhanced by TGF-β1 (Figure S1). To test whether MF activation affected stem cell features, we focused on bone-marrow derived hMSCs (Figure 1A). In standard culture, 17% ± 4.2% of hMSCs expressed α-SMA in F-actin-positive stress fibers, which increased to 32% ± 5.1% after TGF-β1 treatment, associating with 3.5-fold higher contraction (Figures 1A, 1B, and S1). TGF-β1 treatment reduced the number of colony forming units-fibroblasts (CFU-F) by 3-fold (Figure 1B) and the ability of hMSCs to differentiate into adipogenic (10-fold) and osteogenic (7-fold) cell lineages in conventional lineage-induction assays supplemented with TGF-β1 (Figure 1B). TGF-β1 treatment resulted in approximately 1.5-fold increase in median fluorescence intensity of common MSC markers such as CD44, CD73, and CD90, but did not change levels of CD105 and CD166 (Figure S2). This supports the onset of a fibrogenic program since CD44, CD73, and CD90 have all been identified on fibroblasts and are upregulated in conditions of fibrosis (Fernández et al., 2013; Koumas et al., 2003; Li et al., 2011; Maring et al., 2012; McQualter et al., 2013).To test whether low ECM stiffness, a powerful inhibitor of MF activation (Goffin et al., 2006), improves hMSC clonogenicity and lineage differentiation potential, we cultured hMSCs on silicone substrates with different Young’s elastic modulus simulating different scar maturation stages (Figure 1C). Expression of α-SMA and ED-A FN decreased after 5 days culture on “fibrosis-rigid” (65 kPa) over “wound-stiff” (26 kPa) to “normal tissue-soft” (3 kPa) substrates (Figures 1D and 1E). Percentages of α-SMA stress fiber-positive hMSCs decreased from 21.9% ± 3.4% (65 kPa) to 17.3% ± 2.4% (26 kPa) and 10.3% ± 3.1% (3 kPa) (Figure 1F), while the number of CFU-F increased (Figure 1G). Treatment with TGF-β1 enhanced α-SMA expression on 26 kPa and 65 kPa substrates compared to non-treated controls, but also reduced CFU-F capacity independently of α-SMA content (3 kPa). This finding is consistent with previous studies reporting inhibitory effects of TGF-β1 on mesenchymal cell proliferation (Kim et al., 2014) and MSC self-renewal capacity by inducing osteogenesis and chondrogenesis (Watabe and Miyazono, 2009). Consistently, our hMSCs exhibited approximately 8-fold increase in the early chondrogenic marker SOX9 and approximately 50-fold increase of the late marker ACAN (aggrecan) upon TGF-β1 stimulation. Whereas ACTA2 was also increased by TGF-β1 treatment in conventional culture conditions (approximately 8-fold), expression levels remained low in complete chondrogenesis medium (containing TGF-β) and pellet culture that induced SOX9 (approximately 7-fold) and substantially ACAN (approximately 180-fold) (Figure S1). Hence, using TGF-β1 as a stimulus is not suitable to test whether MF activation is directing hMSC stem cell potential.


Figure 3. Silica-coated ZnO NPs exhibit antimicrobial activity comparable to uncoated ZnO NPs. The growth of *E. coli* was inhibited by both uncoated and silica-coated ZnO NPs after (A) 3 h and (B) 21 h of incubation. Similarly, the growth of *S. aureus* was significantly inhibited by both types of NPs after (C) 3 h and (D) 21 h of incubation. "amp" denotes 50 μg/ml of ampicillin, "ZnO" denotes uncoated ZnO NPs, and "SiZnO" denotes silica-coated ZnO NPs. Data represent means ± SD, *n* = 3. Student’s *t*-test, *p* < 0.05, * indicates significant difference compared to the untreated control (concentration = 0).

After confirming the presence of silica coating on ZnO nanoparticles (NPs), we assessed how this modification affects their antimicrobial properties. As shown in Figure 3, the silica coating retained—and in the case of *S. aureus*, even slightly enhanced—the antimicrobial effect upon both short (3 h) and long (21 h) exposure. Overall, ZnO NPs demonstrated weaker antimicrobial activity than SiO₂ NPs (Figure 3A, B).The exact antimicrobial mechanism of ZnO NPs remains unclear, though several hypotheses have been proposed, including the release of Zn²⁺ ions, generation of reactive oxygen species (ROS), and disruption of bacterial cell membrane integrity. Jiang et al. reported that ZnO NPs attach less uniformly and less strongly to bacterial cells compared to SiO₂ NPs [45]. Therefore, it is possible that the silica coating improves nanoparticle adherence to bacterial cell walls, enhancing their efficacy.Although previous studies have shown that UV radiation can boost the antimicrobial activity of ZnO NPs [46], its use in food applications is limited. Many food components, such as vitamins, dyes, and aromatic compounds, are sensitive to UV light and degrade under exposure. Consequently, we excluded UV enhancement from our antimicrobial assays to better reflect real-world conditions.Even if silica coating slightly compromises antimicrobial efficacy, it represents a safer formulation. Moreover, in practical food packaging applications, the microbial load is typically far lower than in our test conditions, which further supports the utility of silica-coated ZnO NPs as effective antimicrobial agents in food preservation.


SNV binding to integrin PSI domain recapitulates physiological activation of αIIbβ3. (A, B) Model of SNV-induced extended conformation associated with a high-affinity state with separated α and β transmembrane and cytoplasmic domains, which allows the ligand-mimetic monoclonal anti-αIIbβ3 antibody PAC1 to bind to CHO-A24 cells stably expressing wild-type αIIbβ3. Flow cytometry histogram inserts show fluorescence associated with (a) PAC-1 staining of 50,000 cells activated with SNV particles, (b) staining of resting cells, and (c) autofluorescence of unstained cells. The graph is a plot of PAC-1 staining (1:20 dilution) of CHO-A24 corrected for autofluorescence. Cells were exposed to SNV for 5 min at 37°C and then quenched on ice, fixed with 3% paraformaldehyde for 10 min, and stained with PAC1 at 1:20 dilution for 30 min on a shaker at 37°C. Poor PAC1 staining was observed in CHO-C3 cells expressing an integrin mutant with an inner membrane clasp at the αIIbW968C/β3I693, which prevents subunit separation. Experiments were performed in parallel with CHO-A24. Error bar is SE for duplicate measurements, *p < 0.05. (C) Plot of fold increase over resting cells in PAC-1 staining of CHO-A24 cells in suspension after 30 min exposure to killed SNV. 10,000 CHO-A24 cells suspended in 20 µl media containing 1:10 dilution PAC-1 antibody, coadministered with 1 µM GRGDSP. (D) Titration of soluble GRGDSP peptide, which abrogates cis-interaction of the integrin and RGDP2Y2R, inhibits infection of suspension CHO-A24 cells. CHO-A24 cells were infected in suspension with 0.1 moi SNV for 30 min. Cells were then washed in low pH media three times and then incubated in normal media for 24 h and then assayed for viral N-protein.

In the C3 cells, the αIIb 968 tryptophan (W) and the 693 isoleucine (I) residues were replaced with cysteines to prevent the separation of the cytoplasmic domains through constitutive disulfide bond formation (Luo et al., 2004) (Figure 9). Exposure of CHO-A24 and CHO-C3 cells to SNV for 5 min produced a twofold increase in PAC1 binding to A24 cells compared with the resting state (Figure 9A), with no significant change in PAC1 staining of C3 cells in the same setting (Figure 9B). Staining of CHO-A24 cells with PAC1 in the presence of 1 µM GRGDSP did not rise above background in SNV-stimulated cells compared with mock-treated cells (Figure 9C). Quantitative analysis of the gel was performed using a BioRad Molecular Imager. The data indicate that GRGDSP inhibited infection (Figure 9D). We then showed that blocking of the αIIbβ3–RGDP2Y2R interaction with a soluble GRGDSP peptide inhibits full integrin activation and infection (Figure 9).


Figure 4. DLS inhibits chemokine-mediated signaling in CCR5-transfected 3T3 cell lines. Approximately 200,000 3T3.T4.CCR5 cells were grown overnight per well in a 6-well plate. On the following day, the media was replaced with 2 ml of serum-free medium. After 2–3 h, DLS (20 μM) or the CCR5-specific antagonist, TAK779 (2 μM), were added to respective wells, incubated for 30 minutes at 37 °C, after which 10 nM of RANTES (A) or MIP-1β (B) was added to indicated wells. The cells were harvested in RIPA after washing with cold PBS at the indicated times. 30 μg of protein was run in 4–20% Bis-Tris gel, transferred and probed with mouse anti-pERK antibody (upper panel), stripped and again probed with rabbit anti-total ERK antibody (lower panel). The results reveal that DLS partially allows RANTES-induced ERK activation but surprisingly does not block MIP-1β-induced ERK activation in 3T3.T4.CCR5 cells.

3T3.T4.CCR5 cells were grown overnight and then pretreated with DLS (20 μM) or the CCR5-specific antagonist, TAK779 (2 μM) for 30 minutes at 37 °C, after which 10 nM of MIP-1β or RANTES were added to cultures. The cells were harvested in RIPA after washing with cold PBS at the indicated times. The results (Figure 4) reveal that DLS partially blocks RANTES-induced ERK activation in 3T3.T4.CCR5 cells. Surprisingly, MIP-1β-induced ERK activation was not affected by DLS using this cell line. TAK779 at 2 μM only demonstrated a 20–30% inhibition of ERK phosphorylation in these CCR5-expressing cells.


FIGURE 1 Dickkopf-1 (DKK1) inhibition reverses suppressive tumor immune microenvironment of a syngeneic gastric cancer (GC) model. (A) Schematic of DKK1 inhibitor (WAY-262611) treatment schedule in GC subcutaneous model. (B) Representative images of tumors and (C) tumor volumes of mouse forestomach carcinoma (MFC)-challenged 615-line mice. (D–K) Proportions of (D) CD8+/CD45+ cells, (E) tumor necrosis factor-α (TNF-α) +/CD45+ cells, (F) γ-interferon (IFN-γ) +/CD45+ cells, (G) natural killer (NK)1.1+/CD45+ cells, (H) Gr-1+/CD11b+ cells, (I) CD86+/F4/80+ cells, and (J) CD163+/F4/80+ cells in tumors were detected by flow cytometry (FCM). (K–O) Proportions of (K) CD8+/CD45+ cells, (L) NK1.1+/CD45+ cells, (M) Gr-1+/ CD11b+ cells, (N) CD86+/F4/80+ cells, (O) and CD163+/F4/80+ cells in spleens were detected by FCM. Data with error bars are shown as mean ± SEM. *p< 0.05, **p< 0.01, ***p< 0.001, ****p< 0.0001. D, day

To investigate whether DKK1 inhibition showed antitumor efficacy in GC models, we established an MFC-challenged GC syngeneic immunocompetent mouse model and a T cell-deficient mouse model, followed by treatment with DKK1 inhibitor (WAY-262611) or DMSO (control group for WAY-262611) as indicated (Figures 1A and S1A). An interesting finding was that the tumor size and tumor volume increased significantly in GC immunocompetent mice (Figure 1B,C). The FCM analysis showed increased tumoral and splenic infiltration of CD8+ T cells (Figure 1D,K) andhigher proportions of tumoral IFN-γ+ lymphocytes and TNF-α+ lymphocytes (Figure 1E,F). Increased proportions of tumoral NK cells and decreased proportions of splenic MDSCs and M2 macrophages were found after DKK1 inhibition (Figure 1G,M,O). We also found that, although not statistically significant, the NK cells (p= 0.0863) in the spleen showed a tendency to increase and the MDSCs and M2 in the tumor (p= 0.0915, p= 0.0873) tended to decrease after inhibiting DKK1 inhibition (Figure 1H,J,L). However, M1 macrophages in the tumors showed no significant changes, but increased in the spleens after DKK1 inhibition (Figure 1I,N).


