Honey bee venom contains various enzymes with wide medical and pharmaceutical applications.Apis mellifera lamarckii) 8.9-fold to a very high specific activity of 6033 U/mg protein using DEAE–cellulose and Sephacryl S-300 columns. The purified bee venom PLA2 is monomeric 16 kDa protein and has isoelectric point (pI) of 5.9. The optimal activity of bee venom PLA2 was attained at pH 8 and 45 °C. Cu2+, Ni2+, Fe2+, Ca2+, and Co2+ exhibited a complete activating effect on it, while Zn2+, Mn2+, NaN3, PMSF, N-Methylmaleimide, and EDTA have inhibitory effect.The phospholipase A2 (PLA2) has been apparently purified from the venom of Egyptian honey bee (The purified bee venom PLA2 exhibited anti-platelet aggregation and anti-coagulation activities which makes it promising agent for developing novel anti-clot formation drugs in future. Apithoperties . PLA2s oent type , 11, 12.ent type . PLA2s hent type , 15. PLAent type . PLA2 caent type , 18. PLAent type . For allApis mellifera lamarckii) were obtained from Assiut Governorate, Egypt. Bee venom was extracted from 500 forager workers that were caught at entry of the colony and immobilized via rapid freezing at – 20 °C. Individuals were dissected, sting devices and venom reservoirs were removed, disrupted in tube with 2.5 ml dH2O, and finally centrifuged at 12000×g for 5 min at 4 °C and supernatant was obtained as crude venom.Honey bees colonies , Phenyl methyl sulfonyl fluoride (PMSF), 1,10 Phenanthroline, bovine serum albumin (BSA), Diethylaminoethyl cellulose (DEAE-Cellulose), marker proteins, Sephacryl S-300, and thromboplastin were from Sigma Chemical . The other chemicals were of analytical grade. Human blood samples were obtained from the laboratory of medical center hospital of our institute.2 (5 μmol), and phenol red (80 μmol). The optical density was first recorded at 558 nm as a blank for each sample. Start the reaction by adding the enzyme solution, incubate for an hour at 37 °C, and then record the decrease in absorbance at 558 nm. One unit PLA2 activity is the amount of enzyme needed to hydrolyze 1 μmol phosphatidylcholine per hour at 37 °C [The reaction mixture of PLA2 activity assay consists of 2.5 ml 7.5 μmol Tris/HCl, pH 7.9 containing phosphatydylcholine (15 μmol), Triton X-100 (18 μmol), CaClat 37 °C .Apis mellifera lamarckii) venom extract was loaded on DEAE cellulose column (6× 2.4 cm i.d.) formerly equilibrated with 0.02 M Tris/HCI buffer, pH 7.8. Venom components were eluted with equilibration buffer containing NaCl gradients (0–1 M) with collection of 5 ml fractions. Fractions were monitored for PLA2 activity at 558 nm and that exhibiting PLA2 activity were collected, lyophilized, and utilized for further purification steps. These concentrated fractions were further loaded on Sephacryl S-300 column (142 cm × 1.75 cm i.d.) earlier equilibrated with 0.02 M Tris/HCI buffer, pH 7.8 with collection of 2 ml fractions. Fractions exhibiting PLA2 activity were stored at – 20 °C and thereafter utilized for studying homogeneity and characteristics of the purified PLA2.All experiments were performed at 4 °C. The crude  as a standard .g for 15 min at 4 °C to separate platelet-rich plasma (PRP) and platelet-poor plasma (PPP) [Blood specimens were collected in sodium citrate and centrifuged at 250×ma (PPP) . Assays ma (PPP) .The prothrombin time (PT) measures the plasma coagulation time at 37 °C in existence of tissue thromboplastin-calcium mixture. Mix 50 μl bee venom PLA2 with 50 μl plasma, incubate for 6 min at 37 °C then add 100 μl pre-warmed calcium-thromboplastin solution at 37 °C for determining the clotting time , 30.Tests were performed in triplicates unless stated otherwise and statistical analyses were performed in calculating average arithmetic mean and standard error (S. E.) .Apis mellifera lamarckii) was observed by PLA2 capability of hydrolyzing the phosphatidylcholine whereas the purification outlines exists in Table −1 PLA2 represented 8.9-folds and 38% yield. A native bee venom PLA2 mass of 16-kDa was deduced via its elution volume from the size-exclusion column.The purification of the PLA2 from the venom of the Egyptian honey bee  to know the suitable temperature for enzyme activity. Honey bee venom PLA2 enzyme showed its maximum activity at 45 °C . The highest activity of bee venom PLA2 was recorded at pH 8.0 Fig. a. The be °C Fig. b. The Kmine Fig. c.Fig. 32+, Ni2+, Fe2+, Ca2+, and Co2+ raised PLA2 activity, while Zn2+ and Mn2+ lowered it  pI at 10.5 . Most hymenopterous venoms PLA2 pI ranging from pH 9 to 12 [Bothrops leucurus snake venom PLA2 pI at 5.4 [Km value of 20 μM phosphatidylcholine  while calcium ion acted as enhancer effect on that of cobra venom [Bee venom components are important in pharmaceutical industry and drug formulations. The honey bee venom has high biological activity and is a better form for certain potential pharmacological sources . Bee ven columns ; and Sep columns . In this columns , who demfication . Egyptias 14 kDa , Europead 20 kDa  and Iran at 7.05  and EuroI at 5.4 . The EgyI at 5.4 . SimilarI at 5.4  and IbraI at 5.4 , low Km ra venom . EgyptiaIn conclusion, the main achievement of this study is the preparation of homogenous PLA2 from Egyptian bee venom by a straightforward purification procedure. Honey bee venom PLA2 could have a role in treating variety of diseases as anti-platelets aggregation and anticoagulant agent. Finally, the current study provides the bee venom PLA2 as a promising agent for developing novel anti-clot formation drugs in future.


Accordingly, inhibitors of peroxiredoxin 2 and catalase significantly decreased erythrocyte HbNO concentration. Conversely, steady-state levels of HbNO were preserved upon supplying erythrocytes with exogenous catalase. These findings support HbNO measurements as indicators of vascular oxidant stress and of NO bioavailability and potentially, as useful biomarkers of early endothelial dysfunction.Oxidative stress perturbs vascular homeostasis leading to endothelial dysfunction and cardiovascular diseases. Vascular reactive oxygen species (ROS) reduce nitric oxide (NO) bioactivity, a hallmark of cardiovascular and metabolic diseases. We measured steady-state vascular NO levels through the quantification of heme nitrosylated hemoglobin (5-coordinate-α-HbNO) in venous erythrocytes of healthy human subjects using electron paramagnetic resonance (EPR) spectroscopy. To examine how ROS may influence HbNO complex formation and stability, we identified the pro- and anti-oxidant enzymatic sources in human erythrocytes and their relative impact on intracellular redox state and steady-state HbNO levels. We demonstrated that pro-oxidant enzymes such as NADPH oxidases are expressed and produce a significant amount of ROS at the membrane of healthy erythrocytes. In addition, the steady-state levels of HbNO were preserved when NOX (e.g. NOX1 and NOX2) activity was inhibited. We next evaluated the impact of selective antioxidant enzymatic systems on HbNO stability. Peroxiredoxin 2 and catalase, in particular, played an important role in endogenous and exogenous H Reduin vivo and proposed it as a surrogate index of vascular NO availability -N-[4-(3-aminopropylammonio)butyl]-amino]diazen-1-ium-1,2-diolate, Spermine-NONOate, Enzo Life Science) for 45 min. A stock solution of the NO-donor was prepared from Spermine-NONOate powder dissolved in NaOH (0.01 M), and diluted in physiological solution (pH 7.4) before immediate addition to cells in hypoxia chamber. The final concentration of 50  μM was chosen based on the concentration-response curve as illustrated in 2, and the RBCs were frozen in calibrated tubes in liquid nitrogen every 15 min during next 45 min for low-temperature EPR measurements.Human blood daily collected was centrifuged and incubated at 37 °C in hypoxia workstation (INVIVO2 in RBCs (under continuous mixing) after reconstitution (50% of haematocrit) and Spermine-NONOate addition as described above. Then 10 μmol/L of H2O2 was added for 15 min incubation at 4% of O2 and the samples were frozen every 15 min (up to 45 min) for the EPR measurements.The antioxidant enzymes  were inhibited respectively by 10 μmol/L of 3-AT and by 25 μmol/L of ConoidinA for 30 min at 4% of O2.92300) at 1% of O2 and at room air (21% of O2). Then, different concentrations of Spermine-NONOate (10-20-50 and 100 μmol/L) were added for 45 min, samples were incubated at 1% of O2 in hypoxia chamber under continuous mixing (with Eppendorf Thermomixer) and afterwards frozen in calibrated tubes in liquid nitrogen for low-temperature EPR measurements.To study the accumulation of HbNO upon incubation with a NO donor, RBCs were collected from venous blood, washed and reconstituted at 50% of haematocrit with isotonic buffer. Erythrocytes were incubated with 50 units of non-PEGylated catalase  or non-PEGylated SOD  at 37 °C during 1 h in a hypoxia workstation  in Hank's Buffered Saline Solution  and incubated for 30 min at room temperature and at 4% of O2 with the following reagents: 3-AT (10 μmol/L), ConoidinA (25 μmol/L), catalase (50 units/ml). CM-H2DCFDA  at 1 μmol/L was added to each sample for additional 25 min incubation at 37 °C and 4% of O2. Samples were washed twice with HBSS and at least 10000 events were analyzed for fluorescence on FACSCantoII™ flow cytometer at room oxygen. For the experiments with antioxidant inhibitors, 10 μmol of H2O2 was added after a first reading and a kinetic curve of ROS formation was measured every 5 min until 45 min for each sample.Isolated human erythrocytes were re-suspended . The SAS 9.4 software was used for mixed model statistical analysis. Median fluorescence intensity (MFI) values were obtained from at least 10000 events analyzed for fluorescence on FACSCantoII™ flow cytometer and normalized in each experiment to a baseline of control unstained samples. Differences were considered statistically significant with a P-value < 0.05.33.13.1.12.- formation using the EPR spin probe, CPH in ghost membranes isolated from healthy human RBCs. Erythrocyte membrane samples were pretreated with vehicle or SOD or NOX inhibitors, or with l-NAME, an inhibitor of nitric oxide synthase (NOS) . VAS2870 preferentially inhibits NOX2 with Ki value of 770 nmol/L. Inhibition of NOX1 and/or NOX2 produced a significant reduction in membrane O2.- production by about 44%; these effects were comparable to that of SOD  B. SOD, ahrocytes ,18, we t samples B. We test of SOD B.Fig. 1E3.1.22 (venous oxygen level) where hemoglobin is mostly in T-state conformation amenable to HbNO complex formation. Isolated erythrocytes were pre-incubated with a NO donor and pre-treated with the NOX inhibitors, VAS2870 or GKT137831, or both, or with SOD before freezing samples for low temperature EPR measurements. The calculated HbNO concentrations and typical EPR spectra are shown in The erythrocyte membrane NOX1 and/or NOX2 generated a significant portion of ROS. Therefore, we tested the stability of HbNO after inhibition of these NOXs. Note that erythrocytes were incubated at 4% of O3.23.2.1RBCs have an extensive antioxidant system as demonstrated by our Western blot analysis illustrated on 3.2.22.- in erythrocyte membranes and preserved HbNO; however SOD activity was expected to proportionally increase the content of H2O2, another ROS source, as product of its dismutase enzymatic activity. We then investigated the potential role of the other intracellular antioxidant enzymes, as identified above, on ROS and HbNO contents in erythrocytes. In particular, we inhibited peroxideroxin2, the most abundant antioxidant enzyme expressed in these cells [50 = 23 μmol/L) [2O2. In absence of added H2O2, inhibition of Prdx2, but not catalase, produced an increase of intracellular ROS production in RBCs (96% increase at 20 min) compared with control cells. However, in RBCs exposed to exogenous H2O2, both inhibition of catalase and Prdx2 produced a strong increase of intracellular DCFDA signal, with catalase inhibition producing a doubling of the signal at 20 min . The HbNO EPR signal decreased with time significantly more in erythrocytes treated with 3-AT or ConoidinA by comparison with untreated RBCs , 3-AT  or ConoidinA .We have shown that SOD decreased the production of Ose cells  with Con μmol/L) ,21, whil μmol/L) . Fig. 3A2O2. To confirm this, we tested the effect of exogenous, non-PEGylated catalase on ROS accumulation (2O2 (2O2) catalase significantly decreased ROS accumulation intracellularly (by 37 ± 0.3% at 20 min compared to control samples)  A. Consissamples) B, as alsse Supp. B suggestse Supp. C.42.