Pascopyrum smithii (Rydb.) Á. Löve], a temperate-region forage and turf grass, using multiple different combinations of N fertilizer [(NH4)2·CO3] and irrigation levels over 3 years to determine optimal field management. We conducted multifactorial, orthogonally designed field experiments with large sample sizes, and measured fertile tillers m-2 (Y1), spikelets/fertile tillers (Y2), florets/spikelet (Y3), seed numbers/spikelet (Y4), seed weight (Y5), and seed yield (Z) to study factors associated with the switch between vegetative and reproductive growth. Fertilization had a greater effect on seed yield and yield components than irrigation. Y1 had the strongest positive effect on Z, whereas Y5 had a negative effect on Z. Irrigation and fertilization affected Z, Y1, and Y5. Fertilizer concentrations were positively correlated with Z, Y1, and Y5, whereas irrigation levels were negatively correlated. The ridge regression linear model results suggested N application rate and irrigation had antagonistic effects on Y1 . We conclude that the optimal amount of N fertilizer and irrigation was 156 kg ha-1 + 115 mm for seed yield, 120 kg ha-1 + 146 mm for spikelets/fertile tillers, and 108 kg ha-1 + 119 mm for seed numbers/spikelets. These results will improve yield and reduce agricultural inputs for P. smithii in semiarid and arid regions, thereby reducing fertilizer pollution and conserving water.It is crucial for agricultural production to identify the trigger that switches plants from vegetative to reproductive growth. Agricultural sustainability in semiarid regions is challenged by nitrogen (N) fertilizer overuse, inadequate soil water, and heavy carbon emissions. Previous studies focused on the short-term effects of a single application of N and water but have not investigated the long-term effects of different irrigation and N fertilizer regimens on crop yields and yield components. N application is routinely coupled with water availability, and crop yields can be maximized by optimizing both. We examined the growth of western wheatgrass [ The leve varied  37, 38], 381 to ectively . Group C1 was measured from 10 randomly selected 1 m row samples; Y2, Y3, and Y4 were measured from 30 randomly selected fertile tillers and spikelets. The seed heads became ripe during the month of August from 2003 to 2005; we separately hand-threshed four samples from a 1-m length of the row, cleaned and weighed seeds from each sample, and determined that seed water content was 7–10% to convert weight values into kg ha-1 seed yield (Z)  among the five yield components. By contrast, Z was negatively affected by Y4 and Y5 during the experimental years (P≤0.05 or P≤0.01), and Z was not affected by Y3 during the experimental years. Y1 was significantly affected by irrigation and N application  was significantly affected by irrigation in all comparisons (P<0.006), and significantly affected by N during 2003–2005  in 2003 and 2004  in all three experimental years (Tables P<0.01) except for Y4 , and for Y2 and Y4 in two experimental years , which were detected using the PROC MIXED analysis of variance (ANOVA) in SAS (N application and irrigation rates strongly affected seed yield (Z) in the three consecutive growing seasons. The F test values of Z in 2003, 2004, and 2005 were much larger for the N rate  than for the irrigation rate , and the water × N interaction was significant for Z  in SAS .3) had a direct effect on all seed yield traits  > Y1 (0.590) > Y5 (–0.394) > Y2 (0.351) > Y4 (–0.260) > Y3 (0.091). The direct effect of X3 on Z was positive and strong  on Y1 was positive and strong in all three experimental years (P<0.0001), with coefficients of 0.144 in 2003, 0.109 in 2004, and 0.201 in 2005.N fertilizer (Xd traits . The tot2) had a weak direct effect on the yield factors Y2, Y3, and Y4. By contrast, irrigation had a strong, highly significant, and direct effect on Y1, Y5, and Z in at least two experimental years , with coefficients of –0.069 in 2003, –0.065 in 2004, and –0.096 in 2005. Therefore, irrigation had the largest effect on Y1. Irrigation significantly affected Y5, with coefficients of –0.089 in 2003, –0.039 in 2004, and 0.255 in 2005. Irrigation had a strong, direct effect on Y1, Y5, and Z, and the total effect in order of absolute value was Y1 (–0.296) > Y2 (–0.192) > Y5 (0.130) > Y3 (0.123) > Z (0.009) > Y4 (0.008)  .2 and X3. The strongest positive indirect effect on Z was from the X2 via X3 path (the coefficient was 0.047 in 2005), and the second strongest indirect effect was on Y5 from the X3 through X2 path (0.045 in 2005). We observed an effect on Z by X2 via X3 (0.025 in 2004) and on Y1 by X2 via X3 (0.024 in 2005). The strongest negative indirect effects were on Z, Y1, and