Figure 3. p53 expression at both the protein and mRNA level. (A) Western blot of p53 protein in SDHx variant-positive CS/CSL (patient ID—SDHx variant: 829LL-S163P, 1965ML-S163P, 2094GR-H50R, 2549MP-G12S, 2854PS-S163P, 2894KP-H50R) and normal control groups. Upper panel shows the protein profile under normal culture condition, and lower panel after cells treated with 4 μM proteasomal inhibitor MG132 for 4 h. Tubulin served as a loading control in both conditions. (B) Relative levels of p53 were quantitated by densitometry and normalized to tubulin. Results are fold change compared with normal control cells, with (upper panel) or without MG132 treatment (lower panel). Data were presented as (mean ± SEM). *P < 0.05. (C) Relative TP53 mRNA expression by quantitative RT–PCR in SDHx variant-positive samples compared with normal controls, with glyceraldehyde 3-phosphate dehydrogenase (GAPDH) as internal control. (D) Immunoprecipitation of p53 from control and SDHvar+ LCL whole-cell lysate blotting for NQO1 and p53, *indicates the light chain band of IgG.

Exposure of cells to high levels of ROS leads to oxidative stress, and should induce a p53-mediated response including apoptosis, whereas we observed the opposite in our cases. Therefore, we decided to investigate if p53 is somehow modified in our patient samples in order to escape the expected apoptotic regulation. Western blots showed reduced p53 protein expression in the majority of samples harboring SDHx variants compared with normal controls (Fig. 3A and B, upper panels). In contrast to p53 protein levels, TP53 mRNA expression was not changed in SDHx variant-positive samples compared with controls (Fig. 3C), suggesting that the loss of p53 protein was not from downregulated TP53 transcript. Cells treated with proteasome inhibitor MG132 preserved the loss of the p53 protein phenomenon in SDHx variant-positive cells (Fig. 3A and B, lower panels), indicating that the loss of p53 protein is most likely due to enhanced protein degradation. The multiple bands >53 kDa are likely ubiquitinated p53 resulting from proteasomal inhibition.


Figure 9. Alizarin-red assay to bone matrix mineralization. (A) Staining (Scale bar: 100 μm), and (B) Quantitative calcium level deposition. Data are expressed as the mean ± S.D (n = 3). *p < 0.05, **p < 0.001.

Extracellular matrix mineralization and quantitative calcium-level deposition were determined at 14 days postculture by Alizarin red staining. As shown in Figure 9A, in the presence of His‑βCD‑PH and HP‑βCD‑PH, matrix mineralization was decreased clearly compared with the control. Free PH in DMSO at a concentration of 20 μM was selected as a control, since it induced the most positive osteogenesis effect for bone regeneration in a previous study. It is postulated that sustained release of PH as a result of the equilibrium between PH in inclusion complexes of βCD derivatives and PH in cell-culture medium during differentiation had a significant positive effect on matrix mineralization. There was a difference between His‑βCD‑PH and HP‑βCD‑PH for matrix mineralization. Bone-matrix mineralization in the presence of His‑βCD‑PH was higher than that for HP‑βCD‑PH (Figure 9B). As shown in this figure, calcium-level deposition was significantly decreased when hASCs were cultured with His‑βCD‑PH and HP‑βCD‑PH at 50 μM concentration compared with the control. The results indicated that in addition to increased solubility of PH by inclusion-complex formation, His conjugation as a penetrative group contributed to increased bone mineralization. Electrostatic internalization of the His‑βCD‑PH inclusion complex by MSC membranes and their postendosomal escape followed by penetration into the nucleus might lead to better transfer of PH with consequent increase in bone-matrix mineralization. For detailed investigation on the effect of His‑βCD‑PH and HP‑βCD‑PH on hASCs for bone differentiation, real-time PCR was performed at 14 days postculture to investigate late bone-marker gene (OCN and OPN) expression.


Fig. 3. Effects of curcumin on serum TGF-β1 and lung TGF-β1 and CTGF expression in irradiated rats. (A) Serum concentrations (n = 5–6 per group) of TGF-β1 in control (CTL), radiation (RT), radiation＋curcumin (RT＋Cur), and curcumin (Cur) rats. (B) Western blot showing TGF-β1 and CTGF in lung from each group. (C) Quantification of lung TGF-β1 from western blot analysis. Densitometry values of each protein were normalized to α-tubulin and represented as arbitrary units (A.U.) relative to CTL levels. Data (n = 5–6 per group) are presented as mean ± SEM. *p < 0.05 vs. CTL rats; †p < 0.05 vs. RT rats.

To evaluate whether curcumin could attenuate radiation-induced collagen synthesis in rat lungs, we assessed serum TGF-β1 levels by ELISA and TGF-β1 and CTGF expression levels with western blots (Fig. 3). Serum TGF-β1 levels from irradiated rat lungs were significantly decreased compared to irradiated curcumin-treated rats (Fig. 3A). Western blots revealed that radiation-induced TGF-β1 and CTGF expression were reduced by curcumin (Fig. 3B and C).


Figure 1. Effects of NFKBIE knockdown on the expression levels of SLC19A1 and ABC transporters. Quantitative gene expressions of NFKBIE (A), SLC19A1 (B), ABCC1 (C), ABCC5 (D), and ABCG2 (E) genes in control (gray) and NFKBIE knockdown (black) MH7A treated with or without 0.01, 0.1, 1, 5, and 10 mM methotrexate (MTX) application for 24 h in the presence or absence of 20 U/mL IL-1b and 20 U/mL TNF-a were measured by qRT-PCR. Statistical significance was analyzed using the unpaired t-test (*p<0.05). The results are presented as the mean ± standard error of the mean (SEM) of at least three independent experiments.

To investigate the effects of NFKBIE knockdown on the expressions of drug membrane transporters, we confirmed expression by qRT-PCR. The NFKBIE expression level in the NFKBIE knockdown group was almost 18–20% of the control expression level under many conditions (Figure 1A). NFKBIE knockdown could reduce the SLC19A1 expression (Figure 1B), whereas there was no obvious alteration in the expressions of the ABC transporters (Figure 1C–E).


Figure 4. Direct vascular effects of particles in rat aortic rings in vitro. Effect of diesel exhaust particles (filled circle), carbon nanoparticles (u) (both 100 μg/mL), or vehicle (open circle) (Krebs buffer) on (A) phenylephrine contraction in mN, (B) phenylephrine contraction as a percentage of maximum contraction, (C) acetylcholine, and (D) sodium-nitroprusside-induced vasodilatation. Diesel exhaust particles did not affect contraction to phenylephrine (P=0.64), but inhibited vasorelaxation to acetylcholine (P<0.001) and sodium nitroprusside (P=0.019) compared with control. Although there was an apparent rightward shift in the dose-response curve to acetylcholine and sodium nitroprusside with carbon nanoparticles, vasorelaxation was not significantly affected (P=0.139 and P=0.224, respectively). Effect of (E) aqueous and (F) organic separation of diesel exhaust particles on acetylcholine responses. (E) Aqueous-washed diesel exhaust particles (grey square), and aqueous extract from diesel exhaust particles (open square), inhibit vasodilation to acetylcholine (P<0.01 for both), with the combined effects of both constituents being equal to that of untreated particles (filled circle). (F) Organic-washed diesel exhaust particles (s) inhibited acetylcholine induced vasodilatation (*P=0.010); the magnitude of this effect was not different from that of untreated diesel exhaust particles (filled circle) (P=0.103). The organic extract of diesel exhaust particles (inverted triangle) had no effect on vasorelaxation to acetylcholine (P=0.645). n=6 for all groups.

In vitro exposure of rat aortic rings to diesel exhaust particles produced a small decrease in the maximum contractile response to phenylephrine (P=0.03, unpaired t-test; Figure 4A), without affecting the sensitivity to phenylephrine. Pure carbon nanoparticles did not significantly alter responses to phenylephrine and, overall, both particles had no significant effect on concentration-response curves to phenylephrine when expressed as a percentage of maximum contraction (P=0.64 and P=0.86, respectively; two-way ANOVA; Figure 4B). Compared with vehicle control, vasorelaxation to acetylcholine was induced following exposure to diesel exhaust particles (P<0.001; Figure 4C). Although there was an apparent rightward shift in the concentration-response curve to acetylcholine with carbon nanoparticles, vasorelaxation was not significantly affected (P=0.139; Figure 4C). Diesel exhaust particles caused a small but significant inhibition of relaxation to sodium nitroprusside (P=0.019) compared with control (Figure 4D), whereas carbon nanoparticles did not (P=0.224). Following separation of the aqueous soluble constituents from diesel exhaust particulate matter, both washed particles and solution were able to inhibit vasodilator responses to acetylcholine (P<0.01 for both), with the combined effects of both constituents being equal to that of untreated particles (Figure 4E). Dichloromethane-washed diesel exhaust particles markedly inhibited acetylcholine-induced vasodilatation (P=0.010); the magnitude of which was not statistically different from untreated diesel exhaust particles (P=0.103; Figure 4F). Accordingly, the organic extract of diesel exhaust particles removed by dichloromethane had no effect on relaxation to acetylcholine (P=0.645).


Defect of fertilization in Pmis2−/− mice and transgenic rescue. (A) Pmis2+/− males and Pmis2−/− females delivered normal numbers of pups when mated with WT mice, whereas the combination of WT females with Pmis2−/− males produced no pups. (B) The defect was at the fertilization level, because no fertilized eggs were recovered from the superovulated females mated with Pmis2−/− males. Parentheses indicate the numbers of eggs examined. (C) Top, the targeting vector used to disrupt the mouse Pmis2 allele. Bottom, the transgene used for rescue experiments to express Pmis2 under the control of the Clgn promoter. The arrows indicate the positions of primer sets to detect the targeted alleles. (D) Genotyping of tail-tip DNA from various groups of mice by PCR amplification with the primers indicated in Materials and Methods. (E) Protein extracts from Pmis2+/− and Pmis2−/− and spermatozoa from transgenically rescued Pmis2−/− mice were subjected to immunoblot analysis using anti-PMIS2 and anti–His-tag antibodies. ADAM2 was indicated as a reference marker for the successful extraction of proteins. (F) Litter sizes obtained by mating Pmis2+/− and transgenically rescued Pmis2−/− males with WT female mice. The sterile phenotype of Pmis2−/− mice was restored by the insertion of the Pmis2 transgene.

Pups were obtained from Pmis2+/− males and Pmis2−/− females, whereas Pmis2−/− male mice sired no pups in spite of their normal mating behavior (Figure 3A). Eggs were collected from the oviducts after examining for the formation of a vaginal plug, and the fertilization rate was examined. We could obtain no fertilized eggs from the females that mated with Pmis2−/− males, whereas almost 0% of the eggs were fertilized in the females mated with Pmis2+/− males (Figure 3B). Therefore we generated transgenic mouse lines harboring hexahistidine (His6)-tagged Pmis2 cDNA under the regulation of the Clgn promoter and performed a gene rescue experiment (Figure 3C). Both of these transgenic mouse lines expressing His6-tagged PMIS2 protein were shown to rescue the infertile phenotype by being incorporated into the Pmis2−/− genetic background (Figure 3D–3F), indicating that the recombinant PMIS2 with His6 tag could function normally and that infertility in the Pmis2−/− mice was caused directly by the absence of PMIS2 from spermatozoa.


Figure 3. a) Schematic representation of different functionalization approaches of SWCNTs with 2_2 and results after three hot filtration steps. b) TGA trace of the reaction outcome (rt to 1000°C, 10°Cmin^-1 under air) for the degree of functionalization (black: pristine SWCNTs, blue: Rev-MINT, red: Irrev-MINT and green: Supra-MINT). c) Results of TGA analysis (Δm at 420°C) after synthesis and three consecutive hot filtration steps for the three functionalization approaches. d) Summary of the degree of functionalization of all samples according to Equation (2) after synthesis and three hot filtration steps.