- in membrane extracts of human erythrocytes; ii. O2.- produced by NOX1-2 decreases steady-state levels of HbNO; iii. among the anti-oxidant armamentarium of RBCs, cytosolic peroxiredoxins (e.g. Prxd2) attenuate the constitutive intracellular production of oxidant species, while both Prdx and catalase cooperate to reduce the intracellular effect of exogenously applied H2O2; iv. the effect of extracellular (exogenously added), but also endogenous catalase  against degradation of HbNO points to extracellularly generated H2O2 as a significant oxidant species influencing steady-state HbNO levels.The main findings from our study can be summarized as follows; i. NOX1, NOX2 and uncoupled eNOS, are the main sources of O2.- [2O2 or another strong oxidant such as peroxynitrite (itself formed by the direct reaction between NO and O2.-), can produce heme in the ferryl form (Hb -Fe4+) or globin radicals that potentiate pro-oxidant activity in RBCs . Besidereviewed , NADPH oreviewed  but littreviewed . In our reviewed ,27. Withdiseases , as welldiseases ,29 and idiseases . NOX4 isses [2O2  and can ses [2O2 . These i2.- could directly react with NO , or its dismutation , which  [l-NAME B, or dirmembrane B. Accord2O2 itself is an oxidant and, according to conventional thinking, can migrate across cell membranes and, in the case of RBCs, could also react with hemoglobin, as well as other protein cysteine residues with low pKa [2O2 can also produce the highly reactive Fe(IV) ferrylhemoglobin, however such formation was shown to produce only small changes in the structure and property of the membrane of isolated erythrocytes [2O2, as previously shown in endothelium [2O2 on erythrocyte redox balance and HbNO?However, H low pKa , and inthrocytes , owing pothelium , is not 2O2, also produced from the dismutation of O2.- by the cytosolic SOD1, can be decomposed by catalase, glutathione peroxidase and by different isoforms of peroxiredoxin . We found immunoblotted signals for all these enzymes in the cytosolic lysates of human RBCs  increased intracellular H2O2, suggesting that Prdx2 mainly controls intracellular H2O2 formation [2O2) ROS detection  concentration of NONOate which may seem to deviate from (patho)physiological concentrations of endogenously produced NO. Note, however, that the final concentration of bioavailable NO in solution may differ from that calculated from the NO donor . Nevertheless, this concentration allowed us to efficiently test the dynamic formation of HbNO in intact erythrocytes with the optimal signal/noise ratio. It is also important to note that we observed reproducible HbNO signals in the range of 100–200 nmol/l in intact erythrocytes freshly drawn from normal volunteers, in absence of incubation with exogenous NO donors . While m2O2, the transport of which may be facilitated by Aquaporin-1 richly expressed in the erythrocyte membrane. This points to HbNO measurements as indicators of vascular oxidant stress, as well as NO bioavailability and potentially, as useful biomarkers of early endothelial dysfunction.In conclusion, our work shows that the steady-state level of HbNO is profoundly influenced by the redox balance between pro-oxidant and anti-oxidant enzymatic systems in human erythrocytes. In particular, we identified a significant contribution of extracellular H


Herein, Mn-based catalysts (MnCl2 and Mn) are adopted and doped into MgH2 to improve its hydrogen storage performance. The onset dehydrogenation temperatures of MnCl2 and submicron-Mn-doped MgH2 are reduced to 225 °C and 183 °C, while the un-doped MgH2 starts to release hydrogen at 315 °C. Further study reveals that 10 wt% of Mn is the better doping amount and the MgH2 + 10 wt% submicron-Mn composite can quickly release 6.6 wt% hydrogen in 8 min at 300 °C. For hydrogenation, the completely dehydrogenated composite starts to absorb hydrogen even at room temperature and almost 3.0 wt% H2 can be rehydrogenated in 30 min under 3 MPa hydrogen at 100 °C. Additionally, the activation energy of hydrogenation reaction for the modified MgH2 composite significantly decreases to 17.3 ± 0.4 kJ/mol, which is much lower than that of the primitive MgH2. Furthermore, the submicron-Mn-doped sample presents favorable cycling stability in 20 cycles, providing a good reference for designing and constructing efficient solid-state hydrogen storage systems for future application.Magnesium hydride (MgH Hydrogen, which occupies higher energy density 142 MJ/kg) 42 MJ/kg ,4. Unfor storage ,12,13. N storage ,16. To c storage ,20,21,22 storage ,25,26,27 storage ,33,34,352 to improve its hydrogen storage properties [2 was decreased by about 100 °C by doping TiF4 and the activation energy was lower by about 96 kJ/mol. Zhang et al. [2-NiCl2 composite and SEM tests exhibited that the addition of NiCl2 was conductive to decreasing the size of MgH2 grains and particles. Ismail et al. [3 into MgH2 to find that the desorption temperature of the MgH2-10 wt% FeCl3 composite was 90 °C lower than that of as-milled MgH2 and the activation energy for hydrogen desorption was also decreased from 166 kJ/mol to 130 kJ/mol. Mao et al. [2/NiCl2 sample could release 5.17 wt% H2 in 60 s at 300 °C and the dehydrogenation activation energy was decreased to 121.3 kJ/mol and 102.6 kJ/mol for MgH2/CoCl2 and MgH2/NiCl2 sample, respectively.Transition metal halides were easy to be obtained and doped to MgHoperties ,39,40,41operties  observedg et al.  successfl et al.  doped Feo et al.  revealed2 could greatly enhance the de/hydrogenation properties. As far as we know, studies about Mn-based catalysts have rarely been researched, thus, it is urgent and interesting to explore the catalytic effect of MnCl2 for the reversible hydrogen storage performance of MgH2. However, there are still some shortcomings about the doping of transition metal halides. On one hand, the really doping amount of transition metal atoms are restricted because of the heavy halogen atoms. On the other hand, there would be a deadweight that Mg may react with halogen elements to form MgCl2 or MgF2 which could affect hydrogen capacity and absorption/ desorption rates [According to the above references, it can be concluded that doping transition metal halides into MgHon rates .2 was investigated and based on the microstructure evidence, submicron-Mn was successfully synthesized via a simple wet chemical method and doped directly to MgH2 to further enhance the hydrogen storage properties of MgH2. Moreover, its catalytic mechanism was explored and discussed in detail.In this work, the catalytic effect of MnCl2) was purchased from Sinopharm Chemical Reagent and Mn powders was commercially purchased from Aladdin Industrial Corporation. Submicron-Mn particles were prepared by a wet-chemical ball milling method. At first, 4 g Mn powders , 12 mL heptane , 0.6 mL oleic acid , 0.2 mL oleylamine , and 240 g balls were mingled in a home-made stainless steel jar under 0.1 MPa of Ar. The mixture was milled at a speed of 400 rpm for 60 h in the planetary ball mill . The treated slurry mixed with another 15 mL heptane, 1 mL oleic acid, and 1 mL oleic acid was then placed in a centrifuge tub. In addition, ethanol was used to centrifuge and wash the mixed solution eight times to remove larger particles and residual organic solvent. Finally, Mn submicron particles (submicron-Mn) can be acquired after vacuum-drying at room temperature for 12 h.Powders of manganese chloride  was placed at 380 °C under the hydrogen pressure of 6.5 MPa to absorb hydrogen for 2 h. The second step was to ball-mill the processed samples at 450 rpm for 5 h. After repeating the above hydrogenation heat treatment, MgH2 can be finally acquired.The MgH2 + x wt% submicron-Mn  and MgH2- MnCl2 composites were prepared by mechanical ball milling at 450 rpm for 2 h under 0.1 MPa of Ar . In order to avoid oxidation and contamination, all samples were handled and transferred in an Ar-filled glove box  where the oxygen /water concentration was kept less than 0.1 ppm.The MgH−1 in a sealed stainless steel reactor. For hydrogenation, the samples were gradually heated from room temperature to 400 °C at an average rate of 1 °C min−1 under 3 MPa H2. For isothermal measurements, the samples were first heated up to the desired temperature and then keeping the temperature constant in the whole test. In order to get the exact values of hydrogenation capacity, the second dehydrogenation measurements were also conducted to verify the accuracy of the values. Moreover, controlling the hydrogen pressure for de/hydrogenation tests well is also important, the isothermal absorption tests were performed at various temperatures under 3 MPa while the isothermal desorption performance was tested at different temperatures under hydrogen pressure below 0.001 MPa.X-ray diffraction (XRD) analyses of all samples were performed on an X’Pert Pro X-ray diffractometer  with Cu K alpha radiation at 40 KV, 40 mA to detect the phase compositions. To avoid air and water contamination, a special container was adopted for transferring and scanning samples. A scanning electron microscopy  with an energy dispersive spectroscopy (EDS) was performed to further characterize morphologies and element distribution of the samples. The hydrogen absorption and desorption properties were tested in a Sieverts-type apparatus. During testing non-isothermal hydrogen desorption properties, about 75 mg sample was heated to 450 °C at a heating rate of 2 °C min2 on the hydrogen storage properties of MgH2, 5 wt% MnCl2 was ball milled with MgH2 to prepare the MgH2 + 5wt% MnCl2 composite, and temperature-programmed desorption (TPD) tests were conducted from room temperature to 450 °C, the results are shown in 2 began to release hydrogen from 315 °C and about 7.45 wt% hydrogen could be desorbed after the non-isothermal test. It is clear that the dehydrogenation temperature shifts to lower temperature after doping MnCl2 powders, which started to release hydrogen at 230 °C and about 6.8 wt% hydrogen could be attained when heating up to 350 °C. XRD measurements were also used to shed light on the microstructure evolution in the desorption process. 2 still dominates the diffraction peaks for MgH2-MnCl2 sample and no apparent new phases appeared during the ball-milling process, indicating that it was only a physical mixture after the ball milling process. After dehydrogenation, it is interesting that the signal of Mn phase emerged at 43° besides Mg phase. It is much likely that MnCl2 reacted with MgH2 during the dehydrogenation process, just as reported in other transition metal halides-modified MgH2 systems [2. In order to confirm this conjecture, submicron-Mn particles were further synthesized and a series of tests were performed.To investigate the catalytic effect of MnCl systems ,44. Ther2.The as-prepared Mn particles were analyzed by XRD and SEM measurements, as shown in 2, isothermal and non-isothermal dehydrogenation tests were operated. As a comparison, the purchased Mn was also doped into MgH2. The non-isothermal dehydrogenation curves in 2 + 5wt% Mn composite was 225 °C, 5 °C, and 90 °C lower than that of MgH2 + 5wt% MnCl2 composite and additive-free MgH2, respectively. Just as expected, after submicron-Mn was doped to MgH2, MgH2 + 5wt% submicron-Mn composite began to release hydrogen at 183 °C, superior to purchased Mn and MnCl2. In order to figure out the best doping amount, different amounts of submicron-Mn were ball-milled with MgH2 and further isothermal and non-isothermal measurements were conducted on the MgH2 + submicron-Mn composites. It could be clearly seen from 2 + 5 wt% submicron-Mn composite possessed onset dehydrogenation temperatures of 183 °C, about 132 °C lower than that of prepared MgH2. As for the MgH2 + 10 wt% submicron-Mn and MgH2 + 15 wt% submicron-Mn composites, the onset desorption temperatures further decreased to 175 °C and 165 °C, respectively. When the temperature rose to 350 °C, about 6.8 wt%, 6.5 wt%, and 6.1 wt% H2 could be obtained for the MgH2 + 5 wt% submicron-Mn, MgH2 + 10 wt% submicron-Mn, and MgH2 + 15 wt% submicron-Mn samples, respectively. Further isothermal dehydrogenation measurements of the above three samples are performed at 275 °C. 2 was desorbed in the first 10 min for the MgH2 + 5 wt% submicron-Mn composite. For the MgH2 + 10 wt% submicron-Mn and the MgH2 + 15 wt% submicron-Mn samples, the values increased to 6.1 wt% and 6.0 wt%. On the contrary, the pristine MgH2 sample could hardly release hydrogen under the same condition.To witness the modification impact of the as-synthesized Mn submicron particles on MgH2. Moreover, the initial dehydrogenation temperature did not decrease obviously after increasing the doping amount of catalyst. From a comprehensive perspective of the dehydrogenation temperature and capacity, the MgH2 + 10 wt% submicron-Mn was chosen for further study. 