Y5 were by X2 via X3 (–0.035 in 2005), and the second strongest indirect effects were on Y4 by X2 via X3 (–0.027 in 2005). X2 via X3 effects on Y3 were positive in 2003 but negative and marginal in the other two experimental years. X2 via X3 effects on Y1, Y2, and Z were positive in all experimental years. X2 via X3 effects on Y4 and Y5 were negative in all three experimental years  > Y1 (0.078) > Y5 (–0.057) > Y2 (0.042) > Y4 (0.040) > Y3 (0.010), which was the same as the effect of N fertilizer  > Y5 (0.031) >Y2 (–0.022) > Y3 (0.015) > Z (0.002) > Y4 (0.001), which was similar to the irrigation effect  as irrigation increased. Under these conditions, irrigation determined Y1  was beneficial for higher floret numbers  and X2 is irrigation (mm).With codes converted into applications of N and irrigation,2 was assigned new means for the total annual water requirement (precipitation and irrigation). The linear model results suggested that irrigation and N application rate had antagonistic effects on Y1. X2 effects were more dynamic than those of X3 , 108 kg/ha and 119 mm for seed numbers/spikelet (Y4), and 144 kg/ha and 115 mm for seed yield (Z), respectively  lines extended in different directions from those of florets/spikelet (Y3) in each year (5 lines in 2005 (green) displayed synergetic effects between irrigation and N supply (k>0), but the Y3 lines in 2005 (green) displayed antagonistic effects (k<0) between irrigation and N supply. All other ridgelines in each year display the same phenomenon   . The cooSeed yield is primarily influenced by climate conditions, environmental factors, and extreme weather events , especia-1 during the three years  .The 18 treatments of N fertilizer ranged from 0 to 480 kg/ha . The 11 levels of irrigation (X(DOCX)Click here for additional data file.


This study shows that a valuable anode material can be produced through conversion of CD by high-temperature carbonization. This approach provides a new way to alleviate environmental problems associated with CD.We develop a high-performance anode formed from carbonized cow dung for bioelectrochemical systems. Thermal gravimetric analysis showed that the CD carbonization process started at 300°C and ended at approximately 550°C; the weight was reduced by 51%. After a heat-treatment at 800°C for 2 h, the treated CD featured a good conductivity and a high specific surface area. The maximum current density of 11.74 ± 0.41 A m Currentor costs . TherefoLooking for cost effectiveness and performance affinity of electrode materials catalyzed the attempts to apply biochar in MFCs. Biochar is the product of heating biomass in the absence of oxygen and this carbonized material was used to be applied as soil amendment to fix nutrition or adsorbent to remove pollutants . RecentlToday there are an estimated 1.4 billion cows around the world, which produce approximately 4.2 million tons of cow dung (CD) per year . To addrIn this study, we show that valuable anode materials can be produced through conversion of CD by high-temperature carbonization. We also investigated the BES the current generation performances of these anodes.2 and thickness of 0.4 cm). The molded CD pieces were carbonized in a tube furnace for 2 h at different temperatures of 400, 600, 800, and 1000°C under a N2 flow rate of 300 mL min-1. After the carbonization was completed, a piece of titanium wire was bonded to the carbonized waste mass with conductive epoxy to ensure an electrical connection. The CD materials, subjected to the different carbonization temperatures of 400, 600, 800, and 1000°C, are denoted as CD400, CD600, CD800, and CD1000, respectively.Dry cow dung (CD)  was purchased as precursor to prepare the electrode material. The CD was mixed with certain proportion of water (g/g = 3/4), packed into a mold (2 cm × 1.3 cm × 0.5 cm), and then dried at room temperature  following the manufacturer’s instructions. Labeled cells were visualized and z-stacks were captured on a Zeiss LSM 780 confocal laser scanning microscope (CLSM). Microbial protein on the electrode was obtained using an ultrasonic cell disrupter  and high-speed freezing centrifuge . The amount of biological protein was determined by an ultraviolet spectrophotometer  according to the Coomassie brilliant blue methods.The surface morphology of the carbonized CD was characterized with a scanning electron microscope . X-ray photoelectron spectroscopy (XPS) was used to measure the elemental compositions. The high resolution O1s (528–546 eV) and C1s (281–300 eV) peaks were measured on an EscaLab 250Xi spectrometer with a monochromatic Al Kα source . Fourier Transform infrared spectroscopy  was used to detect the surface functional groups on the electrode