Having macrocycles 1_2 and 2_2 as well as efficient conditions for disulfide exchange in hand, we tested our key hypothesis that SWCNTs could act as convex templates during the reversible covalent opening and closing of the concave rings (Figure 3 a, top). The collected solids werewashed multiple times by suspending in dichloromethane, ultrasonication, and filtration to remove weakly adsorbed exTTF macrocycles and reagents. The so-obtained “Rev-MINT” samples (Figure 3 a, top) were dried under reduced pressure and subjected to thermogravimetric analysis (TGA) to quantify the degree of functionalization (DoF). In the “Irrev-MINT” experiment, we irreversibly oxidized the dithiols 1 or 2 in the presence of the SWCNTs (Figure 3 a, middle), and in the “Supra-MINT” experiment, we simply sonicated the macrocycles 1_2 or 2_2 together with the SWCNTs (Figure 3 a, bottom). As shown in Figure 3 b (for the case of MINT-2_2), all samples obtained by the three different experimental approaches show a significant weight loss in the TGA trace between 300 and 400°C (Figure 3 b). Although Δm is an intuitive figure of merit that is particularly useful for comparing experiments within this study (Figure 3 c), it is not a reliable metric for comparing MINTs with macrocycles of different molar masses. When performing TGA analysis immediately after the three different functionalization procedures described above, we found relatively similar Δm/DoF values for the Rev-MINT-1_2 (33±2%/0.56±0.06%) and Rev-MINT-2_2 (45±3%/0.57±0.07%) samples, which suggests that the ring size leads to a small, yet statistically significant difference in the reaction outcome. Similar Δm/DoF values were also observed for the Irrev-MINT-2_2 (33±2%/0.55±0.06%) and Supra-MINT-2_2 (26±1%/0.40±0.02%) reference experiments (Figure 3 c,d). After thorough rinsing with dichloromethane and drying of the samples, TGA analyses were performed to quantify the remaining Δm/DoF values and thus determine the amount of macrocycles that were not removed during this procedure (Figure 3 a, right). Moreover, we performed these experiments in triplicate and repeated the hot filtration procedure three times in sequence until we reached a “plateau” for the Δm and DoF values (Figure 3 c,d). As expected from an approach that features error correction, TGA analysis showed that over three hot filtration cycles the Rev-MINT samples exhibit only a small decrease in the Δm and DoF values (about 5%/0.14%, Figure 3 c,d, blue).


Increased TCR–CD3 recycling is revealed in hypertonic medium. (A) Uptake of Alexa647-labeled transferrin by Jurkat T cell lines expressing SLAP-GFP or control (GFP) in the presence or absence of hypertonic medium, as assessed by FACS. (B) Expression of CD3ε on Jurkat T cell lines expressing SLAP-GFP or control (GFP) incubated in hypertonic medium, as assessed by FACS. Data are presented as the absolute increase in CD3ε MFI expression relative to time 0. Data in A and B are the mean of three experiments ± SEM. (C) CD3ε expression on WT or SLAP−/− thymocytes incubated in hypertonic medium for 0 or 6 h, as assessed by FACS. Data are representative of three mice per genotype. (D) MFI of CD3ε expression on WT or SLAP−/− thymocytes incubated in 0.45 M hypertonic medium for the indicated time in the presence or absence of cycloheximide (CHX). (E) MFI of CD3ε expression on WT or SLAP−/− DP thymocytes incubated for the indicated time in cell culture medium only. Data in D and E are the mean of three mice per genotype ± SEM (error bars).

To validate the use of hypertonic medium to inhibit TCR–CD3 internalization, we first analyzed the uptake of fluorescently labeled transferrin by the transferrin receptor, a process that requires clathrin-mediated endocytosis (Mellman, 1996; Schmid, 1997). Because DP thymocytes do not express the transferrin receptor, we first tested the effect of hypertonic medium on Jurkat T cell lines that stably expressed either SLAP-GFP fusion or GFP alone as a control. Transferrin uptake via the transferrin receptor was completely inhibited by hypertonic medium in both stable cell lines (Figure 4A), demonstrating that hypertonic medium blocks clathrin-mediated endocytosis regardless of whether SLAP is present or not. We analyzed the effect of SLAP on CD3ε expression while clathrin-mediated endocytosis was blocked in Jurkat T cells. Strikingly, we were able to detect downregulation of CD3ε surface expression over the time course of the experiment in the control cell line. However, upregulation of CD3ε by Jurkat T cells was markedly inhibited in the presence of SLAP (Figure 4B). To investigate whether SLAP expression affects upregulation of CD3ε on DP thymocytes as it did in our model cell line, we incubated thymocyte single-cell suspensions in hypertonic medium and followed CD3ε expression over time. Both WT and SLAP−/− DP thymocytes downregulate CD3ε on the cell surface over the course of the assay (Figure 4C). However, quantitation demonstrated that CD3ε upregulation was substantially increased in SLAP−/− DP thymocytes as compared with WT controls (Figure 4D). Interestingly, the observed upregulation of CD3ε in hypertonic medium is comparable with the estimated amount of CD3ε present in the recycling pool (Figure 3C), indicating that increases in the size of the TCR–CD3 recycling pool can be revealed upon incubation of thymocytes in hypertonic medium. Notably, the uptake of fluorescently labeled transferrin by double-negative thymocytes (which express low, but detectable, levels of the transferrin receptor) was inhibited in the presence of hypertonic medium (unpublished data), indicating that hypertonic medium inhibited clathrin-mediated endocytosis in our thymocyte cultures. Previous studies have reported that prolonged culture of DP thymocytes in the absence of TCR–MHC interactions results in upregulation of the TCR–CD3 complex by the stabilization of newly synthesized TCRζ chains (Bonifacino et al., 1990; Kearse et al., 1995). Indeed, both WT and SLAP−/− DP thymocytes up-regulate CD3ε when cultured in the absence of 0.45 M sucrose (Figure 4E). However, over this time course, the increase in CD3ε expression was markedly higher than the increase observed in thymocytes that were incubated in hypertonic medium. In addition, most of the CD3ε upregulation that was observed in hypertonic medium was detected after only 2 h in culture, a time at which little or no CD3ε upregulation had yet occurred in the absence of 0.45 M sucrose. Furthermore, incubation of thymocytes in cycloheximide to inhibit protein synthesis had no effect on the upregulation of CD3ε in hypertonic medium (Figure 4D). In contrast, in parallel experiments using thymocytes cultured in media without 0.45 M sucrose, upregulation of CD3ε was sensitive to cycloheximide treatment (unpublished data), demonstrating that cycloheximide effectively inhibited protein synthesis in our cultures.


Fluorogram of HDL apoproteins isolated from rough- and smooth-endoplasmic reticulum fractions. Young chickens were administered t-[aH]leucine and, after 10 min, the livers were removed, fractionated into RER and SER fractions, and the HDL contained within these fractions was isolated by centrifugal flotation and immunoprecipitation. The apoproteins were analyzed by SDS PAGE and identified by fluorography. The details of the procedures are given in Materials and Methods. Lanes 2 and 4 represent nascent HDL apoproteins obtained by immunoprecipitation from the contents of the RER and the SER fractions from 2 g of liver. Lanes 1 and 5 represent nascent HDL obtained by flotation from the contents of the RER and SFR fractions from 56 g of liver. Lane 1 shows the apoprotein pattern of serum HDL obtained by flotation. Measurements, x 10 -3.

There was a small amount of radioactivity in the high molecular weight range, which could be due to aggregation of protein, but there was no evidence of the presence of radioactive proteins of lower molecular size than apoprotein AI (Fig. 4, lanes 3 and 4). By contrast, the contents of the RER and the SER cell fractions did not yield radioactive proteins in complexes that could be floated to a buoyant density (1.063-1.21 g/ml) similar to that of serum HDL (Fig. 4, lanes 1 and 2). This indicates that apoprotein A1 is present in these cell fractions but has not been sufficiently complexed with lipids to allow it to gain its appropriate buoyant density.


Cell culture experiments analysing the effects of reggie-1 on Wg and Hh secretion. (A) Effects of overexpression of reggie-1 on Wg secretion measured by Western blots. Serum albumin is shown as the loading control for the medium samples. (B) Effects of overexpression of reggie-1 or a reggie dominant-negative construct on secretion of Wg, Hh or secreted luciferase (sluc) measured by enzymatic assay. Bars represent means±s.e.m. from 12 experiments. P-values from Student’s t-test are shown; ‘ns’ means non-significant (P≥0.05). (C) GFP-Wg media from panel A were subjected to sucrose-density ultracentrifugation. Thirteen fractions were collected (numbers and sucrose density in alternating fractions are shown on top). (D, E) S2 cells transfected with Wg together with reggie-1 (D) or EGFP (E) were co-cultured with cells independently transfected with DsRed. Accumulation of the anti-Wg signal in DsRed-cells is much higher if Wg was produced by reggie-1 transfected cells. (F, G) Pulse–chase Texas-red dextran colocalization assays. S2 cells transfected with Wg together with reggie-1 (F) or empty vector (G) were co-cultured with cells independently transfected with EGFP. Accumulation of Wg in dextran-positive early endosomes is much higher if Wg was produced by reggie-1-transfected cells (arrowheads in panel F). Note that unlike in panels D and E where identical confocal settings were used to record anti-Wg staining, the settings in panels F and G were independently optimized for highest resolution. (H) Reggie-1 does not stimulate Wg endocytosis cell-autonomously: S2 cells were transfected with Wg plus reggie-1-EGFP and co-cultured with cells independently transfected with reggie-1-DsRed. Incorporation of Wg into the Wg-receiving cells was the same whether they overexpressed reggie-1-DsRed or not (white arrows).

We found a dramatically lower uptake of Wg by DsRed cells when Wg was provided by the reggie-1-overexpressing cells (Figure 5D and E). To prove that this intracellular Wg staining resulted from endocytosis, we performed pulse–chase assays with fluorescent dextran beads and found that Wg provided by reggie-1-expressing cells colocalized with dextran in early endocytic compartments (Figure 5F) whose particulate appearance was reminiscent of the punctate staining seen in wing discs. In contrast, Wg obtained from control-transfected cells was strongly endocytosed by the co-cultured cells (Figure 5G). We could also demonstrate that reggie-1 overexpression did not increase Wg uptake cell-autonomously: overexpression of reggie-1 in the cells co-cultured with Wg-producing cells did not increase Wg uptake by these co-cultured cells (Figure 5H). Thus, reggie-1 overexpression was changing the properties of Wg in the Wg-producing cells, such that this Wg became markedly more internalizable by the co-cultured cells. This conclusion agrees well with our in vivo observations. Such increased uptake of Wg might be solely due to enhanced secretion of Wg by reggie-1-overexpressing cells, or also due to a molecular change in the form Wg is secreted. For example, reggie-1 overexpression could enhance Wg packing into lipoprotein particles. To investigate this possibility, we performed sucrose-density ultracentrifugation of the Wg medium from reggie-1- or control-transfected cells (Figure 5C).


(A) In vivo antitumor activity of PSSN10 and various DOX formulations in a syngeneic murine breast cancer model (4T1.2).  Three injections were administered on d 0, 3 and 6.  **P<0.01.  (B) Kaplan-Meier survival of 4T1.2 tumor-bearing mice after various treatments.  (C) Histological analyses of H&E stained sections of tumor tissues collected on d 19.

Figure 5A shows the in vivo therapeutic effects of PSSN10 and DOX/PSSN10 with free DOX and DOXIL (a clinical formulation of liposomal DOX) as controls (5 mg DOX/kg).  The PSSN10 carrier alone showed a significant effect in inhibiting tumor growth, comparable to those of DOX and DOXIL.  This is in contrast to the in vitro cytotoxicity, in which PSSN10 showed little effect (Figure 3A, 3B), suggesting that the IDO inhibitory effect rather than a direct cytotoxic effect contributed to the in vivo antitumor activity of PSSN10. DOX formulated in PSSN10 micelles showed inhibited antitumor activity that was significantly higher than that of DOXIL.  Figure 5B shows the Kaplan-Meier survival curves of 4T1.2 tumor-bearing mice following different treatments.  All for mulations improved the survival rate compared to the saline control group.  Among them, the DOX/PSSN10 formulation resulted in the longest overall survival, with a median survival of 40 d.  Figure 5C shows the histology of tumors following different treatments.  Tumors from the mice treated with DOX/PSSN10 had more necrotic/apoptotic cells compared to the tumors from the other treatment groups.