2 + 10 wt% submicron-Mn composite at 250 °C, 275 °C, and 300 °C, respectively. The MgH2 + 10 wt% submicron-Mn composite could quickly release 6.6 wt% hydrogen in 8 min at 300 °C . At 275 °C, this sample could desorb the same amount hydrogen with 20 min. Furthermore, about 6.0 wt% H2 could be acquired at 250 °C.According to the results of TPD curves, it could be concluded that the addition of submicron-Mn could remarkably improve the hydrogen desorption kinetics of MgH2+ submicron-Mn composites were also investigated. As shown in 2 + 10 wt% submicron-Mn sample could absorb H2 even at room temperature and about 5.5 wt% hydrogen could be re-absorbed when heating up to 250 °C. However, the dehydrogenated MgH2 sample sluggishly took up hydrogen from 186 °C. The hydrogen absorption curves of MgH2 + 10 wt% submicron-Mn at relatively low temperature were also performed, shown in 2 + 10 wt% submicron-Mn sample still absorbed 1.8 wt% hydrogen within 40 min. When the temperature went up to 75 °C, the hydrogen uptake of the submicron-Mn-doped sample amounted to 2.3 wt% under the same condition. After hydrogenation at 100 °C for 40 min, the fully dehydrogenated composite could absorb 3.2 wt% hydrogen. As a comparison, non-isothermal hydrogenation measurement of MgH2 sample were also conducted  model for the gas-solid reaction were adopted to simulate the evolution of kinetics [α is the fraction of Mg converted to MgH2 with time, k is an effective kinetic parameter, and n is the Avrami exponent. The numerical values of n and nlnk carried out by fitting the JMAK plots are shown in In addition, the Ea values of the hydrogenation reaction were calculated to further explore the improved kinetics of hydrogenation for MgHkinetics ,47. Figukinetics .ln[−ln. Compared with our previous study [2 + nano-Fe samples, which had an evident decrease in the first 20 cycles. In general, the degenerating cycling properties caused by that MgH2 particles tend to grow and aggregate during the process of thermolysis [2; other technics should still be explored to improve the cycling properties.In order to realize the practical application of hydrogen energy, preserving long-term kinetics is considered as one of the important indexes to evaluate the practicability for hydrogen storage materials. In this case, as shown in us study , the cycrmolysis . Thus, a2 and Mn particles were doped as catalysts into MgH2 to improve its hydrogen storage properties and the submicron-Mn particles exhibited superior catalytic effect. The MgH2 + 10 wt% submicron-Mn composite started to release hydrogen at 175 °C and about 6.6 wt% hydrogen could be obtained within 8 min at 300 °C. For absorption performance, the dehydrogenated MgH2 + 10 wt% submicron-Mn sample began to absorb H2 at room temperature the completely dehydrogenated sample could assimilate 3.0 wt% H2 within 30 min under 100 °C while the dehydrogenated MgH2 needed a high temperature of 210 °C to absorb the same amount of H2. Besides, the Ea of hydrogen absorption of MgH2 was reduced to 17.3 ± 0.4 kJ/mol because of the addition of submicron-Mn. Moreover, the MgH2 + 10 wt% nano-Mn composite exhibited good cycling performance that 89% of initial hydrogen could still be maintained after 20 cycles.In summary, MnCl


P2X1, P2X4, P2X5, P2X7, and PANX1 were expressed at the mRNA level in bovine PMN. Additionally, NEFA-induced NET formation was completely abolished with exposure to NF449, a P2X1 antagonist, and partially inhibited by treatment with etomoxir, an inhibitor of fatty acid oxidation (FAO). Our results suggest that OA and LA induce NET formation and ATP release via PANX1 and activation of P2X1. These new data contribute to explaining the effects of NEFA high concentrations during the transition period of dairy cattle and further understanding of pro-inflammatory effects and outcome of postpartum diseases.Non-esterified fatty acids (NEFAs) such as oleic acid (OA) and linoleic acid (LA) are associated with a higher incidence of infectious diseases such as metritis and mastitis during the bovine peripartum. Fatty acids can induce an increase in the release of ATP, and changes in the expression levels of purinergic receptors in bovine polymorphonuclears (PMN) during peripartum have also been reported. PMN respond to inflammatory processes with production of ROS, release of proteolytic and bactericidal proteins, and formation of neutrophil extracellular traps (NETs). NETs formation is known to require ATP production through glycolysis. Studies have shown that the above-mentioned metabolic changes alter innate immune responses, particularly in PMN. We hypothesized that NEFAs induce the formation of NETs through ATP release by Pannexin 1 and activation of purinergic receptors. In this study, we found that OA and LA induce NET formation and extracellular ATP release. Carbenoxolone, a pannexin-1 (PANX1) inhibitor, reduced OA- and LA-induced ATP release. We also found that  In the transition period of dairy cows, which includes 3 weeks pre- and 3 weeks postpartum, there is a high demand for nutrients and a decrease in feed ingestion, producing a transient negative energy balance . This pein vitro. For instance, OA is reported to induce intracellular calcium mobilization, MAPK phosphorylation, superoxide production, and release of granules containing CD11b and MMP-9 (Polymorphonuclear leukocytes cells (PMN) are the principal cell type of the host innate immune system and are regarded as the first line of defense against pathogens . PMN kilnd MMP-9 , 16. Mornd MMP-9 . HoweverP2X7, P2Y2, and P2Y11 was increased, whereas that of pannexin 1 (PANX1) was decreased, in bovine PMN 3 days postpartum  and according to the current Chilean Animal Protection Laws.Four Holstein Friesian heifers with body weights of 280–310 kg from the herd of the University Austral of Chile were used for the experiments. The heifers were fed twice a day with 1.0 kg of commercial concentrate, Cosetan® , and grazed in naturalized pastures composed of g for 20 min at 20°C. The plasma and buffy coat layers were aspirated and discarded, while the remaining red blood cells and PMN were resuspended in cold Hank's Balanced Salt Solution . Blood was transferred to Falcon tubes (15 ml) and centrifuged again at 1,000 × g for 20 min at 20°C, and the remaining phlogistic layer was removed by aspiration with a Pasteur pipette. The erythrocytes were then separated twice by rapid hypotonic lysis with a cold, aqueous phosphate buffer solution (5.5 mM NaH2PO4 and 8.4 mM HK2PO4 at pH 7.2) in a 2:1 ratio. Next, isotonicity was restored with a solution of hypertonic phosphate  in a 1:1 ratio, and then centrifuged at 600 × g for 10 min at 20°C. The PMN pellet was resuspended and washed twice with cold HBSS, being centrifuged each time at 500 × g for 10 min at 20°C. Finally, the pellet was resuspended in 10 ml of cold HBSS and 100 μl of cells was incubated with 5 μM propidium iodide  in HBSS with Ca2+ for 5 min at room temperature (RT). After incubation, 900 μl of HBSS with Ca2+ was added, and purity, counts, and viability were assessed using flow cytometry .Blood was collected aseptically by jugular venipuncture into BD Vacutainer® tubes  containing acid citrate dextrose (ACD). The tubes were gently shaken for 5 min and then centrifuged at 1,000 × 6) were suspended in HBSS with Ca2+ (0.897 mM) and exposed to 1 μM NF449   was added. NETs were quantified using 485/520 nm excitation/emission wavelengths in a Thermo Scientific Varioskan Flash  and expressed as relative fluorescence units (RFUs). Measurements were performed in duplicate.PMN  , 10 μM ctol, UK) , 28, 0.1tol, UK) , 0.001–1tol, UK) , 10 μM Gtol, UK) , 16, andtol, UK)  for 15 mtol, UK)  for 60 m5) were preincubated with NF449 (1 μM) or carboxenolone (10 μM) for 15 min at 37°C, or with 10 μM etomoxir for 60 min at 37°C, and then stimulated with 300 μM OA or LA for 30 min. The PMN were fixed in a 2% paraformaldehyde solution for 30 min at RT, followed by blocking with a 1% BSA solution for 2 h, and incubation with an anti-histone H4 (citrulline 3) antibody  overnight at 4°C. The next day, samples were washed three times with sterile HBSS and incubated with an Alexa Fluor 405- or Alexa Fluor 488-conjugated anti-rabbit secondary antibody  for 2 h at RT. To visualize the nuclei, the covers were stained with PicoGreen (1:200 in HBSS with Ca2+) or Sytox Orange  for 30 min at RT. Images were acquired with a confocal microscope .PMN (2 × 105) were resuspended in 250 μl of HBSS with Ca2+ and treated in two sets of experiments. In the first set, PMN were stimulated with 200 μM OA or LA, or vehicle (0.088% DMSO) for 0, 15, 30, or 60 s to obtain the time of the maximum ATP release for both NEFAs . In the second set, the PMN were treated with 10 μM carbenoxolone (CBX) or vehicle (0.01% DMSO) for 15 min at 37°C and then were stimulated with 300 μM OA or LA, or vehicle (0.088% DMSO) for 15 s . Immediately after finishing the time of stimulation, the PMN were maintained on ice for 5 min. Subsequently, the cells were centrifuged at 600 × g for 5 min at 4°C. A 10-μl aliquot of the supernatant was incubated in 100 μl of the kit mix for the determination of ATP levels, according to the manufacturer's instructions. Finally, the samples were incubated at RT for 15 min, and the luminescence was measured using Varioskan Flash (Thermo Fisher Scientific). The data were normalized using the mean of 60 s of measurement of the control group.ATP levels were determined using a commercial ATP Determination Kit . PMN . Samples were treated with Turbo DNase-Free (Thermo Fisher Scientific). For cDNA synthesis, 500 ng of total RNA was reverse-transcribed using M-MLV Reverse Transcriptase Kits . Real-time PCR assays were performed using Takyon Rox SYBR® MasterMix dTTP Blue  and primers specific for bovine P2X1–7, PANX1, gap junction protein alpha 1 (GJA1), and housekeeping ribonucleoprotein S9 (RPS9). The primers used for the PCR reaction were as follows: P2X1 forward 5′-CTGTGCAGAGAACCCGGAAG−3′ and reverse 5′-CGTTGAAGGCCACACACTTG-3′; P2X2 forward 5′-GCGTTCTGGGACTACGAGAC-3′ and reverse 5′-CACGATGAACACGTACCACAC-3′; P2X3 forward 5′-CTACTTCGTGGGGTGGGTTT-3′ and reverse 5′-ATGACTCGGTTGGCATAGCG-3′; P2X4 forward 5′-GGTGCGTGTCATTCAATGGG-3′ and reverse 5′-AAAGCAGGCTTTGGCACTTC-3′; P2X5 forward 5′-GGGAGTGGGGTCTTTCTTCTG-3′ and reverse 5′-TTCACCTTTCCGTTCCCCTG-3′, P2X6 forward 5′-AACTTCAGGACAGCCACACA-3′ and reverse 5′-TGACCAGGATGTCGAAACGG-3′; P2X7 forward 5′-GCTGCAGCTGGAATGATGTTT-3′ and reverse 5′-AAAGAGCACCACGTGGAAGAG-3′; PANX1 forward 5′-TTGACTTGAGAGACGGTGCC-3′ and reverse 5′-TGGCTTTCCTGTGAACTTTGC-3′; GJA1 forward 5′-GAGTGCCTGGGCTTGCTTTT-3′ and reverse 5′-TTGCCTGGGTACTGCTCTTTCT-3′; and RSP9 forward 5′-GCTGACGCTGGATGAGAAAGACCC-3′ and reverse 5′-ATCCAGCACCCCGATACGGACG-3′. The following conditions were used on a StepOne Plus real-time PCR system (Thermo Fisher Scientific): one cycle at 95°C for 10 min and 35 cycles of 95°C for 15 s, 60°C for 30 s, and 72°C for 30 s. This was followed by analysis of dissociation (melting) curves to ensure primer specificity. Relative changes in gene expression were determined by the ΔCT method , and the efficiency was calculated as [10(−1/slope)]. To P2X1 R2 = 0.96; Slope = −3.30; % efficiency = 100%, and efficiency was 2.01. To P2X4 R2 = 0.99; Slope = −3.40; % efficiency = 95%, and efficiency was 1.97. To P2X5 R2 = 0.96; Slope = −3.50; % efficiency = 110%, and efficiency was 1.93. To P2X7 R2 = 0.99; Slope = −3.03; % efficiency = 110%, and efficiency was 2.14. Finally, to PANX1 R2 = 0.99; Slope = −3.30; % efficiency = 100%, and efficiency was 2.01.Total RNA was isolated from 5 × 10T method . The effp-value < 0.05 was considered significant.The results are illustrated in bar graphs or dot plots as means ± S.E.M. of four independent experiments. One-way analysis of variance (ANOVA) was performed, and Fisher's LSD multiple comparison test was applied, using a significance level of 5%. When assumptions of normality or homogeneity of variance were not met according to the Shapiro–Wilks or Brown–Forsythe test, respectively, Kruskal–Wallis ANOVA and Dunn's multiple comparison test were used. All statistical analyses were performed using GraphPad Prism v7.0 . A Exposure to 200 and 300 μM OA  or LA F increaseTo assess the mechanism associated with OA- or LA-induced NET formation, we measured the concentrations of extracellular ATP released within 60 s by PMN stimulated with 200 μM of both fatty acids. Our results showed that OA and LA induced a peak of ATP release between 15 and 30 s .PANX1 mRNA was expressed in bovine PMN  channel receptors with differing functions in immune cells .PA, CM, RB, and MH designed the experiments. PA, CM, JQ, IB, KÁ, GD, AT, and CH performed the experiments. PA and CM prepared the manuscript. PA, CM, MH, and RB analyzed the data. All authors have read and approved the final version of this manuscript.The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.


Many reports indicate that in the cell S100A6 has an influence on the organization of actin filaments, but so far no direct interaction between S100A6 and actin has been shown. In the present study we investigated binding of S100A6 to actin and the actin–tropomyosin complex. The analyses were performed on G- and F-actin and two tropomyosin isoforms—Tpm1.6 and Tpm1.8. Using purified proteins and a variety of biochemical approaches we have shown that, in a Ca2+-bound form, S100A6 directly interacts with G- and F-actin and with tropomyosin, preferentially with isoform Tpm1.8. S100A6 and tropomyosin bind to the same population of filaments and the presence of tropomyosin on the microfilament facilitates the binding of S100A6. By applying proximity ligation assay we have found that in NIH3T3 fibroblasts S100A6 forms complexes both with actin and with tropomyosin. These results indicate that S100A6, through direct interactions with actin and tropomyosin, might regulate the organization and functional properties of microfilaments.S100A6 is a low molecular weight Ca S100A6 gene expression is cell specific and, in vitro, can be induced by multiple agents, among others, by stress factors2. The protein is present in different mammalian tissues and cells. Particularly, its high expression is found in mouse stomach and chicken gizzard4. As to cell distribution, the protein is expressed mainly in fibroblasts and epithelial cells5. Many reports which appeared up to now have shown that S100A6 plays multiple cellular functions6. For instance, it has been demonstrated, that S100A6 overexpression leads to higher cell proliferation rate and sensitizes cells to apoptosis8. On the other hand, diminished level of S100A6 not only inhibits proliferation but also alters cell morphology9. In particular, S100A6 deficiency in fibroblasts has led to changes in the shape, adhesion and motile properties of these cells, probably due to rearrangements in actin stress fiber architecture and/or reorganization of microfilaments associated with tropomyosin10.S100A6 is a Ca12. The isoforms segregate to different cell compartments and diversify microfilaments’ properties such as stability, dynamic polymerization and depolymerization, interactions with myosin motors and other actin-binding proteins14. S100A6 was shown to interact in vitro with smooth muscle tropomyosin15, therefore it is possible that changes in microfilament architecture observed in S100A6-deficient NIH3T3 fibroblasts were coordinated by tropomyosins. Although many reports have indicated that S100A6 has an influence on the rearrangement of actin filaments, so far, no direct interaction of S100A6 with actin was clearly demonstrated. It was also not known whether tropomyosin affects S100A6 interaction with microfilaments.Tropomyosins are actin regulatory proteins, which polymerize on both sides of the filament, stabilize the filament and control interactions of actin with other binding partners. In mammalian cells about forty tissue-specific and constitutive isoforms of tropomyosin are expressed16. For such studies we used purified proteins and applied different biochemical approaches. To check whether the interactions also occur in the cell we applied proximity ligation assay (PLA) in NIH3T3 fibroblasts.The goal of this work was to check whether S100A6 directly interacts with actin, tropomyosin and the actin–tropomyosin complex. Since, in the cell, actin is in a dynamic equilibrium between a monomeric (G-actin) and filamentous (F-actin) form, we examined interaction of S100A6 with both forms of actin, and with F-actin covered by various tropomyosin isoforms. We selected two non-muscle tropomyosin isoforms, Tpm1.6 and Tpm1.8, which co-localize with actin filaments in different cell compartments2+-dependent manner and eluted in buffer containing EGTA were analyzed by immunoblotting using anti-actin antibody. As it can be seen in Fig. To check whether S100A6 is capable of binding to actin, a pull-down assay with the use of CNBr-Sepharose-S100A6 affinity resin and a lysate prepared from NIH3T3 fibroblasts, was conducted. Proteins bound to the resin in a Ca2+-dependent. For that, purified G-actin was applied to the CNBr-Sepharose-S100A6 affinity resin in a buffer containing Ca2+. Proteins bound in the presence of Ca2+ were then eluted in a buffer containing EGTA. As it is shown in Fig. In the next step we established whether S100A6 interacts with G-actin and/or with F-actin, and if the binding is direct and Ca2+.Interaction between S100A6 and G-actin was additionally tested by an ELISA assay, which was done with the use of purified proteins. As illustrated in Fig. 2+. As shown in Fig. 2+ , but not in the absence of Ca2+  or in the control sample without EDC . Thus, the results from pull-down assay, ELISA and cross-linking indicate that the S100A6–G-actin interaction is direct and Ca2+-dependent.The interaction between S100A6 and G-actin was also examined by chemical cross-linking of both proteins using EDC in the presence or in the absence of Caapp), the fractional saturation of F-actin with S100A6 was drawn as a function of free S100A6  of S100A6 binding to F-actin. However, one should note that the experimental points obtained in the presence of Tpm1.8 are scattered, which affects fitting the data and decreases significance of the differences in the binding parameters.Most of the microfilaments present in various types of cells are associated with tropomyosinThe presence of S100A6 and each tropomyosin isoform on the actin filament at saturation point is illustrated in Fig. 2+, followed by elution of bound proteins with a buffer containing EGTA. As shown in Fig. To find the reason of the increased affinity of S100A6 to F-actin covered with Tpm1.8, we checked whether S100A6 directly interacts with tropomyosin in vitro and in NIH3T3 fibroblasts. For that, purified Tpm1.6 and Tpm1.8 were applied to CNBr-Sepharose-S100A6 affinity resin in a buffer containing Ca2+-dependent. In this assay binding of Tpm1.6 to S100A6 was very weak.To further analyze direct interaction between S100A6 and tropomyosin isoforms, an ELISA assay was performed using purified proteins. The results illustrated in Fig. 2+. Proteins bound in the presence of Ca2+ and eluted in a buffer containing EGTA were then analyzed by SDS-PAGE and immunoblotting using anti-tropomyosin antibodies . No such band was eluted from the CNBr-Sepharose-empty resin .Subsequently, we tested S100A6–tropomyosin interaction in cell lysate obtained from NIH3T3 fibroblasts. For that the protein lysate was applied to the CNBr-Sepharose-S100A6 affinity resin in a buffer containing Caies Fig. A. In theFinally, we checked whether S100A6 can form complexes with tropomyosin in the cell. For that we performed proximity ligation assay (PLA) in NIH3T3 fibroblasts Fig. B. The ob2+-binding protein belonging to the S100 family6. At present, this family consists of 25 members which exhibit tissue- and cell- specific distribution. S100 proteins, through interactions with various target proteins, are involved in many cellular functions such as regulation of Ca2+ homeostasis, proliferation, differentiation, apoptosis, inflammation, migration/invasion etc. Of note, S100A6 ligands include some components of the cytoskeleton such as tropomyosin, tubulin or CacyBP/SIP6, which suggests that S100A6 is involved in cytoskeletal organization and dynamics. This possibility is supported by the earlier observation that S100A6 deficiency in fibroblasts led to altered cell morphology, rearrangement of actin stress fibers, increased adhesion and motility of these cells10. Because cell shape and motility depend on actin filament dynamics, the above mentioned observations suggest that S100A6 is involved in the maintenance of microfilament architecture. However, it was not known whether S100A6 affects actin filaments through signaling pathways, activation of accessory proteins or through direct interactions with actin. Thus, in this work we examined this issue and showed for the first time that S100A6 binds to actin directly and in a Ca2+-dependent way. Moreover, our studies revealed that interaction between S100A6 and the actin filament is facilitated by tropomyosin—a regulatory protein, which controls actin interactions with many binding partners.S100A6  is a Ca19. Because this process is based on a transition from monomeric G-actin to filamentous F-actin, we checked whether S100A6 binds to G-, F- or both forms of actin. Pull-down assay, zero-length cross-linking and ELISA revealed that S100A6 binds to purified monomeric G-actin. Co-sedimentation assay demonstrated that S100A6 binds also to F-actin. For saturation of the filament, 5-molar excess of S100A6 was required, which indicates weak interactions with the apparent binding constant (Kapp) in the micromolar range, however, at this stage of investigations, the binding stoichiometry is unknown. Our experiments revealed a substantial degree of cooperativity of S100A6 binding to the filament. The cooperativity may arise from direct interactions between S100A6 molecules bound along the actin filament, which is possible in view of the S100A6 ability to form dimers20. An alternative explanation is that binding of S100A6 induces conformational changes in actin subunits, which are spread along the filament and facilitate S100A6 binding. Structural studies are necessary to get more information on the mechanism of F-actin-S100A6 interactions.We have shown that S100A6 forms complexes with actin in NIH3T3 fibroblasts. Pull-down assay with the use of CNBr-Sepharose-S100A6 affinity resin indicated that S100A6 bound to actin in cell lysates while the PLA assay revealed a dotted pattern of the S100A6–actin complexes visible throughout the cytoplasm of NIH3T3 fibroblasts. Fibroblasts are highly motile cells, in which actin filaments undergo dynamic polymerization and depolymerizationThe experiments with purified, G- and F-actin, proved that the binding did not require any accessory proteins. Also, our data showed that S100A6 binds both to β-actin present in the cell lysate and to purified skeletal muscle α-actin used in in vitro assays. This suggests that interaction of S100A6 with actin is a universal feature of this protein, and does not depend on the actin isoform, which may differ in various cell and tissue types.9. Because smooth muscle tropomyosin was shown to interact with S100A6 in vitro15, tropomyosin can be considered as a potential regulator of S100A6–actin interaction. Fibroblasts express numerous tropomyosin isoforms which segregate to different cellular compartments21. Tropomyosin isoforms, Tpm1.6 and Tpm1.8, selected for this study were shown to associate with different