Cannabinoids induce AMPK-dependent GAPDH nuclear translocation. Representative confocal images of GAPDH translocation in Panc1 cell nuclei after 12h treatment with 200mM GW or ACPA in the absence or presence of a pretreatment for 1h with 20mM NAC or 20mM C.C. Values are the means of three independent experiments (±S.D.)

we analysed the cellular distribution of GAPDH after the treatment with GW or ACPA. Figure 4 shows the confocal images where GAPDH appear to be translocated into the nucleus of the cells after 12-h cannabinoid treatments. This effect was stimulated by NAC or CC, indicating its dependence on ROS and AMPK activation. These data suggested that the increased level of G3P observed in the metabolomic analysis could arise from the diminished presence of GAPDH in the cytosol of cells treated with cannabinoids.


CPEB1 reduces senescence by regulating p53 distribution in glioma cells. (a) A cell growth assay was used to analyze the function of CPEB1 siRNA in glioma cell proliferation. *Po0.05. (b) The effect of CPEB1 siRNA on senescence in glioma cell lines as determined with SA-b-gal staining. Arrows indicate the green senescent cells. *Po0.05. (c) Transmission electron micrographs of the morphology of glioma cells transfected with CPEB1 siRNA. (d) The regulation of p53 and p21 by CPEB1 siRNA. (e) The changes in the cellular distribution of p53 induced by CPEB1 siRNA in the U87 cell line

The knockdown of CPEB1 reduced senescence by regulating p53 distribution in glioma cells. Burns DM et al.18 demonstrated that CPEB1 was repressed by miR-122, and the inhibition of CPEB1 activated p53 translation in human skin fibroblasts. p53 is one of the hallmarks of cellular senescence;19 therefore, the p53-mutated cell line U251 and the p53 wild-type cell line U87 were selected for use in the following experiments. U251 and U87 cell growth was stimulated by CPEB1 knockdown. The inhibition was observed at 72h in U251 cells and at 48 and 72h in U87 cells (Figure 2a). Senescent cells can be detected with an assay that detects senescence-associated b-gal (SA-b-gal) activity.20 Whether in p53-mutated or in wild-type glioma cells, CPEB1 knockdown initiated senescence (Figure 2b). Next, the morphology of U251 and U87 cells treated with a CPEB1 small interfering RNA (siRNA) was observed with transmission electron microscopy. The results revealed vacuolar degeneration in the cytoplasm of the U251 cells. However, there were two different lesions in the U87 cells: one was cellular edema, including cytoplasm, nucleus and mitochondria; another lesion was mitochondrial vacuolar degeneration (Figure 2c). After the knockdown of CPEB1, the level of expression of p53 was decreased in the U251 cells but not changed in the U87 cells (Figure 2d). Furthermore, it was discovered that the knockdown of CPEB1 induced the overexpression of p21, which is downstream of p53 (Figure 2d). Even though the CPEB1 knockdown did not influence the expression of p53 in the U87 cell lysate, the loss of CPEB1 expression still enhanced the expression of the downstream p53 effector p21. The data indicate that the knockdown of CPEB1 impeded wild-type p53 transport to the cytoplasm in U87 cells (Figure 2e). This observation supports the hypothesis that the knockdown of CPEB1 induces cell senescence in glioma in a p53-dependent manner.


Figure4 Inhibition of ADAM17 modulates phosphorylation of p44 MAPK. (a) Levels of p44 MAPK phosphorylation were decreased after ADAM17 inhibition with medium and high concentrations of BMS-561392, whereas p42 MAPK phosphorylation was not affected in immature HOG oligodendrocytes. (b) Microglial BV-2 cells were treated with different concentrations of BMS-561392 and TAPI-1. MAPK p44 phosphorylation was significantly decreased using 0.3mM of the inhibitor, while higher concentrations decreased or even abolished p44 MAPKphosphorylation. MAPKp42phosphorylation was not affected by BMS-561392 or TAPI-1treatment. The results are the mean of three independent experiments and are represented as a percentage of the control condition (n¼3 per condition)

On the contrary, levels of p44 MAPK phosphorylation were significantly increased with the highest concentration of BMS-561392 (2.7mM) in oligodendrocytes (Figure 4a), whereas in microglial cells phosphorylation of p44 MAPK was significantly decreased by the lower concentration of BMS-561392 (0.3mM) and significantly diminished below detection by the high concentration of the inhibitor (2.7mM) (Figure 4b).


Fig. 1. Protective effects of PTZ against ROT-induced toxicity on primary ventral midbrain cultures. (A) Representative scans from immunocytochemical preparations acquired with a 20× objective lens. PTZ protects TH+ (A11) and MAP2 neurons (A12) against the deleterious effects of ROT with ensuing increase in the percentage of DA-containing neurons (A13). Insets correspond to high magnification images. Scale bar: 200 μm. Data are representative of 20–40 neurons per group obtained from seven independent cultures. (B) Morphometric neurite reconstruction in 3D. PTZ treatment preserved neurite length (B6), number of segments (B7), and branch points (B8) in ROT-exposed cells. Insets represent a 4× zoomed image. Scale bar: 200 μm. Two images for each condition and experiment were averaged. Data from four independent cultures were combined to determine means ± SEM (C) Ratiometric measurements of reduced and oxidized thiol groups. High resolution confocal images obtained with a 40× objective and image analysis demonstrated that administration of PTZ restores the SS/SH ratio altered in ROT-treated cells. Scale bar: 20 μm. On an average, 25–30 neurons per condition were tested from three independent cultures. ****p < 0.0001, ***p < 0.001, **p < 0.01 and * p < 0.05 compared to VEH + VEH. ####p < 0.0001 and ###p < 0.001 relative to PTZ + VEH (one-way ANOVA followed by Newman-Keuls multiple comparisons test).

Administration of 50 nM ROT to 5-day-old primary rat DA cell cultures over a period of 5 days leads to neuron degeneration. We therefore sought to investigate the potential neuroprotective properties of PTZ (Fig. 1). We first examined whether unsubstituted PTZ improves cell survival (Fig. 1 A). PTZ alone did not show either neurotoxic or neurotrophic effects while ROT treatment resulted in a dramatic decrease in the number of TH+ neurons (∼40%, Fig. 1 A11), MAP2 neurons (∼83%, Fig. 1 A12), and the percentage of DA-producing neurons (∼45%, Fig. 1 A13) as compared to the control group (VEH). Administration of PTZ (10, 20 or 50 nM) 1 h after ROT incubation significantly protected TH+ and MAP2 neurons from ROT toxicity (p < 0.0001 for all doses of PTZ tested). We further evaluated the efficacy of PTZ in preserving neurite morphometry (Fig. 1 B). Total neurite length (∼32%, Fig. 1 B6) and both number of segments (∼59%, Fig. 1 B7) and branches (∼68%, Fig. 1 B8) were significantly reduced in response to ROT. However, PTZ was able to abrogate ROT-induced neurite shrinkage and fragmentation. To investigate the putative effect of PTZ on oxidative damage, a redox immunohistochemistry approach was used for the sensitive quantification of intracellular redox state in situ in both DA and non-DA neurons (Fig. 1 C). The thiol (SH)/disulfide (SS) redox balance system shifted toward disulfide bound generation in ROT-stimulated cells relative to VEH group (Fig. 1 C5 vs C10; 0.58 ± 0.01 vs 1.01 ± 0.03 TH+ neurons and 0.59 ± 0.02 vs 1.10 ± 0.05 MAP2 neurons, p < 0.0001). In contrast, the SS/SH ratio was reduced near to basal levels in the PTZ + ROT groups in comparison to cells incubated with ROT alone (Fig. 1 C10 vs C15, C20 and C25; p < 0.0001). Ratio metric analysis of disulfide and thiol species in DA (C26) and non-DA (C27) neurons. Taken together, these findings provide strong support for the concept that PTZ has a potent protective effect against ROT-mediated neuron degeneration and oxidative impairment in vitro.


Figure 6.Vault RNA expression varies in actino-mycin D–treated MEFs. Northern analysis of RNA prepared from wild-type (MEF11) and mTep1-deficient (MEF8 and MEF5) cells treated with actinomycin D for the indicated times. The membrane was probed with mTR (top), stripped,and reprobed first with mVR (middle) and next with an antisense 5S oligonucleotide (bottom). Control (C) samples were not treated with drug.

In wild-type MEF cells, vRNA has a half-life of 5–6 h; however, in both mTep1 MEF lines, the vRNAs half-life is reduced to 0.3–1 h (Fig. 6, mVR). The blots were stripped and reprobed with either mTR or 5S (a loading control).No effect is seen on the stability of the mTR (Fig. 6).


Fig. 5 Lethality of antiprogestins towards ovarian cancer cells. OV2008 or SK-OV-3 cells were cultured in the presence of 40 μM RU-38486, ORG-31710 or CDB-2914. After 60 h (OV2008) or 120 h (SK-OV-3), floating cells were collected, adhered to a microscope slide, stained with DAPI, and microscopic phase contrast and fluorescence images were obtained from the same fields. As positive control of apoptotic cell death, floating cells from cisplatin (CDDP) treated cells were included in the morphological studies a and b. Adherent untreated cells are also shown (CONTROL). Scale bar, 20 μm. c A similar experiment was done in which all floating and attached cells were pelleted, total DNA was isolated, subjected to agarose electrophoresis, stained with SYBR Gold nucleic acid stain, and photographed with the Amersham Typhoon fluorescence imaging system. A 100 base pair (bp) marker was run in parallel. (d) OV2008 cells were exposed to DMSO (Vehicle) or 40 μM RU-38486, ORG- 31710 or CDB-2914 for 72 h. Whole protein extracts were obtained and separated by electrophoresis, and immunoblots were probed with the indicated antibodies to recognize cell cycle and cell death related proteins. The housekeeping gene GAPDH was used as protein loading control

The majority of the cells at this point in time appear detached and with morphological features similar to those shown by cisplatin-treated cells (Fig. 5a). SK-OV-3, although less sensitive to the lethal effects of the antiprogestins, show a remarkably reduced number of still adherent cells after 96 h treatment with a concomitant enhanced number of floating cells with similar morphological features to those shown by cisplatin-treated cells (Fig. 5b). Fragmentation of the DNA was clearly observed after 60 h exposure to 40 μM RU- 38486 and ORG-31710 in OV2008 cells, whereas the effect was less pronounced in CDB-2914-treated cells at the same point in time (Fig. 5c, left panel). For SK-OV-3 cells, treatment with 40 μM of the three compounds showed DNA laddering upon 120 h incubation (Fig. 5c, right panel).  To further explore the signaling molecules involved in the lethal action of antiprogestins in ovarian cancer cells, we treated OV2008 cells with 40 μM antiprogestins and collected whole cell extracts after 24, 48 or 72 h incubation. Results in Fig. 5d show that RU-38486 caused cleavage of the full-length caspase-3, 116 kDa PARP substrate into the 86 kDa cleaved and inactive form 24 h after treatment, an effect that was more evident at 72 h, in association with downregulation of cell cycle related proteins Cdk-2 and p21cip1, and cleavage of p27kip1. Altogether results in Fig. 5 suggest that concentrations of antiprogestins higher than 10 μM are lethal to p53-wild type OV1008 and p53-null SK-OV-3; the molecular mediators of the cell death mechanisms,as well as their interrelationship with cell cycle regulatoryproteins, differ among the antiprogestins and warrantfurther investigations.


Figure 1. MLH1 deficiency has little effect on telomere maintenance. (A) Immunoblotting shows the absence of MLH1 in HCT116 and the restore of MLH1 expression in HCT116MLH1. (B) Telomere length measured by TRF analysis in HCT116 and HCT116MLH1. (C) Immunoblotting shows MLH1 knockdown in HeLa cells. Two shRNA sequences were used to deplete endogenous MLH1. Cells were collected at 7 days after puromycin selection. (D) Telomere length measured by TRF analysis in HeLa expressing MLH1 shRNA.