microfilaments. While Tpm1.6 segregates to contractile and non-contractile stress fibers22, Tpm1.8 is recruited to dynamic filaments at the base of ruffling membranes23 and de-branched filaments of lamellipodia24. Tpm1.6 and Tpm1.8 are products of the TPM1 gene and are identical in sequence except for the N-terminal segment. In spite of the similarity, these isoforms differ in the regulation of actin interactions with actin-binding proteins such as myosin and cofilin25.Earlier studies demonstrated a connection between S100A6 expression level and the specific architecture of the actin cytoskeleton. In fibroblasts transfected with anti-sense S100A6 retrovirus, microfilaments associated with tropomyosin were arranged diagonally across the cell, which was in contrast to the parallel organization in the control fibroblasts expressing normal level of S100A626. The results of CNBr-Sepharose-S100A6 affinity and ELISA assays performed with the use of purified tropomyosin isoforms demonstrated much stronger interaction of S100A6 with Tpm1.8 than with Tpm1.6. Furthermore, the CNBr-Sepharose-S100A6 affinity resin pulled-down only a low molecular isoform of tropomyosin from the NIH3T3 cell lysate. One has to note that in the cell lysate the mouse monoclonal anti-tropomyosin antibody, clone TM311, detected high and low molecular weight tropomyosin isoforms. There is a high probability that the low molecular weight isoform which was pulled-down from the cell lysate was either Tpm1.8 or Tpm1.9, because Tpm1.8 and Tpm1.9 (an isoform which differs from Tpm1.8 in the sequence of an inner segment) are the only two low molecular weight products of the TPM1 gene found in non-nervous cells16. This suggests that for specific interactions with tropomyosin S100A6 prefers the N-terminal segment encoded by the alternative exon 1b, which is present in low molecular weight tropomyosin isoforms.In this work we demonstrated that both tropomyosin isoforms facilitated binding of S100A6 to actin filaments by decreasing S100A6 concentration required for half-maximal saturation. This might be a result of conformational changes in F-actin brought about by binding of tropomyosin. Such possibility finds support in the results of our recent study showing tropomyosin-induced changes at the interface between actin subunits along and across the filament, which affected binding of cofilin-1 to actin. As mentioned above, in cells, Tpm1.8 associates mostly with highly dynamic microfilaments24. Because in this work we demonstrated that Tpm1.8 directly binds to S100A6, we propose that it recruits S100A6 to the filaments which undergo dynamic polymerization and depolymerization required for cell proliferation. As depletion of S100A6 in NIH3T3 fibroblasts resulted in profound stress fibers formation and stronger adhesion10, one can suppose that S100A6 is important for actin filament dynamic rearrangements. Specific isoforms of tropomyosin, e.g. Tpm1.8, can assist S100A6 in this function. We have also demonstrated that S100A6 binds to monomeric G-actin, therefore it is possible that binding of S100A6 to G-actin within the cell compartment containing dynamic actin filaments can enhance depolymerization and shortening of the filament. Because all interactions are Ca2+-dependent, S100A6 can be engaged in rebuilding of the actin cytoskeleton in connection with Ca2+ signaling pathways. This and the previous data are consistent with the idea that S100A6 regulates the architecture of the actin cytoskeleton, however, more studies are necessary to decipher exact mechanisms of this process.The observation that S100A6 forms complexes with actin and tropomyosin in NIH3T3 fibroblasts strongly supports the idea that interactions between these proteins regulate cytoskeletal organization in vivo27. In turn, the S100A8/S100A9 tetramer was found to promote F-actin cross-linking29. Ca2+-dependent interaction of S100A8/S100A9 with components of the cytoskeleton were shown to be involved in migration, degranulation or phagocytosis of activated monocytes and neutrophils. Another member of S100 family, S100A12, a protein constitutively expressed in neutrophils and inducible in macrophages and smooth muscle cells, modulates interaction between cytoskeletal elements and the cell membrane31. Also, S100A2, a protein mainly present in epithelial cells, was proposed to play a direct modulatory role in the organization of the actin cytoskeleton via a Ca2+-sensitive interaction with muscle and non-muscle tropomyosins32. Thus, regulation of the cytoskeleton organization by S100 proteins occurs via interaction with actin and actin binding proteins, and via interaction with tubulin. For instance, it has been reported that S100A8/S100A9 control microtubule reorganization during transendothelial migration of phagocytes29 while S100A6 affects microtubules in Wharton's jelly mesenchymal stem cells33. Taken together these data show that members of S100 family emerge as important regulators of the cytoskeleton and multiple cellular functions driven by the cytoskeleton dynamics.Our results place S100A6 among the family of actin-binding proteins and extend the list of S100 proteins, which interact with actin cytoskeleton. Another example of an S100 protein interacting with cytoskeletal components is S100A4, which was shown to bind non-muscle myosin heavy chain IIA, tropomyosin and actin, and through these interactions it was proposed to increase cell migration/invasivenessIn conclusion, our studies show that S100A6 may contribute to the regulation of the organization and functional properties of the microfilaments through direct interactions with actin and tropomyosin.10. Cells were grown in Dulbecco’s Modified Eagle’s Medium (DMEM) (Sigma Aldrich) supplemented with 10% fetal bovine serum (FBS) (Gibco), 100 U/ml penicillin, 100 µg/ml streptomycin (Sigma Aldrich) in an incubator at 37ºC and 5% CO2. Cells were passaged when confluent and used for experiments 24 h later. For the pull-down assay the protein lysates from NIH3T3 fibroblasts were prepared in buffer containing 20 mM Tris pH 7.5, 50 mM NaCl, 1 mM DTT, 2 mM CaCl2, 1 mM MgCl2 and protease inhibitors (Roche). Cells were homogenized using a syringe with a needle . Lysates were centrifuged and the supernatant fractions (300 µg of protein estimated using Bradford’s procedure) were applied to the CNBr-Sepharose-S100A6 or CNBr-Sepharose-empty resins.NIH3T3 mouse fibroblasts were cultured as described by Słomnicki and Leśniak34. S100A6 concentration was determined spectrophotometrically at 280 nm using the absorption coefficient 0.439 mg ml−1 cm−1 at 280 nm and molecular weight 10 000 Da.Purification of the S100A6 protein was performed as described by Słomnicki et al.35. It was kept in the form of G-actin in a buffer containing 2 mM HEPES, pH 7.5, 0.2 mM ATP, 0.1 mM CaCl2, 0.2 mM DTT and 0.02% NaN3 (G-buffer). The concentration of G-actin was determined spectrophotometrically by using the absorption coefficient 0.63 mg ml−1 cm−1 at 290 nm and molecular weight 42 000 Da.Actin was isolated from New Zealand rabbit back and hind leg muscle acetone powder according to Spudich and Watt36. The cells were harvested and the proteins were purified by ion exchange chromatography as described before37. Tpm1.6 had an Ala-Ser N-terminal extension, a modification that was previously introduced to increase actin affinity of this isoform38. Both tropomyosin isoforms were unacetylated at the N-termini. Concentration of the tropomyosin was determined spectrophotometrically at 280 nm using molar extinction coefficient 17 880 for Tpm1.6 and 8 940 for Tpm1.8. The coefficients were calculated from the amino acid sequences in Expasy (https://web.expasy.org/protparam/).Tropomyosin isoforms (Tpm1.6 and Tpm1.8) were expressed in bacteria BL21 (DE3) cells. Expression of Tpm1.6 and Tpm1.8 was induced with IPTG33. G-actin, Tpm1.6, Tpm1.8 or BSA (400 ng/well) were coated onto 96-well microtiter plates in 50 μl of Coating Buffer . After overnight incubation at 4 °C, the solution was removed and the wells were washed with TBS-T. The remaining adsorption sites were blocked with TBS  containing 10% BSA for 3 h at room temperature. Then, increasing amounts of S100A6 were added in 100 μl of Reaction Buffer  or , in the actin or tropomyosin-binding assay, respectively. After overnight incubation at 4 °C, the solution was removed. Following the washing procedure with TBS-T, polyclonal anti-S100A6 antibodies, diluted 1:500,  were added and incubation was carried out for 3 h at room temperature. This was followed by addition of a secondary goat anti-rabbit antibody (Sigma Aldrich) diluted 1:7,000. The analysis of bound antibodies was performed by colorimetric detection with TMB peroxidase EIA substrate kit (Bio-Rad). The absorbance at 450 nm was measured using a Bio-Rad microplate reader.ELISA assay was performed according to Jurewicz et al.39. S100A6 (30 μM) was mixed with G-actin (7.5 μM) in 15 μl of a buffer containing 2 mM HEPES, pH 7.6, 0.5 mM ATP and 0.2 mM CaCl2 or 2 mM EGTA. EDC (Sigma Aldrich) was added from fresh stock to 3 mM final concentration and activated by 2 mM NHS. In control reactions, S100A6 and G-actin were cross-linked separately or were incubated without the cross-linker. After 1 h incubation at 25 °C, the reaction was terminated by addition of DTT to a final concentration of 11 mM. Half of the reaction mixture was analyzed by 10% SDS-PAGE stained with Coomassie brilliant blue R250.Crosslinking with the use of a zero-length cross-linking reagent, EDC, was performed as described by Jurewicz et al.g before the assay and its concentration was measured in the resulting supernatant. Binding of S100A6 to 3 µM F-actin alone or saturated with 0.5 µM Tpm1.6 or Tpm1.8 was analyzed by addition of S100A6 at concentrations from 1–20 µM (corresponding to 0.3–6.7 S100A6:F-actin molar ratio). The assay buffer contained 10 mM Tris, pH, 7.5, 50 mM NaCl and 1 mM CaCl2. Protein mixtures were incubated for 15 min at room temperature and then ultracentrifuged for 1 h at 200,000×g. The proteins in the pellets and in supernatants were separated on 12% SDS-PAGE. The gels were stained with Coomassie brilliant blue G250. Protein bands were scanned in ChemiDoc MP (Bio-Rad) and their density was quantitated using the EasyDens software . The concentration of unbound S100A6 (free S100A6) remaining in the supernatant was calculated from the standard curve obtained on the basis of the density of S100A6 samples of known concentrations. The saturation curves were plotted as an increase in S100A6:actin densitometric ratios in the function of S100A6:F-actin molar. The binding curves were obtained by fitting the experimental points either to liner regression  or to hyperbolic equation (S100A6 binding to F-actin–tropomyosin complex) in SigmaPlot, version 12.5 . In order to compute the apparent binding constant (Kapp) and the Hill cooperativity coefficient (αH), the S100A6:actin densitometric ratios were normalized by dividing each S100A6:actin ratio by the maximal value of the ratio at saturation. The thus calculated fractional saturation of the actin filament with S100A6 was plotted as a function of free, unbound S100A6 concentration measured for each sample in the supernatant. The experimental points were fit to the Hill equation  but blocked with Tris, was prepared.The coupling of purified recombinant S100A6 protein to CNBr-Sepharose  was carried out according to the manufacturer instructions as described by Słomnicki et al.2 or in a buffer containing 10 mM Tris pH 7.5, 30 mM NaCl, 2 mM MgCl2, 1 mM CaCl2, respectively, and the samples were incubated on a rotating wheel for 1 h at 4 °C. The resins were extensively washed with the same buffer and in the case of tropomyosin additionally with this buffer but containing 150 mM NaCl, 500 mM NaCl and finally with 1 M NaCl. Then, proteins bound in a Ca2+-dependent manner were eluted with buffer containing 4 mM EGTA instead of CaCl2, precipitated with cold acetone and analyzed by SDS-PAGE. Gels were then stained with Coomassie brilliant blue R250.G-actin or Tpm1.6/Tpm1.8 (purified as described above) were applied to the CNBr-Sepharose-S100A6 affinity resin in a buffer containing 2 mM HEPES, 0.2 mM ATP and 0.2 mM CaCl2, 1 mM MgCl2 and protease inhibitors (Roche), and then applied to the resins as described above. After incubation on a rotating wheel for 2 h at 4 °C, the resins were extensively washed with this buffer and then with the same buffer but supplemented with 250 mM NaCl. The proteins bound in a Ca2+-dependent manner were eluted with buffer containing 4 mM EGTA instead of CaCl2. The elution fractions were precipitated with cold acetone and analyzed by SDS-PAGE and immunoblotting using anti-actin and anti-tropomyosin antibodies (see below).In the second experimental approach, protein lysate from NIH3T3 fibroblasts was prepared in buffer containing 20 mM Tris pH 7.5, 50 mM NaCl, 1 mM DTT, 2 mM CaCl40. Separated proteins were electrotransferred onto nitrocellulose membrane and identified using mouse monoclonal anti-actin-peroxidase , mouse monoclonal anti-tropomyosin antibodies, clone TM311 . After washing with TBS-T buffer  the blots were allowed to react with goat anti-mouse IgG (Jackson Immunoresearch Laboratories) secondary antibodies conjugated to horseradish peroxidase, diluted 1:10,000. After three washes with TBS-T and two washes with TBS blots were developed with the ECL chemiluminescence kit (Amersham Biosciences) followed by exposure against an X-ray film.Gel electrophoresis with 10%, 12% or 15% (w/v) polyacrylamide containing 0.1% SDS (SDS-PAGE) was performed by the method of Laemmli33. NIH3T3 fibroblasts were grown on glass cover slips previously coated with poly-l-lysine (50 μg/ml) for 24 h. The reaction with primary antibodies  1:60 and anti-actin  1:400 or anti-tropomyosin  1:400 was carried out for 2 h at room temperature. After washing, the incubation with oligonucleotide-conjugated antibodies: anti-rabbit PLA plus and anti-mouse PLA minus probes was conducted for 1 h at 37 °C in a humidity chamber. All following steps were performed according to the manufacturer protocol and with the reagents and media provided in the PLA kit. Immunofluorescence staining was analyzed under a Zeiss Spinning Disc Confocal Microscope (Carl Zeiss GmbH).For visualization of S100A6–actin and S100A6–tropomyosin complexes, a proximity ligation assay (PLA)  was appliedSupplementary information.


Thermoanerobacter italicus (Ta.it.I1). We show that Ta.it.I1 is highly active for ribozyme activity, forming linear hydrolytic intron products. Addition of purified IEP switches activity to the canonical lariat forming splicing reaction. We demonstrate that the Ta.it.I1 group IIC intron coordinates the progression of the forward splicing reaction through a π–π′ interaction between intron domains II and VI. We further show that branched splicing is supported in the absence of the IEP when the π–π′ interaction is mutated. We also investigated the regulation of the two steps of reverse splicing during intron mobility into DNA substrates. Using a fluorescent mobility assay that simultaneously visualizes all steps of intron integration into DNA, we show that completion of reverse splicing is tightly coupled to cDNA synthesis regardless of mutation of the π–π′ interaction.Group II introns are mobile genetic elements that perform both self-splicing and intron mobility reactions. These ribozymes are comprised of a catalytic RNA core that binds to an intron-encoded protein (IEP) to form a ribonucleoprotein (RNP) complex. Splicing proceeds through two competing reactions: hydrolysis or branching. Group IIC intron ribozymes have a minimal RNA architecture, and splice almost exclusively through hydrolysis in ribozyme reactions. Addition of the IEP allows the splicing reaction to form branched lariat RNPs capable of intron mobility. Here we examine ribozyme splicing, IEP-dependent splicing, and mobility reactions of a group IIC intron from the thermophilic bacterium  Group II introns are self-splicing ribozymes that reside within the genomes of bacteria  and the Group II introns are divided into three major classes based on IEP sequence and RNA secondary structural features: IIA, IIB, and IIC . While aStructural studies of group IIB introns have revealed that both protein and RNA tertiary interactions play pivotal roles in positioning DVI between the two steps of splicing. In the pre-second step of reverse splicing  DVI is positioned by RNA–RNA interactions with DIC, and protein–RNA interactions with the IEP . In thisThermoanerobacter italicus (Ta.it.I1) and its IEP can be readily reconstituted in vitro, and use this system to investigate the transition between the steps of both forward and reverse splicing. We demonstrate that the Ta.it.I1 group IIC intron requires a basic amino acid patch within the IEP to promote the formation of branched splicing products, and uses the π–π′ interaction to transition between the steps of the forward splicing reaction. We also find that mutation of the π–π′ interaction allows lariat formation independent of the IEP. We developed a fluorescent assay to investigate Ta.it.I1 retromobility into DNA. Using this assay, we show that the engagement of the RT domain of the IEP in cDNA synthesis mediates the transition between the steps of the reverse splicing reaction during Ta.it.I1 integration into DNA substrates.In this study, we show that a group IIC intron from the thermophilic bacterium 8-SUMO-IEP was expressed in E. coli by auto-induction  to assist ribozyme splicing and facilitate intron mobility into DNA targets. To reconstitute IEP-dependent splicing in vitro, an amino-terminal fusion of Hisnduction , followenduction A. The elactivity B. As a ced to WT B. The puTa.it.I1 intron RNA was prepared by in vitro transcription using T7 RNA polymerase . Many group II introns have limited activity in vitro, most likely due to RNA misfolding. Thus, the almost complete  group IIC intron demonstrated that NH4Cl and KCl monovalent ions retain the same ribozyme active site architecture  to prime cDNA synthesis . DNA substrate was then added to the RNP product in the presence or absence of dNTPs and incubated at 37°C. Incubation of the RNP in the absence of dNTPs primarily produced the lariat-3′ DNA exon mobility intermediate and free 5′ DNA exon products . We next investigated if the π–π′ interaction is also important for the progression of the reverse splicing reaction into DNA substrates. We did not observe any differences in reverse splicing activity of mutant RNPs compared to WT  could switch splicing preference from hydrolysis to branching  E. coli cells. Expression was performed with 1.5 L of Terrific broth auto-induction media  and treated with PMSF protease inhibitor at a 1/1000 dilution. The cells were lysed on ice by sonication: 15 bursts of 8 sec separated by 1-min rests. The cell lysate was cleared at 25,000g at 4°C, added to 2 mL Ni-IDA agarose beads (GoldBio) and allowed to bind in a conical tube while turning end-over-end for 30 min at 4°C. The resin was collected at 500g at 4°C and washed with 30 mL of lysis buffer. This process was repeated three times to preclean the resin prior to column loading, which was observed to significantly reduce RNase contamination of the IEP preparation. After prewashing, the resin was loaded onto a 20 mL column, washed with 30 mL of wash buffer , followed by a wash with 2 M NaCl, 20 mM Tris-HCl (pH 7.5), 5 mM β-mercaptoethanol to remove copurifying nucleic acids bound to the IEP. The YAAA mutant IEP was particularly prone to copurification with non-specific nucleic acids judged by OD 260 nm, which may have caused higher background in PicoGreen RT assays (see below). After rinsing the resin with lysis buffer, the protein was eluted with ∼15 mL of elution buffer . The IEP was desalted/buffer exchanged in desalting buffer  on a HiPrep Desalting FPLC column (AKTA). IEP was further purified by size exclusion chromatography using a Sephacryl S-200HR column. The final protein was concentrated at 4000g followed by addition of 5 mM DTT and glycerol to a final concentration of 50%. The IEP was stored at −20°C.pET11a His-SUMO-on media  containiTa.it.I1 IEP was diluted 1:5, 1:10, 1:20, 1:50, and 1:100 in dilution buffer . An amount 5 µL of each RT dilution was added to 20 µL of poly(rA) oligo(dT)18 substrate and incubated at 25°C for 30 min before quenching with 2 µL of 220 mM EDTA. An amount of 12.5 µL of each reaction was then transferred to one well of a 96 well plate where 86.5 µL of Picogreen reagent was added. An amount of 12.5 µL of dilution buffer was used as a negative control. Fluorescence was measured with a Synergy H4 Hybrid plate reader (BioTek). To normalize mutant Ta.it. RT activity to WT Ta.it. RT activity, the concentration of each RT was determined by a Bradford assay and each were diluted to 800 µg/mL. RT reactions were set up with 5 µL of 800 µg/mL RT (WT or mutant), to 20 µL of poly(rA) oligo(dT)18 substrate, and 2 µL of RNaseA and performed as described above.Ta.it.I1 specific 10× T7 buffer containing 5 mM MgCl2. After transcription and subsequent DNase I and Proteinase K digestion, the transcripts were filtered, concentrated, and buffer exchanged by several rounds of centrifugation at 4000g using an Amicon 100 kDa cutoff ultrafiltration centrifugal filter unit.RNA transcription was performed with sequence confirmed plasmid DNA and T7 RNA polymerase . During Ta.it.I1 WT RNA, 5 mM fresh DTT, 50 µL of splicing buffer  and made up to 100 µL volume with Millipore water. Splicing reactions were performed by preincubation of the splicing buffer with or without IEP for 5 min at the assay temperature and initiated by addition of the RNA. Hydrolytic splicing assays were performed without IEP under 100 mM MgCl2 buffer conditions. Maturase splicing assays were performed with ∼1.5× molar excess of Ta.it.I1 IEP. Reactions were phenol-CIA extracted and ethanol precipitated with 0.3 M NaOAc, pH 5.2 with a linear polyacrylamide carrier. Extracted RNA was resuspended in water and 2× formamide/2 mM EDTA loading buffer before heating at 95°C for 3 min. Samples were resolved on 4% polyacrylamide (19:1 acrylamide:bisacrylamide)/7 M urea gels, stained with ethidium bromide, and imaged on the Syngene G:Box imaging system.Standard splicing assays were performed with 1 µg of 2, 250 mM NH4Cl, 50 mM Tris-HCl (pH 7.5) buffer. A 10 µg Ta.it.I1 RNA IEP-assisted splicing reaction in pull-down buffer: 250 mM NH4Cl, 10 mM MgCl2, 1 mM β-mercaptoethanol, 50 mM Tris-HCl (pH 7.5). The reaction was added to the blocked beads and allowed to bind end-over-end at room temperature for 30 min. The resin was collected by centrifugation at 500g for 2 min and washed with 500 µL of pull-down buffer at room temperature for 15 min three times. Pulled-down RNPs were eluted with pull-down buffer supplemented with 300 mM imidazole. Eluted RNPs were phenol-CIA extracted, ethanol precipitated with 0.3 M NaOAc, pH 5.2 with a linear polyacrylamide carrier, resolved on 4% polyacrylamide/7 M urea gel, and stained with ethidium bromide.Ni-IDA-agarose beads (GoldBio) were blocked with 100 ng/µL BSA and 1 µM pMALc5x-Rev oligonucleotide in 10 mM MgClTCGGCGCGG, underlined position indicates the internal FAM label position.Mobility assays were performed with multiple different oligonucleotides labeled with fluorescent probes to eliminate the need for radioactivity. DNA substrate was designed with a 5′ end DNA stem–loop labeled with Cy5 and a 3′ end hairpin labeled with FAM. The sequence of the substrate, synthesized by IDT, was: Cy5GCAGTCTAAAAGTAATTTTAGACTGCTTTTTTATTTTTTCCGCGCTRNPs for mobility reactions were assembled by IEP-dependent splicing reactions under standard maturase conditions stated above at 37°C for 15 min. Mobility reactions were performed with 500 nM substrate with or without 1 mM dNTPs or dATP. Controls included were substrate alone and reactions treated with 50 U RNase H. All reactions and controls were incubated at 37°C for 15 min before phenol-CIA extraction and ethanol precipitation with 0.3 M NaOAc (pH 5.2). Reactions at higher temperatures did not support mobility, likely due to melting of the stem–loops in the DNA substrate. Time course assay reactions were stopped using stop solution . Extracted RNA was resuspended in water and 2× formamide/2 mM EDTA loading buffer before heating at 95°C for 3 min. Samples were then resolved on 4% polyacrylamide/7 M urea gels and scanned for fluorescence on a Typhoon imaging system (GE Amersham).