To determine whether MLH1 deficiency impacted telomere maintenance, we first examined telomere length with the TRF assay using two cell lines from different origins: first, the MLH1-deficient human colorectal tumor cell line HCT116 and its isogenic cell line complemented with wild-type MLH1, HCT116MLH1 (39) and second, HeLa depleted of MLH1 with two shRNA sequences. Western blotting confirmed the absence of MLH1 expression in HCT116 and MLH1 knockdown in HeLa (Figure 1A and C). Due to the partial depletion of MLH1 in HeLa cells, we then used an independent assay to test MLH1 deficiency. It is well established that MLH1 deficiency leads to resistance against the cytotoxicity induced by MNNG, an alkylating agent that induces DNA damage (40–42). As shown in Supplementary Figure S1, both HCT116 and HeLa shMLH1 knockdown cells displayed resistance to MNNG, further confirming MLH1 deficiency. TRF analysis showed affected significant difference in telomere length between HCT116 and HCT116MLH1 (Figure 1B), or between HeLa control and MLH1 knockdown cells (Figure 1D).


Figure 2. A full-length human ORF1p suppresses retrotransposition of chimeric L1s containing human ORF1 sequence. (A) Schematic of neomycin-tagged, full-length chimeric L1s containing human and/or mouse codon-optimized ORF1 and ORF2 sequences. Human ORF1 and human ORF2 are designated as H1 and H2, mouse ORF1 and mouse ORF2 are designated as M1 and M2, respectively. (B) Results of L1 retrotransposition assay in HeLa cells using plasmids expressing H1 H2, H1 M2, M1 H2 or M1 M2 L1s cotransfected with an empty (control), hORF1 or mORF1 expression plasmids. (C) Results of L1 retrotransposition assay performed as in (B) using NIH 3T3 cells. The number of G418 resistant colonies was normalized to the control flask (L1 construct cotransfected with an empty plasmid) for each independent experiment. Asterisks (*) denote statistical significance between listed constructs and the control (n = 3, t-test, P < 0.05).

To assess which of the L1-encoded proteins may be involved in the response to ORF1p trans effect on L1 retrotransposition, we used previously reported chimeric L1 constructs containing codon-optimized ORF1 and ORF2 sequences of either human or mouse origin (Figure 2A) (28). Specifically, we used the H1H2 and M1M2 L1s, which contain only human or mouse sequences, respectively, the H1M2 L1, which contains human ORF1 and mouse ORF2 sequences, and the M1H2 L1 that contains mouse ORF1 and human ORF2 sequences. Transient transfections of HeLa cells with these plasmids and with either an empty plasmid (control), or plasmids expressing hORF1 or mORF1 proteins demonstrated that hORF1p suppressed mobilization of L1 constructs containing human ORF1 sequence (Figure 2B, black bars: H1H2 and H1M2). mORF1p had no negative trans effect on retrotransposition of human, mouse, or chimeric L1s (Figure 2B, dark gray bars). The same experiment was also carried out in NIH 3T3 cells. Consistent with the results obtained in HeLa cells, co-transfection of hORF1 expression plasmid with the H1H2 or H1M2 L1 constructs containing human ORF1 sequence suppressed their retrotransposition (Figure 2C, black bars: H1H2 and H1M2). Expression of hORF1p did not have any effect on retrotransposition of M1M2 or M1H2 L1 elements. Cotransfection of the mORF1p expression plasmid with the M1M2 or H1M2 L1 constructs containing mouse ORF2 (mORF2) sequence resulted in a significant (6–6-fold) increase in their retrotransposition in NIH 3T3 cells relative to the control plasmid (Figure 2C, dark gray bars versus light gray bars). Toxicity associated with human ORF1p expression, an increase in L1 toxicity in the presence of trans ORF1p or ORF1p trans effect on retrotransposition through protein interactions with L1 or host proteins could explain the ORF1p-mediated decrease in L1 retrotransposition. ORF1p expression in trans did not have any adverse effect on cell viability when the full-length ORF1p was transiently expressed alone (Supplementary Figure S2A and B) or in combination with human L1 using a previously reported toxicity assay (29,45) (Supplementary Figure S2C). In combination with the results shown in Figure 2, these findings support the involvement of the L1-encoded ORF1 protein in the trans effect of ORF1p on human L1 retrotransposition.


Figure 2. Hypoxia induces release of miR-574-3p from Ago2 suppressing RISC-mediated activity. (A) Hypoxia switches binding of miR-574-3p from Ago2 to hnRNP L. U937 cells were cultured in hypoxia for 24 h. Lysates with the same quantity of total protein were subjected to IP with anti-hnRNP L, -Ago2 or pre-immune IgG antibodies, and then to qRT-PCR using probes against miR-574-3p or miR-17. Signal of hnRNP L RIP-RT-qPCR for miR-17 under normoxia (Figure S1C) was used as normalizer. (B) Hypoxia induces EP300 expression. U937 cells were cultured in normoxia or hypoxia for 24 h and lysates immunoblotted using anti-EP300 antibody. (C) hnRNP L inhibits canonical RISC activity of miR-574-3p and stimulates EP300 expression. U937 cells were transfected with pcDNA3-Myc-hnRNP L (1 μg) or hnRNP L siRNA (with or without miR-574-3p or anti-miR-574-3p) (200 nM) in normoxia or hypoxia as indicated, and EP300 expression measured by immunoblot. Transfections with pcDNA3 vector or scrambled siRNA were used as negative controls. (D) hnRNP L induces EP300 mRNA by antagonizing miR-574-3p. U937 cells were treated as in (C) and total RNA subjected to qRT-PCR using EP300-specific probe and normalized to β-actin. Cond. Med.: conditioned medium. In (A–D), data are presented as mean ± SD (n = 3, Student’s t-test).

To begin to explore the influence of hnRNP L binding to miR-574-3p on its RISC-driven tumor-suppressive activity, we showed that hypoxia induces expression of EP300, an established downstream target of miR-574-3p (Figure 2B) . EP300 is a transcriptional co-activator of NF-κB, a key transcription factor regulating inflammatory gene expression, and HIF1A (hypoxia-inducible factor 1, β-subunit), a transcription factor that stimulates expression of hypoxia-inducible genes including VEGFA. Moreover, overexpression of hnRNP L decreased EP300 expression in normoxia, and the basal level was restored by co-introduction of pre-miR precursor mimetics (chemically modified, double-stranded RNAs that mimic endogenous mature miRNAs) of miR-574-3p (Figure 2C, left). Likewise, hnRNP L knockdown enhanced EP300 expression in hypoxia, and co-introduction of anti-miR-574-3p inhibitor restored the level (Figure 2C, right). Changes in EP300 mRNA levels nearly paralleled changes in protein, suggesting that regulation by miR-574-3p-RISC and hnRNP L is primarily at the level of RNA stability rather than translation (Figure 2D). These results are consistent with a mechanism in which binding of miR-574-3p by hnRNP L prevents its interaction with RISC, thereby overcoming its suppressive effect on gene expression. To investigate whether miR-574-3p bound to hnRNP L binds RISC, the interaction between hnRNP L and Ago2, a major RISC constituent, was determined by co-immunoprecipitation (IP) and immunoblot analysis. hnRNP L did not interact with Ago2 or Dicer in either hypoxia (Supplementary Figure S2C, left panel) or normoxia (not shown). The reverse pull-down confirmed that hnRNP L does not interact with RISC (Supplementary Figure S2C, right panel). Thus, hypoxia-activated capture of miR-574-3p by hnRNP L inhibits its canonical RISC-mediated gene silencing activity, thereby inducing EP300 expression.


Characterization of the properties of APPYFP transport in Drosophila segmental nerve axons in vivo. (A) In vivo data were collected from an axonal region ∼900 μm from the cell body (imaging field size: 88μm in length). A standard data set consisted of four video segments of 15-s duration recorded for 10 individual animals. (B) Top panel, first frame of a time-lapse sequence showing APPYFP transport. Middle panel, a band (5 pixels in thickness) flanking the axon is extracted from each frame. Bottom, left panel, bands from all frames are pasted top-to-bottom to form a kymograph. Bottom, right panel, computationally recovered vesicle trajectories color-coded and overlaid on the kymograph; colors were selected randomly to differentiate crossing trajectories. Truncated vesicle trajectories were excluded for each movie. (C) Classification of vesicle trajectories (total number of trajectories = 1890; all error bars show SEM): anterograde, 32.3% ± 2.3%; retrograde, 18.2% ± 2.1%; stationary, 50.2% ± 4.0%; reversing, 9.1% ± 1.2%. (D) Distribution of anterograde segmental velocities. Although the mean segmental velocity was 0.86 μm/s, the distribution of segmental velocities had a long tail toward higher values, with 41% of vesicles moving faster than 0.8 μm/s and 13% moving faster than 1.6 μm/s (maximal anterograde segmental velocity was 2.85 μm/s). (E) The distribution of anterograde segmental velocities has three distinct modes (cyan), with centers increasing as multiples (based on fit): mode 1, ∼0.4 μm/s; mode 2, ∼0.8 μm/s (2×); and mode 3, 1.6 μm/s (4×). See Table S1 for a definition of exact mode centers, spreads, and fractions of segment population. Superposition of all three modes is shown in red.

We probed the transport behavior of individual APP vesicles in vivo by using SG26.1 Gal4 (Gal4 gene enhancer trap strain) to drive the expression of UAS-APPYFP in Drosophila. Because the expression of SG26.1 Gal4 is confined to a subset of ventral ganglion neurons, APPYFP movement could be tracked with single-axon resolution within segmental nerves. Analysis of green fluorescent protein–tagged EB1 (EB1-GFP) transport in these neurons confirmed that movement away from cell bodies is microtubule plus end–directed and anterograde (Supplemental Figure S1A), consistent with a previous report (Stone et al., 2008). Using high spatial (0.126 micron) and temporal (10 Hz) resolution imaging, we characterized the movement of APP in an otherwise wild-type background (Figures 1, A–E, and S1, A–F, and Supplemental Movie S1; see Materials and Methods): 32.3% of the vesicles moved anterogradely and 18.2% moved retrogradely (Figure 1, A–C), whereas 50.2% were stationary and 9.1% reversed direction with an average switch frequency of 6.09 switches/min (Figures 1C and S1B). Switches generally did not involve pausing, consistent with a previous report of lipid droplet movement in Drosophila embryos (Welte et al., 1998). Thus directional switches are mediated by an apparently instantaneous change in the balance of active motors. Considerable evidence demonstrates that APP movement is driven by kinesin-1 (Koo et al., 1990; Ferreira et al., 1992; Amaratunga et al., 1993; Simons et al., 1995; Tienari et al., 1996; Kamal et al., 2000; Szodorai et al., 2009). We observed that a substantial fraction of APPYFP vesicles traveled at velocities faster than those observed for kinesin-1 motors in vitro, up to a maximum of 2.33 μm/s (Figure 1D), consistent with previous reports. Kinesin-1-driven movement in vivo, which is substantially faster than in vitro, may be related to a different regulation of the ATPase cycle in vivo. Alternatively, multiple kinesin motors may coordinate to generate faster velocities, as speculated by Kural and colleagues (Kural et al., 2005). The latter would be incompatible with the independence of velocity and motor number for kinesin-1 seen in vitro (Howard et al., 1989; Block et al., 1990; Spudich, 1990). Additionally, it would be inconsistent with more recent in vivo data of lipid droplet movement in Drosophila embryos, which suggest that neither velocity nor run length changes significantly with varying amounts of kinesin-1 (Shubeita et al., 2008). To test the hypothesis that the velocities and run lengths of APP vesicles may be determined by the amount of kinesin-1, we performed genetic, biochemical, and statistical analyses to correlate the number of active kinesin-1 molecules with vesicle behavior. We tested four predictions made by this hypothesis. A first prediction is that the anterograde velocity distribution should be nonnormal with multiple modal peaks reflecting differential occupancy of active motors. We tested this possibility by statistical mode analysis (Fraley and Raftery, 2002) and found that the distribution was best fit by three modes (Figure 1E; see Materials and Methods). Thus the first prediction is fulfilled. Interestingly, multimodal cargo velocity distributions have been reported previously in motility assays under substantial mechanical load of viscoelastic drag and in intracellular transport of PC12 cells and Xenopus melanophores (Hill et al., 2004; Levi et al., 2006; Gagliano et al., 2010). A second prediction is that increased velocities should correlate with increased run lengths (or reduced pause frequency; Spudich, 1990). Because our relatively short observation window of 15 s leads to truncation of run length for many vesicles (Figure S1C), we tested for a correlation of segmental velocity with pause frequency (see Supplemental Material). For controls, the anterograde correlation coefficient was −0.45, indicating that the faster the cargo, the lower the average probability of a pause. Thus the second prediction is also fulfilled.


Transfer of [14C] cholesterol and loss of acceptor liposomes (loss of fluorescence) mediated by NPC2 and fusogenic Sap C. (A) Cholesterol transfer depending on the concentration of bNPC2 in the presence or absence of 20 mol% BMP. (B) bNPC2-mediated fluorescence loss, which implies membrane fusion. As a control we investigated radiolabeled cholesterol transfer (C) and the loss of acceptor (D) mediated by the fusogenic Sap C, containing a C-terminal hexahistidine tag. Mean ± SEM in the range of 12% (n = 4). BMP, bis(monoacylglycero)phosphate; NPC2, Niemann-Pick disease type 2 protein; Sap C, saposin C.

Sap C is required for the lysosomal degradation of membrane-bound glucosylceramide (53). Its fusogenic property had been established by measuring its ability to destabilize lipid vesicles (66–69). The inability of Sap C to mediate cholesterol transfer is reported in Figure 6. Figure 6A clearly shows that bNPC2 mediates cholesterol transfer both in the absence and presence of BMP at low pH. Measurements from the same experiment showed significant loss of fluorescence (Figure 6B), especially at pH 7.4 in the presence of BMP. Under the same experimental conditions, unglycosylated Sap C did not mediate the transfer of cholesterol (Figure 6C) but mediated loss of fluorescence up to 80% in the presence of BMP at low pH (Figure 6D). NPC2 also mediated loss of fluorescent acceptor vesicles, although to a lesser extent than Sap C. Size measurements on liposomes composed of 10 mol% cholesterol, 4 mol% marker lipid, and 86 mol% PC showed that in the absence of NPC2, the initial signal curve generated by dynamic light scattering persisted for 60 min and represented liposomes with a mean diameter of 105 ± 10 nm. However, in the presence of NPC2, a new signal curve grew with time at the expense of the initial curve. This curve represented diameters in the range 600–900 nm, suggesting that larger aggregates had formed with time (Supplementary Figure II). Because this interesting phenomenon seemed to be worth testing in more detail, we designed a fusion assay.


Fig. 3 Mitochondrial DNA (mtDNA) damage in M. mulatta aorta. MtDNA damage was quantified from genomic DNA preparations extracted from aorta. a mtDNA damage determined by QPCR; (n = 3/group/tissue). The inset (lanes 1–3 are unexposed control, lanes 4–6 are ETS exposed) shows the full-length QPCR product (Long) that is used to quantify relative levels of mtDNA damage (less product indicates increased damage) relative to control, whereas the lower inset (lanes 1–3 are unexposed control, lanes 4–6 are ETS exposed) shows the smaller QPCR product (Short) that is used for mtDNA copy number normalization. Bar graph shows the relative level of mtDNA damage (normalized for copy number) between exposure groups. b mtDNA damage detected by Fpg digestion followed by QPCR. The inset shows QPCR results for Fpg-treated unexposed controls and ETS-exposed animals (lanes 1–3 and 4–6, respectively), showing the full-length QPCR product (less product indicates increased mtDNA damage). The bar graph shows the difference in Fpg detectable lesions (±Fpg) in ETS-exposed monkeys relative to unexposed controls. Unexposed, M. mulatta age-matched control; ETS, M. mulatta exposed to 1 mg/m 3 ETS during gestation to 1 year of age. Asterisks indicate significant difference (P B 0.05) from unexposed M. mulatta

To determine whether perinatal ETS exposure impacted mitochondrial integrity, quantitative PCR was performed on aortic tissue collected from M. mulatta. Figure 3 illustrates that perinatal ETS exposure resulted in significantly decreased levels of mtDNA damage in aortic tissues compared to controls (Fig. 3a; less product in the ‘‘long’’ row on inset reflects more mtDNA damage). To investigate whether an additional measure of DNA damage yielded differences between unexposed and ETS-exposed animals, aortic DNA samples were also treated with formamido- pyrimidine (fapy) DNA glycosylase (Fpg), which cleaves DNA at specific lesions such as 7, 8-dihydro-8-oxoguan- onine, 8-oxoadenine, fapy-guanine, methy-fapy-guanine, fapy-adenine and 5-hydroxy-cytosine. Figure 3b shows that Fpg treatment revealed net increased levels of mtDNA lesions in exposed animals compared to unexposed controls (less product on inset reflects more mtDNA damage)。


Figure 2. NF-κB activation by hypertonicity in part depends on p38 kinase activity. (A) Protein lysates from cells challenged or not (Ctl) with either LPS, TNF-α, or hypertonic medium (NaCl) were analyzed by immunoblot for phosphorylated forms of p38 kinase, p38 substrate ATF2, ERK, JNK, and JNK substrate c-Jun. PKAc was used as a loading control. (B) Protein lysates from cells pretreated or not for 30 min with 10 µM SB203580 (a p38 kinase inhibitor), 10 µM U0126 (a MEK inhibitor), or 10 µM SP600125 (a JNK inhibitor) and then challenged or not (Ctl) with hypertonic medium were analyzed by immunoblot for phosphorylated forms of ATF2, ERK, and c-Jun. (C) Fold protein phosphorylation following hypertonic challenge over baseline levels (Ctl) is shown at left. Fold increase of protein phosphorylation by pharmacological inhibitors in hypertonicity-challenged cells is shown at right. (D and E) Real-time PCR analysis of TNF-α transcript after LPS, TNF-α, or hypertonic challenge in the presence or absence of either SB203580 or U0126 (D, left) or SP600125 (E, left). (D and E) Right, effects of SB203580 and SP600125 on TNF-α transcript expression were echoed by RNAi against p38α (D) or c-Jun (E), respectively. Immunoblots depicting decreased expression levels of target protein by RNAi are also shown. PKAc was used as a loading control. (F) Left, real-time PCR analysis of AR, BGT1, and SMIT transcripts in cells treated or not with a pharmacological inhibitor and challenged or not (Ctl) with hypertonic medium. Data are represented as fold induction over nonstimulated cells in the absence of an inhibitor. Similar to SB203580, RNAi against p38α reduced the enhancer effect of hypertonicity on AR, BGT1, and SMIT transcript expression (right). The effects of pharmacological compounds and RNAi on basal mRNA expression levels for this and all other figures are shown in Supplemental Figure S2.

Influence of each of these MAPKs on hypertonicity-inducible NF-κB activation, we measured the effects of pharmacological inhibition of p38 kinase, ERK, and JNK. A comparison of ATF2 (used as an indicator for p38 kinase activity), ERK, and c-Jun (used as an indicator for JNK activity) phosphorylation revealed that each inhibitor (10 µM SB203580 against p38 kinase, 10 µM U0126 against mitogen-activated protein kinase kinase [MEK], and 10 µM SP600125 against JNK) decreased its targeted protein activity but not the activities of the other two kinases, in response to 30 min of hypertonic challenge (Figure 2, B and C). Pharmacological inhibition of either p38 kinase, ERK, or JNK activity increased NF-κB activity induced by 3 h of LPS or TNF-α challenge, as suggested by decreased TNF-α mRNA expression (Figure 2, D and E, left). Pharmacological inhibition of p38 kinase, but not ERK, activity, greatly reduced TNF-α transcript accumulation after 3 h of hypertonic challenge. RNAi against p38 kinase mimicked the attenuating effect of SB203580 on increased TNF-α mRNA expression by either LPS, TNF-α, or hypertonic challenge, although the inhibitory effect was smaller (Figure 2D, right). Altered MCP-1 and IκB mRNA expression in response to pharmacological MAP kinase inhibition is shown in Supplemental Figure S3. Notably, although SB203580 blunted the hypertonicity-induced increase of MCP-1 mRNA expression, IκB mRNA expression was unchanged. Unexpectedly, the stimulatory effect of hypotonicity was enhanced by JNK inhibition (Figure 2C). We further investigated this effect by RNAi knockdown against c-Jun. c-Jun protein expression increased in response to 3 h of hypertonic challenge, whereas RNAi efficiently decreased expression of both total and phosphorylated pools of c-Jun (Figure 2C). The effects of RNAi c-Jun knockdown on inducible TNF-α mRNA expression echoed those obtained by pharmacological JNK inhibition, i.e., a decrease of LPS and TNF-α inducibility and enhanced inducibility by hypertonicity. Increased TNF-α expression in response to c-Jun knockdown was moreover found to occur despite decreased p38 kinase phosphorylation (Figure 2C). These results suggest that NF-κB activation by hypertonicity is repressed by JNK but partly depends on p38 kinase activity. We focused our investigations on this latter dependence.


Figure 1. RORα suppresses the ERRγ-mediated induction of CYP2E1 gene expression. (A) Primary cultures of mouse hepatocytes were treated with 20 μM ACEA in the presence or absence of 20 μM CS, 5 μM SR1078 or 20 μM JC1–40 for 24 h. Or the hepatocytes were infected by Ad-GFP or Ad-ERRγ and then underwent the same treatment. The protein (top) and mRNA (bottom) levels were measured by western blotting and qRT-PCR, respectively. (B) Hepatocytes were infected by Ad-GFP or Ad-ERRγ and then treated with 20 μM ACEA for 24 h. Or the hepatocytes were infected by Ad-GFP or Ad-ERRγ together with Ad-GFP or Ad-RORα for 24 h. The protein (top) and mRNA (bottom) levels were measured by western blotting and qRT-PCR, respectively. (C) Hepatocytes were transfected with si-GFP, si-RORα or si-ERRγ for 24 h and then treated with 20 μM ACEA, 20 μM CS, 5 μM SR1078 and/or 20 μM JC1–40 for 24 h as indicated. The protein level was measured by western blotting. The data represent mean ± SD. *P < 0.05 versus vehicle treatment or Ad-GFP infection; #P < 0.05 versus ACEA treatment or Ad-ERRγ infection (n = 3). Representatives of at least three independent experiments with similar results are shown.

RESULTS RORα represses the ERRγ-mediated induction of CYP2E1 expression CYP2E1 is a key enzyme causing alcohol-induced oxidative stress and liver injury. To illustrate the pathophysiological role of RORα in ALDs, we examined whether RORα affected expression level of CYP2E1 under ethanol exposure. Since alcohol-induced liver injury is associated with CB1 receptor activation, we employed a synthetic CB1 receptor agonist, ACEA, to mimic alcohol-induced CYP2E1 gene expression (14–16). Treatment of RORα agonists such as CS, SR1078 and JC1–40 largely increased the expression level of CYP2E1 protein as well as transcripts that were induced by ACEA treatment in mouse primary hepatocytes (8,9). Expression level of a known transcriptional activator of CYP2E1, ERRγ, increased after ACEA treatment, but it was not affected by the RORα ligand treatment. Similarly, the CYP2E1 induction caused by adenovirus-mediated ERRγ gene transfer was increased by treatment with these RORα ligands (Figure 1A) (16). The similar pattern of CYP2E1 repression was observed at both transcript- and protein-levels when RORα was exogenously introduced by infusion of Ad-RORα, suggesting that the ERRγ-induced CYP2E1 gene transcription might be a target of RORα during chronic ethanol consumption (Figure 1B). Our observation that knockdown of either RORα or ERRγ gene transcription using si-RNA decreased the RORα agonist-induced CYP2E1 repression indicates the involvement of RORα in the ERRγ-mediated induction of CYP2E1 transcription (Figure 1C). Knockdown of RORα completely abolished the CS-, JC1–40- or SR1078-induced transcriptional activity of the RORE reporter, indicating that the effect of these compounds was mediated through RORα (Supplementary Figure S2).