We show that 4.1N−/− mice were born at a significantly reduced Mendelian ratio and exhibited high mortality between 3 to 5 weeks of age. Live 4.1N−/− mice were smaller than 4.1N+/+ mice. Notably, while there were no significant differences in organ/body weight ratio for most of the organs, the testis/body and ovary/body ratio were dramatically decreased in 4.1N−/− mice, demonstrating selective effects of 4.1N deficiency on the development of the reproductive systems. Histopathology of the reproductive organs showed atrophy of both testis and ovary. Specifically, in the testis there is a lack of spermatogenesis, lack of leydig cells and lack of mature sperm. Similarly, in the ovary there is a lack of follicular development and lack of corpora lutea formation, as well as lack of secretory changes in the endometrium. Examination of pituitary glands revealed that the secretory granules were significantly decreased in pituitary glands of 4.1N−/− compared to 4.1N+/+. Moreover, while GnRH was expressed in both neuronal cell body and axons in the hypothalamus of 4.1N+/+ mice, it was only expressed in the cell body but not the axons of 4.1N-/- mice. Our findings uncover a novel role for 4.1N in the axis of hypothalamus-pituitary gland-reproductive system.Protein 4.1N, a member of the protein 4.1 family, is highly expressed in the brain. But its function remains to be fully defined. Using 4.1N Two highly conserved domains, the 4.1-ezrin-radixin-moesin (FERM) domain and the spectrin/actin-binding domain (SABD) are responsible for interactions of 4.1 family of proteins with the plasma membrane proteins and with the spectrin–actin cytoskeleton, respectively5. The third conserved domain in the 4.1 family is the carboxy-terminal domain (CTD). It has been shown that all members of the protein 4.1 family undergo extensive tissue specific alternate splicing generating multiple protein 4.1 isoforms. In the case of 4.1N, the 135 kDa 4.1N is the predominant isoform identified in the brain, whereas a smaller 100 kDa isoform is enriched in peripheral tissues, such as the enteric neurons of the intestinal tract and adrenal gland6.Protein 4.1N7. Furthermore, the binding of 4.1N to PIKE prevents its interactions with nuclear PI3K, which can influence the regulation of PI3K by NGF8. Interaction of protein 4.1N with dopamine receptor is required for the localization or stabilization of dopamine receptors at the neuronal plasma membrane9. Among dopamine receptors, protein 4.1N interacts specifically with D2 and D3 dopamine receptors. Moreover, the protein 4.1N-binding region of inositol 1,4,5-trisphosphate receptor type 1 (IP3R1) is necessary and sufficient for the localization of IP3R1 at the basolateral membrane domain in polarized MDCK cells10. Both the CTT14aa and CTM1 sequences of IP3R1 can bind to 4.1N 11, suggesting that the spectrin-actin binding domain of 4.1N can serve as a linker between IP3R1 and actin filaments. This actin filament-dependent regulation of IP3R1 diffusion may be important for the spatiotemporal regulation of intracellular Ca2+ signaling which in turn mediates neurite formation14. Interestingly, 4.1N does not interact with the IP3R in an epithelial cell line WIF-B, implying cell specific interaction between protein 4.1N and the IP3R and tissue-specific mechanism in shaping the pattern of Ca2+ waves in various cell types15. In primary hippocampal cultures, mouse 4.1N is enriched at the discrete sites of synaptic contact, co-localizing with the postsynaptic density protein of PSD95 and GluR1, suggesting a potential functional role for 4.1N as a component of the cytoskeletal architecture of excitatory synapses16. 4.1N also directly interacts with the GluR1 subunit of the AMPA receptor and co-localizes with AMPA receptors at excitatory synapses17.In vitro cell biological studies have identified multiple cellular functions for protein 4.1N. Anti-proliferative actions of NGF in PC12 cells has been shown to be due to binding of 4.1N to the nuclear mitotic apparatus protein (NuMA) following its translocation to the nucleus and preventing the mitotic actions of NuMA−/− mice and performed phenotypic characterization of these mice to begin to decipher the function of 4.1N in vivo. The 4.1N−/− mice were born at lower birth rate and showed slow growth with marked hypoplasia of reproductive system. These findings in conjunction with documentation of focal expression pattern of 4.1N in pituitary and hypothalamus suggest a specialized function for 4.1N protein in neuroendocrine system.In contrast to our understanding of the multiple functions of 4.1N in various cell types based on in vitro cell biological studies, function of 4.1N in vivo is yet to be defined. To address this issue, we generated 4.1N+/−) expressing ~ 50% of the normal level of 4.1N were fertile and phenotypically indistinguishable from wild type mice  cells generated by the International Gene Trap Consortium. A gene trap cassette with a reported insertion site between exon 1A and exon 2 was confirmed by genotype analysis using primer pairs Fig. , confirm6. Neither the 95 kDa or 135 kDa 4.1N isoforms were found in heart, liver, skeletal muscle and testis.The protein 4.1N undergoes tissue specific alternate splicing generating multiple isoforms. We compared the expression of 4.1N in different parts of brain and other tissues by Western blot analysis. Western blot analysis of hypothalamus tissue with 4.1N-specific polyclonal antibody revealed a doublet, with the predominant band migrating at 135 kDa and a less prominent lower molecular weight band. A similar pattern was seen for olfactory and cerebrum, whereas the lower molecular band was more prominent than the 135 kDa band in cerebellum  of males and 16 of 38 (43%) of females died between 3 to 5 weeks of age, whereas all their wild-type and hetero-type littermates were alive at that time. These young 4.1N−/− mice were marasmus, weak with movement defects, and with growth retardation. Detailed examination of the 4.1N−/− mice revealed that they were passive and when gently lifted by their tail, their movements were powerless relative to those of their wild-type counterparts. When tested for their ability to walk on a rod or on top of a container, some of the 4.1N−/− mice did not move at all while others were able to proceed for a short distance and tired quickly which was in marked contrast to wild type mice that were able to accomplish these tasks readily. The observed neurobehavioral defects in 4.1N−/− mice suggest a potential role for 4.1N in brain function.Because both male and female 4.1N−/− mice compared with wild-type littermates  in 4.1N+/+ mice. For the female, the 4.1N−/− adult ovary weight was 0.0073 ± 0.00085% of body weight, while it was 0.014 ± 0.0013% (p = 0.0005) in 4.1N+/+mice. We also noted that the 4.1N−/− mice displayed a mild decrease in liver weight compared with the control wild-type littermates (p < 0.05). No significant difference for the ratio of organ to body weight was noted for other organs including heart, lung, spleen and kidney in 4.1N−/− mice.As the mice aged, we noted a marked age-dependent decrease in size and body weight of 4.1Ntes Fig. . Body weles Fig. A and femles Fig. B at 3–5 les Fig. . We then+/+ Fig. . A detai−/− male mice with significant changes in mature sperm and in the number of sperm. The seminiferous tubules were reduced in diameter. Spermatogenic cells and spermatocytes were present, but spermatids were not observed. The number of mature spermatozoa was dramatically decreased , while this decrease in female knockouts was not significant (p = 0.222). Measurements of LH showed reduced levels in female knockouts compared with wild type (p = 0.0317), while differences in male mice was modest (p = 0.0620) between 4.1N−/− and 4.1N+/+ mice .To explore the potential causes for abnormalities in the reproductive system, we measured the serum levels of follicle-stimulating hormone (FSH) and luteinizing hormone (LH) in 4.1Nice Fig. . Analysip = 0.010, while t−/− mice, we examined pituitary tissue from 4.1N+/+ and 4.1N−/− mice. The most common cell types of the pituitary pars distalis are growth hormone (GH)-secreting cells. H&E staining show that acidophils are rather diffusely distributed in the secretory cells of normal mice. The basophilic gonadotrophs are responsible for the production of FSH and LH. We noted a decrease in the number and staining intensity of the chromophils, especially acidophils in 4.1N−/− mice. The secretory cells were smaller with fewer eosin granules in cytoplasm in 4.1N−/− mice compared with 4.1N+/+ mice  axis. Within the HPG axis, reproductive maturation and function is coordinated by the release of GnRH from a neuronal population in hypothalamus which also consists of other clusters such as GHRH secreting neurons−/− mice suggests that the absence of these hormone signals in the pituitary might be attributed to impaired secretion or migration of the releasing hormone granules from the hypothalamic to the pituitary via the hypophyseal-hypothalamic portal system (HHPS). Interestingly, immunohistochemical study confirmed that GnRH are, indeed, absent in the axons of hypothalamus neurons in 4.1N−/− mice. The GnRH immunoreactive signal was present both in neuronal cell body and axons with punctate pattern in the median eminence of the hypothalamus in 4.1N+/+ mice. In the 4.1N−/− mice, the expression of GnRH was not detected in the axons, although some reactivity was observed in the cell body of GnRH neurons, indicating that GnRH fails to migrate through axons in 4.1N−/− mice. Furthermore, the decrease of GnRH in pituitary can partly explain why 4.1N−/− mice have lower FSH and LH levels and developmental defects in the reproductive system. These findings suggest that 4.1N is required, directly or indirectly, for the hormone transport and secretion in neuroendocrine system.Our documentation of decreased GnRH and GHRH expression in pituitary pars intermedia in 4.1N−/− male and female mice is different. In contrast to the significant decrease in FSH level in 4.1N−/− male mice, the decrease in FSH levels in 4.1N−/− female mice was not statistically significant. This may in part be a reflection of the fact that serum FSH levels are generally lower in female rodents than in males27, and there is considerable individual variability in the serum FSH levels masking the difference between the mutant and wild-type mice. The individual variability and stability could account for the finding that the serum GH level showed no statistically significant difference between groups. However, the body weight of 4.1N−/− mice was reduced even in older mice and the expression of GHRH and GH as well as the number of secretory granules were decreased in 4.1N−/− pituitary, suggesting that the slow growth phenotype of 4.1N−/− mice is likely to be of complex etiolgoy.It is interesting to note that the extent of changes in FSH