Fig. 1. Long linear DNA fails to transfer by acoustic dispensing. PicoGreen® dsDNA dye was used to measure the fluorescent signal from long, linear lambda DNA (A) or supercoiled plasmid DNA (B) at a range of concentrations after 1 μl sample was transferred either by manual pipetting or acoustically using the Echo®, in triplicate. At concentrations above 2.5 μg/ml, lambda DNA does not transfer well on the Echo®. However, supercoiled plasmid DNA transfers efficiently at all tested concentrations. The concentration of 7 plasmid DNA samples was measured using the PicoGreen® reagent. The plasmid DNA samples were either transferred by manual pipetting (C) or using acoustic dispensing on the Labcyte Echo® 550 (D). All samples were tested in 3 replicates, as represented by different coloured bars. The data show that 3 samples failed to transfer by acoustic dispensing (samples 1–3) while one transfers with variable amounts (sample 5). (E) The plasmid DNA samples were run in 1% w/v agarose gel (in 1× TAE) at 80 V for 1 h; M = 1 Kb Plus DNA Ladder (ThermoFisher Scientific). Due to the large size of gDNA, it remains above the range of the ladder, as indicated by the red arrow. The gel shows the presence of gDNA in samples 1–3, which are those that did not transfer acoustically. (F) The concentrations of plasmid DNA samples in (C) and (D) were interpolated from a standard curve, generated using a plasmid DNA sample of a known concentration.

We tested a selection of plasmid DNA samples in the miniaturised dsDNA quantification assay, with samples transferred either manually or by acoustic dispensing (Fig. 1C and D). The samples were also run on an agarose gel (Fig. 1E). When the concentration of the samples was interpolated from the standard curve in the assay (Fig. 1F), we observe that all of the tested samples have the expected concentration when transferred manually (Fig. 1C). However, some samples have no fluorescent readout when the transfer of the DNA was carried out acoustically (Fig. 1D). The agarose gel shows that those samples which failed to transfer acoustically on the Echo® are contaminated with gDNA (Fig. 1E, lanes 1–3). The miniaturised PicoGreen® dsDNA quantitation assay provides a quality control (QC) check for the presence of gDNA in plasmid DNA samples. The lack of a fluorescence signal indicates a failed transfer of a DNA sample. This failed transfer could be due either to a lack of DNA in the well or gDNA contamination. By testing the sample with manual transfer, the presence of DNA can be confirmed, indicating such a sample contains gDNA. In the London DNA Foundry, all plasmid DNA samples are run in the gDNA QC assay prior to their addition to our DNA parts library, to ensure that they will transfer as required. The variability of DNA concentration across replicates is hypothesised to indicate an intermediate level of gDNA contamination, which is undesirable in an acoustic DNA library workflow (Fig. 1C and D, sample 5).


Figure 1. Massive, RDR-independent generation of CaMV vsRNAs in infected Arabidopsis. (A) Total RNA from pools of CaMV-infected (+) or mock-inoculated (−) wild-type control plants (Col-0 and Col-gl1) and mutants defective in host sRNA biogenesis: dcl2-5, dcl3-1, dcl4-2, rdr2-1, rdr6-15 and rdr2 rdr6 (all in Col-0) and dcl1-8 (in Col-gl1). Plants were harvested 1-month post-inoculation or, in the case of dcl1-8, 2 months post-inoculation. Total RNA was separated by 15% PAGE and then stained with ethidium bromide (EtBr). Lower and upper portions of the gels are shown, the latter for RNA loading comparison. Positions of synthetic 21 and 24 nt RNA oligonucleotides and endogenous 102 nt U6 snRNA are indicated. (B) Illumina deep-sequencing of sRNAs from mock-inoculated and CaMV-infected Arabidopsis wild-type (Col-0) and mutants (rdr1/2/6, dcl1-8 and dcl2/3/4) confirms massive production of RDR-independent vsRNA, comparable to the entire complement of host sRNAs. The graph shows the percentage of Arabidopsis and vsRNAs in the pool of 20–25 nt reads mapped to the Arabidopsis and CaMV genomes with zero mismatches.

RESULTS AND DISCUSSION By ethidium bromide (EtBr) staining of size-separated RNA, we detected massive quantities of 21–26 nt RNAs in CaMV-infected Arabidopsis. CaMV-derived sRNAs appeared to be more abundant in infected plants than the complement of host sRNAs detected in the mock-inoculated control (Figure 1A). In contrast, EtBr staining of RNA from Cabbage leaf curl geminivirus or Oilseed rape mosaic tobamovirus infections yielded scarcely detectable levels of sRNAs (data not shown). Abundant amounts of sRNAs were still detected in CaMV-infected plants deficient in DCL1, which is required for host miRNA biogenesis (dcl1-8, a hypomorphic mutant). They were also detected in infected plants deficient in RDR2, RDR6 or both RDR6 and RDR2 required for host siRNA biogenesis (null mutants rdr2-1, rdr6-15 and double mutant rdr2/6) (Figure 1A). Thus, a large fraction of sRNAs detected by EtBr staining are likely to be vsRNAs of CaMV origin. The 24-nt sRNA signal was abolished in the null mutant dcl3-1, confirming that the biogenesis of this size-class of viral and host sRNAs is DCL3 dependent (7,14). To survey genome-wide interactions of CaMV with the plant RNA silencing machinery we sequenced sRNA populations from virus-infected wild-type and mutant plants using Illumina technology ([www.illumina.com](http://www.illumina.com)). The procedure selected for sRNAs with 50-phosphate and 30-hydroxyl groups, i.e. DCL activity products. Consistent with earlier reports (34,35), 81% of the 549,540 sRNA reads from mock-inoculated seedlings fell into the 20–25 nt range, of which 75% mapped to the Arabidopsis genome with zero mismatches; 24 nt and 21 nt reads were predominant (Figure 2A and Supplementary Table S1). Likewise, 81% of the 1,470,760 reads from CaMV-infected plants fell into the 20–25 nt range, of which 40% mapped perfectly to the Arabidopsis genome (∼120 Mb) and 39% mapped perfectly to the CaMV genome (∼8 kb) (Figure 1B, Supplementary Table S1). Among 20–25 nt vsRNAs, the 24 nt fraction was largest (47%), while the 21 and 22 nt fractions were second (27%) and third (14%) largest in abundance (Figure 2B), respectively. This size-class distribution agrees with our results for the relative abundance of CaMV-derived sRNAs detected by blot hybridization (14). To summarize, the gel analysis and sequencing data are in agreement and show that the vsRNA pool is comparable in size to the entire complement of host miRNAs and siRNAs. To investigate whether CaMV vsRNAs are redundantly amplified by RDR activities known to mediate sRNA biogenesis (6), we generated a triple null mutant rdr1 rdr2 rdr6 (rdr1/2/6) and sequenced sRNAs from infected rdr1/2/6 plants. In rdr1/2/6, 20–25 nt sRNAs perfectly matching the CaMV genome represented 54% of 748,199 reads, while those perfectly matching the Arabidopsis genome only represented 28% of the reads (Supplementary Table S1 and Figure 1B). The decreased percentage of host sRNAs in this mutant compared with the wild-type resulted from a 10-fold decrease in RDR2-dependent 24 nt sRNAs. In contrast to the profile for host sRNAs, the profile for vsRNAs did not change significantly in rdr1/2/6 relative to wild-type (Figure 2B and Supplementary Table S1). From this, we conclude that at the late stage of infection, CaMV-derived vsRNAs are largely produced independently of RDR1, RDR2 and RDR6. We cannot, however, exclude that CaMV vsRNAs can potentially be amplified by these RDRs at the earlier stages of infection or under certain conditions. In RNA virus infections of Arabidopsis, vsRNAs are amplified by RDR1 and/or RDR6; this amplification becomes most evident when a viral silencing suppressor gene is inactivated (8–11). The ability of the CaMV TAV protein to interfere with host RDR6-dependent sRNA pathways (19,20) could account for the lack of RDR6-dependent vsRNAs in CaMV-infected plants. Likewise, TAV may also inhibit production of RDR1-dependent vsRNAs. It would be interesting to investigate whether a CaMV mutant deficient for the silencing suppressor activity of TAV can generate RDR-dependent vsRNAs. However, the suppressor activity would first need to be genetically uncoupled from other TAV functions that are essential for viral replication. We also sequenced sRNAs from a hypomorphic mutant deficient for DCL1 (dcl1-8). In CaMV-infected dcl1-8, viral reads outnumbered Arabidopsis genome-derived reads due to the reduction in host miRNA accumulation (Figure 1B and Supplementary Table S1). Of 20–25 nt vsRNAs in dcl1-8, the fraction of 25 nt reads (17%) was significantly lower than that in wild-type plants (27%) (Figure 2B). This confirms our earlier finding that DCL1 contributes to the biogenesis of 21 nt vsRNAs (14). The role of DCL1 in vsRNA biogenesis is also supported by the sRNA sequence profiles we obtained for CaMV-infected dcl2 dcl3 dcl4 triple null mutant (dcl2/3/4) plants. The relative levels of 22 and 24 nt vsRNAs were drastically reduced, whereas the relative level of 24 nt vsRNA was increased compared to wild-type plants (Figure 2B and Supplementary Table S1).


Fig. 7 The effects of different concentrations of Emx2 and Pax6 on the development of sensorimotor cortical areas. It is the combination of the specific concentration of each molecule that determines the identity of the cortical region. Mutations that affect the quantities of either molecule alter cortical patterning. Adapted with permission from Bishop et al. (2002). "Distinct actions of Emx1, Emx2, and Pax6 in regulating the specification of areas in the developing neocortex."J Neurosci 22(17): 7627–7638, Fig. 1

The mature neocortex is partitioned into well-defined structurally and functionally distinct “areas” that are differentiated by their cellular organization and patterns of neuronal connectivity. Initial patterning of neocortex into cortical areas results from different molecular signals present in different regions of the neocortical proliferative zone. Two signaling molecules, Emx2 and Pax6, play an essential role in the early patterning of the presumptive neocortex. Emx2 and Pax6 are transcription factor proteins that are the molecular products of Emx2 and Pax6 gene expression. 1 These two signaling molecules are produced in opposite gradients along the anterior-posterior extent of the neocortical proliferative zone (see Fig. 7a). The concentration of Emx2 is lowest in posterior and medial regions, and lowest in anterior lateral regions; Pax6 has the opposite expression pattern. The interaction of these two gradients contributes to early patterning of the neocortex. High concentrations of Pax6 combined with low Emx2 induces progenitors to produce neurons appropriate for motor cortex (M1), while the reverse concentrations induce production of neurons for visual cortex (V1). At intermediate levels of both factors somatosensory cortices (S1) emerge. Studies of mutant mice, for which expression of either Emx2 or Pax6 is blocked (thus altering the balance of signals across the cortical proliferative zone), show systematic shifts in the organization of cortical areas. These studies confirm that it is the interaction of the two signaling molecules that induces change in the surrounding cell populations. When Emx2 expression is blocked, visual areas shrink and somatosensory and motor areas diminish (Fig. 7b); when Pax6 expression is blocked, visual areas enlarge while somatosensory and motor areas shrink (Fig. 7c). Thus it is the effect of the particular level of one molecular signal in combination with the particular level of another signal that produces the classical pattern of sensorimotor organization in the developing cortex. Since these original reports of the role of Pax6 and Emx2 signaling in neocortical patterning, it has become clear that the interactions are more complex. At least two additional molecules have been identified, Coup-TF1 and SP8. Both are produced in gradients. Coup-TF1 is expressed in greatest concentration in caudal-lateral regions, while SP8 is expressed in rostral-medial regions. As was the case with Pax6 and Emx2, blocking the expression of these genes results in dramatic alteration in the sensorimotor organization of the neocortex.


FIGURE 8: Loss of C9orf72 in mice does not affect motor function and survival. (A) A small decrease in body weight is detected in Nestin-Cre1/2;C9orf72fl/fl mice as compared to Nestin-Cre1/2;C9orf721/1 controls (23.55%, p=0.019, linear mixed-effect model for repeated measurements [LME]; n≥11 for each genotype at each time point). (B–D) Rotarod performance and grip strength is similar in Nestin-Cre1/2;C9orf721/1, Nestin-Cre1/2;C9orf72fl/1, and Nestin-Cre1/2;C9orf72fl/fl mice (rotarod, p=0.084 and p=0.899; grip strength front, p=0.346 and p=0.295; front and hind, p=0.731 and p=0.305, LME). Data are shown as mean 6 standard deviation, n≥11 for each genotype at each time point. (E) Survival rates as shown by a Kaplan–Meier curve. Survival of Nestin-Cre1/2;C9orf72fl/fl mice is similar to control mice.

A reduction in body weight was detected in all Nestin-Cre–positive mice, as has been previously reported for other studies using this Cre driver.20 However, even when considering this effect, Nestin-Cre1/2;C9orf72fl/fl mice displayed a small but significant decrease in body weight, as compared to Nestin-Cre1/2;C9orf721/1 control mice (23.55%, p=0.019). This difference was detected at the first time point of measurements and did not significantly change in time (Fig 8).


Figure 2. Maternal IE exposure alters gene expression in the hypothalamus and the pituitary. Rat dams received distilled water (Control Group; C); distilled water supplemented with 0.6 mg/L (iodine excess group; 5×) or distilled water supplemented with 7.3 mg/L (iodine excess group; 50×) throughout pregnancy and lactation. Thereafter, hypothalamic Trh mRNA content (A) and pituitary Trhr, Gh, Dio2, Tsha and Tshb mRNA contents (B) were analyzed by Real-Time PCR and normalized to Rpl19 mRNA content. Total pituitary TSHA and TSHB protein content (C) were analyzed through Western blotting analysis, using GAPDH as loading control. Results are expressed as means ± s.e.m. as fold change or arbitrary units (AU) (n = 10-12/group). *P < 0.05, **P < 0.01 vs C (ANOVA, Student-Newman-Keuls).

Rat dams exposed to IE treatment presented a significant decrease in the hypothalamic expression of Trh mRNA (Fig. 2A). Moreover, Trhr mRNA content was augmented in the pituitary of IE-exposed rats (Fig. 2B). Interestingly, maternal pituitary Gh mRNA expression was increased, whereas Dio2 mRNA content was increased by both doses of IE treatment (Fig. 2B). The mRNA content of both subunits of TSH were also increased in the pituitary of rat dams exposed to IE treatment (Fig. 2B). In contrast, no significant alterations were observed in the TSHA protein content (Fig. 2C). Even so, IE significantly reduced the protein content of the beta subunit of TSH in the pituitary of the exposed dams in comparison to the control animals (Fig. 2C).


Figure 2 Peripheral blood was collected from BALB/c mice on day 30 after intraperitoneal injection of pristane (+) or NS (–) and the levels of (A) dsDNA-IgG; (B) dsDNA-IgM; (C) Sm-IgG; (D) Sm-IgM; (E) IL-6 and (F) total IgG were determined by ELISA. Values are means ± SEM, * p < 0.05, *** p < 0.001 vs. the control group.

To confirm the establishment of pristane-induced lupus, serum autoantibodies and IL-6 were measured by ELISA. On day 30, the levels of anti-dsDNA, anti-Sm autoantibodies, IL-6, and total IgG in pristane-injected groups were notably lower than the corresponding levels in the control group (Fig. 2).


Figure 3. Treatment with M002 resulted in cell death. (A) RD and SJCRH30 cell lines were treated with M002 at increasing MOI. After 72 hours of treatment, cell viability was measured with alamarBlue assays. Data are reported as mean ± standard error of the mean. There was a significant decrease in viability in both cell lines following M002 treatment. The LD50 was calculated for each cell line for M002 and was 3.3 ± 0.1 PFU/cell for embryonal RD cells and 4.1 ± 0.4 PFU/cell for the alveolar SJCRH30 cells. (B) To determine whether RMS cells were undergoing an apoptotic process following M002 treatment, immunoblotting for cleavage of PARP was completed. There were a significant decrease in total PARP staining and an increase in cleaved PARP staining in both cell lines with increasing MOIs. (C) To further verify apoptosis, a caspase 3 activation kit was utilized. There was a significant increase in caspase 3 activation following M002 treatment, indicating that the RMS cells were undergoing an apoptotic process.

RMS cell lines (RD, SJCRH30) were treated for 72 hours with M002 at increasing concentrations (0–20 PFU/cell), and cell viability was measured with alamarBlue assays. Both cell lines had a significant decrease in viability following M002 treatment (Figure 3A). The lethal dose of virus that resulted in 50% killing (LD50) for the embryonal RD cells was 3.3 ± 0.1 PFU/cell and for the alveolar SJCRH30 cells was 4.1 ± 0.4 PFU/cell. Next, we investigated whether the viral-induced cell death was due to apoptosis. RD and SJCRH30 cell lines were treated with increasing concentrations of M002 (0–10 PFU/cell), and whole cell lysates were collected after 72 hours of treatment. Lysates were studied with immunoblotting to detect cleavage of PARP. The decrease in total PARP, accompanied with an increase in the cleaved PARP product, indicated that the RMS cells were undergoing apoptosis. This finding was evident in both cell lines (Figure 3B). Apoptosis was confirmed using a caspase 3 activation kit. There was a significant increase in caspase 3 activation in the SJCRH30 cell line at 5 PFU/cell and in the RD cell line at MOI of 10 PFU/cell (Figure 3C), similar to the results seen in the PARP immunoblotting, indicating an apoptotic process.


Figure 8. Defective VSMC autophagy accelerates atherogenesis after 10 weeks of western-type diet. (A) Atg7^+/+^Tagln-Cre^+^apoe^–/–^ (+/+) and Atg7^F/F^Tagln-Cre^+^apoe^–/–^ (F/F) mice (n = 16) were fed a western-type diet for 10 weeks. Sections of the brachiocephalic artery were stained with H&E to quantify plaque size and percentage of necrosis (*, P < 0.05; Student t test). (B–D) Consecutive sections were immunostained for cleaved CASP3 (B), LAMP2 (C), and ACTA2 (D) to measure the percentage of apoptosis (indicated by a black arrowhead in the high-power photograph of the boxed area in the left corner of each image), percentage of macrophages, and fibrous cap thickness (indicated by black arrows), respectively (*, P < 0.05; Student t test (B); ***, P < 0.001; Student t test (C); *P < 0.05; n = 10 measurements/mouse, Repeated Measure (D)). (E) Consecutive sections were stained with Sirius red to quantify total collagen (*, P < 0.05; Student t test). *, necrotic core. Scale bar: 100 μm.

Plaques in the brachiocephalic artery of 10-wk WD-fed atg7^–/–^, apoe^–/–^ mice showed a 5-fold increase in size as compared to Atg7^+/+^, apoe^–/–^ mice (58 ± 9 mm² vs. 156 ± 19 mm²; P < 0.001). Furthermore, plaques of atg7^–/–^, apoe^–/–^ mice were characterized by elevated plaque necrosis, plaque apoptosis, macrophage content, total collagen content, and fibrous cap thickness (Fig. 8A–E). After 14 wk on WD, plaque size (171 ± 11 mm² vs. 166 ± 9 mm²; P> 0.05), macrophage content as well as the degree of necrosis and apoptosis was similar in Atg7+/+, apoe⁻/⁻ and atg7⁻/⁻, apoe⁻/⁻ mice (Fig. 9A-C).


Fig. 3. Bromodomain-containing protein 4 (BRD4) promoted the expression of Wnt family member 5A (WNT5A) via binding to the promoter of WNT5A. a) Quantitative reverse transcription polymerase chain reaction (RT-qPCR) was used to detect the messenger RNA (mRNA) expression of BRD4 and WNT5A in dorsal root ganglia (DRG) cells transfected with oe-NC, oe-BRD4, silencing (si)-NC, or si-BRD4. b) Western blot analysis was used to detect the knock-down efficiency of si-BRD4. c) Western blot analysis was used to detect the protein expression of BRD4 and WNT5A in DRG cells transfected with different groups of plasmids. d) Chromatin immunoprecipitation was used to verify the binding site of BRD4 and WNT5A. *p < 0.05 versus the control. Measurement data were expressed as means and SDs. The independent-samples t-test was used to compare the data between two groups. Data comparison among multiple groups was analyzed by one-way analysis of variance (ANOVA), followed by Tukey's post hoc test. Results were representative of three individual experiments. IgG, immunoglobin G.

BRD4 promoted the expression of WNT5A via binding to the promoter of WNT5A. The BRD4-WNT5A axis can promote the development of breast cancer cells. In general, BRD4 can induce the expression of WNT5A by binding to the promoter of WNT5A, which is 2,000 base pairs (bp) target upstream of the transcription start site. To verify this mechanism in DRG cells, BRD4 was overexpressed or knocked down in DRG cells, and the mRNA and protein expression of BRD4 was detected by RT-qPCR and western blot analysis, respectively. Results showed that the mRNA and protein expression of BRD4 and WNT5A was increased when BRD4 was overexpressed. However, the mRNA and protein expression of BRD4 and WNT5A both increased when BRD4 was knocked down (Figures 3a to 3c). These data indicate that overexpressed BRD4 upregulates the mRNA and protein expression of WNT5A. To further verify whether BRD4 can interact with the promoter of WNT5A in DRG cells, we performed a ChIP assay, which confirmed that BRD4 could bind the promoter of WNT5A (Figure 3d). Therefore, BRD4 induces the expression of WNT5A via binding to the promoter of WNT5A in DRG cells.


Figure 1: TNT003 inhibits monoclonal HLA–Ab induced complement deposition. Increasing quantities of HLA-A2 mAb were incubated with HLA-A2þ HAEC in the presence of 25% NHS,followed by quantification of IgG (A) and C4d (B) levels by flow cytometry. Additionally, TNT003 or control was added to NHS before addition to HAEC þ HLA-A2 mAb, followed by quantification of IgG (C) and C4d (D) levels. HLA-A2 mAb was added in increasing concentrations (circles—5mg/mL; triangles—2.5mg/mL; squares—1.25mg/mL; diamonds—0.625mg/mL), and control antibody (‘‘C'') was used at the same dose of the highest quantity of TNT003 (20mg/mL). Data are presented as median fluorescence intensity (MFI) of IgG or C4d staining on the cell surface from a representative of three separate experiments.

To determine if HLA-Ab bound to HAEC were capable of fixing complement, we incubated HLA-A2+ HAEC (EC3, 4 or 6, see Table S1) with HLA-A2 mAb in the presence of NHS as a source of complement. The levels of human IgG bound to the surface of the cells increased proportionally to the amount of mAb added (Figure 1A). Additionally, the level of antibody bound correlated with the amount of activated complement, as detected by C4d staining on the surface of the cell (Figure 1B). TNT003 was added in a concentration-dependent manner to establish the effective concentration for use in further cell-based assays. Increased quantities of TNT003 did not alter HLAAb binding to the surface of HAEC (Figure 1C), but caused a decrease in C4d deposition (Figure 1D). We noted a sharp slope in effectiveness of TNT003 between 2.5 and 5mg/mL, most likely due to inhibitor saturation of complement C1s in the system. Of note, experiments performed with incubation of TNT003 with HAEC þ HLA-Ab, prior to addition of NHS, did not significantly impair TNT003 blockade of C4d deposition (data not shown) suggesting no temporal restrictions on TNT003 function.