and LH levels in 4.1N28. The molecular identities of the complexes responsible for transporting axonal hormone are not well understood. A subclass of semaphorins such as Sema4B and Sema4C characterized by a PDZ-binding motif at their carboxy-terminus that mediates their interaction with the post-synaptic density protein PSD-95/ SAP90, SFAP75/Norbin and SEMCAP-1/GIPC have been implicated in clustering of semaphorins, axon guidance and neurite outgrowth31.HPG axis, whose abnormal hormone secretion or transport is associated with clinical feature involving delayed or absent puberty and reproductive dysfunction. Disorder in the development of the GnRH neuroendocrine system leads to absent GnRH secretion or migration, resulting in heterogeneous reproductive disorders such as congenital hypogonadotropic hypogonadism in humans with infertility or decreased fertility33. GluR1 and GluR3 are synthesized by GnRH neurones in preparation for the enhanced release of GnRH34. Our study expands the role of 4.1N to the hypothalamic-pituitary regions, providing further evidence that 4.1N maybe linked to the axonal cytoskeleton which is required to ensure the migration of releasing hormone. It is likely that protein 4.1N plays this role through direct or indirect interactions with other cytoskeletal proteins such as αII- and βII-spectrin which have been shown to contribute to the organization of neurofilaments36. However, precise interactions in specific axons need to be confirmed in future studies.In line with this pattern, 4.1N co-localizes with the postsynaptic density protein of PSD95 and GluR1 in primary hippocampal cultures and interacts with the GluR1 subunit of the AMPA receptor at excitatory synapses, suggesting that 4.1N may function as a component of the cytoskeletal architecture of excitatory synapses−/− mice exhibit unique phenotypes that include low birth rate, slow growth and hypoplasia of reproductive organs due to defects in the neuroendocrine system. Protein 4.1N could be an important component of axons, not only at the hypothalamus, but also in the pituitary region. The presence of 4.1N may be necessary for the transmission of releasing hormone at hypothalamic-pituitary-axon route. The 4.1N knockout mice are likely be a valuable tool for exploring 4.1N function in certain neuroendocrine disorders.In summary, 4.1N37. Briefly, embryonic stem (ES) cells, containing a gene trap cassette inserted downstream of exon 1A and upstream of exon 2 in the 4.1N gene, were obtained from the International Gene Trap Consortium (ES cell line FHCRC-GT-S12-6H1). ES cells were microinjected into blastocysts and implanted into recipient female mice to permit development of the embryos into chimeras at the University of California, San Francisco, transgenic facility, and subsequent breeding was performed to select founders with germ line transmission of the gene trap allele. The mice were initially bred and screened at the Lawrence Berkeley National Laboratory (LBNL) animal facility and then maintained at the animal facility of New York Blood Center under specific-pathogen-free conditions. All animal experiments and protocols were reviewed and approved by Institutional Animal Care and Use Committees from both the New York Blood Center and Lawrence Berkeley National Laboratory, and conducted in accordance with the New York Blood Center and Lawrence Berkeley National Laboratory guidelines and regulations. Mice were backcrossed more than 9 generations into a C57BL/6 background, and, in all experiments, comparisons were made between littermate wild-type (4.1N+/+) and homozygous mice (4.1N−/−). The genotyping was routinely performed by PCR as described below. In total, we examined forty-four 4.1N−/−  and the same number of wild-type and heterozygous mice. The mice were mainly divided into two groups: 3–5 weeks old and 6–9 weeks old. A few mice older than 9 weeks were also analyzed.4.1N knockout mice were generated using the similar method for generation of 4.1G knockout mice as described previouslyGenotyping was performed by multiplex PCR to identify mice inheriting the targeted 4.1N gene and to distinguish wild type and heterozygous mice from homozygous. The wild type 4.1N allele was specifically amplified using primers 4.1N-F1: 5′-TTGAGTTCCAGGACAGCCAGGGTTAC-3′ and 4.1N reverse: 5′-AGAGAGCCTTGAAAGAGACCAGACGAG-3′), while the mutant allele carrying the gene trap insertion was detected using primers 4.1N-F2 (5′-GTGATGAACTGATGGAAGGATAGCC-3′) and β-geo-Rev (5′-AGTATCGGCCTCAGGAAGATCGCAC-3′).Genomic DNA was extracted from mouse-tails. Mice were anesthetized with pentobarbital and perfused through the heart with PBS (pH7.4) containing 4% paraformaldehyde (PFA). After sacrificing the mice, various tissues or organs were rapidly dissected and fixed in 10% phosphate-buffered formalin solution. The testis were fixed in Bouin’s solution. The tissues were subsequently embedded in paraffin, and serial sections (4-μm thick) were cut and mounted on slides. The sections were stained with hematoxylin–eosin (HE). Immunohistochemistry was performed using various antibodies. The stained sections were examined microscopically. The number of testis and ovary analyzed were from 8 male and 6 female, separately. The other tissues were from 6 male and 6 female. Histopathology was assessed independently by two pathologists who were blinded to the different groups. Microscopic semiquantitative scoring was calculated on a scale of 0 to 4 as follows: 0 , 1 , 2 , 3 , and 4 .38.Polyclonal antibody against 4.1N was raised in rabbit using His-tagged recombinant N-terminal headpiece of 4.1N as antigen. The sequence of the peptide is unique to 4.1N and does not share any homology with 4.1R, 4.1G or 4.1B. The antibody was affinity purified on Sulfolink Coupling Gel . Antibody specificity was examined by Western blotting using recombinant 4.1R, 4.1G, 4.1N and 4.1B. As demonstrated in our previous published study, the 4.1N antibody was highly specific for 4.1N and did not recognize other members of the protein 4.1 family6. Briefly, tissues from adult 4.1N+/+ and 4.1N−/− were homogenized by sonication in ice-cold lysis buffer  in 1.5 ml tubes. After centrifugation at 900×g for 5 min, the supernatant was collected and the protein concentration measured by Bradford method using bovine serum albumin as standard. 30 μg of proteins were run on 8% Tris–glycine gels and subsequently transferred onto nitrocellulose membrane . Membranes were blocked for 1 h at room temperature in blocking buffer (PBS + 0.1% Tween-20 + 4% nonfat dry milk + 1% BSA) followed by overnight incubation at 4 °C with primary 4.1N antibody (1: 2000 dilution) and rabbit polyclonal anti-GAPDH antibody (1: 5000 dilution) in blocking buffer. After washes with 0.1% PBST, membranes were incubated for 1 h at room temperature with rabbit HRP-conjugated secondary antibody  diluted 1:5000 in blocking buffer. Following extensive washing, membranes were processed using chemiluminescent reagent .Western blot was performed as described previously6. Briefly, 4.0 μm thick sections were de-paraffinized in xylene and dehydrated using graded ethanol solutions. Endogenous peroxidase activity was quenched with peroxidase blocking reagent for 6 min. The tissue sections were pretreated with target retrieval solutions of high pH (for 4.1N immunostaining) or with low pH  and heated in microwave for 15 min. Following three washes with PBS, the sections were incubated with specific antibodies, at a dilution of 1:100 for anti-4.1NHP, 1:50 for anti-GH, 1:50 for anti-GnRH 39, 1:200 for anti-GHRH, overnight at 4 °C. After several washes in PBS, the sections were incubated with HRP-conjugated secondary antibody  for one hour at room temperature followed by exposure to the chromogen  system for 1–5 min. Slides were counterstained with haematoxylin and examined by light microscopy . A semiquantitative scoring system was used. The intensity of immunoreactivity was measured on a 0–3 scale. A score of 0 indicated no reactivity, a score of 3 denoted an intensity of cell reactivity with dark brown. The percentage positivity was based on as follows: A score of 0 indicated no cell immunopositivity, 1, 1–10% positive cells, 2, 11–40% positive cells, 3, 41–70% positive cells and 4, > 70% positive cells. The final scores for intensity and percentage of immunopositivity for each antibody were multiplied to give a number from 0 to 12.Immunohistochemical analysis was performed as described previouslyPituitary samples were fixed with 3% glutaraldehyde in 0.1 M sodium cacodylate buffer, pH 7.4, for 2 h at room temperature and rinsed in the same buffer several times. Following post-fixation in 1% osmium tetroxide solution, the samples were dehydrated with increasing concentrations of ethanol and small pieces of the tissue were embedded in EMBed-812 media  and cured at 60 °C for 24 h. Ultra-thin sections (65–70 nm) were cut with an MT-XL ultra-microtome , and stained with a Uranyl acetate solution followed by Reynold’s Lead Citrate Stain solution. Sections were imaged using a Tecnai G2 Spirit BioTWIN Transmission Electron Microscope . The area ratio of secretory granules in cytoplasm were calculated with QWin image analysis software (Germany)., tissue specimens, ~ 1 mm3 in size, were collected and fixed in the mixture of 4% paraformaldehyde and 0.1% glutaraldehyde in 0.1 mol/L cacodylate buffer, pH 7.4, for 1 h at 4 °C. After washing with 0.1 M cacodylate buffer and treatment with 50 mM ammonium chloride to quench residual aldehydes, the fixed tissue was dehydrated and embedded in LR-White Resin . Ultrathin sections of embedded tissue were cut with a diamond knife and mounted on to parlodion coated nickel grids. Non-specific binding was blocked by applying nonimmune serum to the sections, followed by incubation overnight at 4 °C with 4.1N primary antibody diluted 1:10 in 0.05 mol/L Tris-buffered saline (TBS), supplemented with 0.1 mol/L bovine serum albumin (BSA). After thorough rinsing with TBS containing 0.1% BSA and 0.5% Tween 20, the sections were incubated with 15-nm gold-conjugated goat-anti-rabbit IgG (Amersham Biosciences) diluted 1:20 in TBS for 2 h at room temperature. Following thorough rinsing with TBS, the sections were treated with 4% aqueous uranyl acetate to enhance contrast and imaged using the transmission electron microscope. For control, either the primary antibody was omitted or nonimmune serum was used.For immunogold labelingMice serum were collected and sent to Laboratory of Comparative Pathology, Memorial Sloan Kettering Cancer Center and Weill Medical College at Cornell University for hormone assay. FSH and LH levels were measured by the radio immunoassay and GH levels by ELISA enzyme immunoassay.All numerical data were presented as mean ± SD. Statistical analysis were performed by Student’s t-test and one-way ANOVA, values of P < 0.05 were considered to be significant.Supplementary Figures.