samples. Thermogravimetric analysis (TGA) measurements were performed on a TA Q600 thermogravimetry/differential scanning calorimetry (TG/DSC) instrument to measure the weight loss over the temperature range 0–1200°C in a nitrogen atmosphere. Specific surface areas of the CD-based electrodes were measured through Quadrasorb SI . The biofilms developed on electrodes were subjected to the LIVE/DEAD BacLightAll the electrochemical tests were performed in a three-electrode system using a potentiostat . The CD-based electrodes were used as the working electrode, with a graphite plate electrode as the auxiliary electrode, and an Ag/AgCl reference electrode in 50 mL sterile M9 solution. Electrochemical impedance spectroscopy (EIS) was measured at open circuit potential with an amplitude of 10 mV over a frequency range of 100 kHz to 100 mHz. The Nyquist plots were fitted with a suitable equivalent electrical circuit .4Cl, 0.1 g L-1; NaCl, 0.5 g L-1; KH2PO4, 4.4 g L-1; K2HPO4, 3.4 g L-1; MgSO4, 0.1 g L-1, NaHCO3, 2 g L-1), trace elements , and 1 g L-1 of sodium acetate as an electron donor , followed by CD1000 and CD600, at 62.18 m2 g-1 and 54.30 m2 g-1, respectively. The specific surface area of CD400 was relatively low at 1.34 m2 g-1. All electrode surfaces after carbonization showed completely hydrophilic surfaces with contact angles of 0°  was porous and the pore size of carbons particles was approximately several tens of micrometers, which is likely beneficial for microbial adhesion. Among the samples that have been carbonized at different temperatures, the surface of CD800 appeared to be the roughest. The volume of CD1000 decreased slightly after the carbonization and the pore sizes in the carbon particles were smaller than those of CD800 Figure . Convers° Figure . TherefoAs shown by the XPS results Figure , all theThe high-resolution O1s spectra could also be divided into three component peaks, assigned to C = O species at 531.4 eV, O-C = O at 532.4 eV, and C-O-C 533.15 eV  when the temperature was increased to 600°C. The conductivity also improved with the increase of carbonization temperature. Notably, the conductivity remained stable when the carbonization temperature was greater than 800°C . EIS was measured and then fitted by the equation R1 + R2/Q2 + R3/Q3, where R1, R2 and R3 represent ohmic resistance, charge transfer resistance, and diffusion resistance of the anode, respectively   , and carup time) . Among oIn addition, a thick biofilm covered the whole surface of the CD electrodes  and could be seen in CLSM images Figure . However-2 (∼5 A m-2), which is half that of our CD anode system. Moreover, the current output performance of our CD anodes also considerably outperformed anodes based on other carbonized biomass, such as tubular bamboo (5.00 A m-2), loofah sponge (1.90 A m-2), textile (8 A m-2) (-2) and graphite (∼26.1 US m-2), the price of our CD electrodes was only ∼16.1 US m-2  . In addi8 A m-2) . The ove-2) was obtained for CD treated at 800°C. This study proposed carbonized cow dung could be used as a high-performance anode in BESs, which could convert CD waste into an economical and environmentally friendly electrode material.In this study, we have tested the current output performance of an anode formed from carbonized CD treated at different temperatures (400–1000°C) under a nitrogen atmosphere. The results of our TG/DSC analyses showed that the CD carbonization process started at 300°C and stopped at approximately 550°C. Furthermore, for carbonization at temperatures above 600°C the CD electrodes possessed high conductivity. The maximum current output performance (11.74 ± 0.41 A mHF, DS, and MW designed the study. HF and ZG wrote the manuscript. HF, ZG, and WC did much of the testing. JW, YJ, HQ, and XY helped with the data analysis. XZ and MW helped to improve the quality of the manuscript. All authors read and approved the final manuscript.JW was employed by the company Zhejiang Lantu Environmental Protection Co., Ltd., China. The remaining 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.


We find that treatment of transforming growth factor β (TGF-β)-differentiated monocytic (THP-1) cells with a PAK4-targeted inhibitor significantly reduces podosome formation and induces the formation of focal adhesions. This switch in adhesions confers a diminution of matrix degradation and reduced cell migration. Furthermore, reduced PAK4 expression causes a significant reduction in podosome number that cannot be rescued by kinase-dead PAK4, supporting a kinase-dependent role. Concomitant with PAK4 depletion, phosphorylation of Akt is perturbed, whereas a specific phospho-Akt signal is detected within the podosomes. Using superresolution analysis, we find that PAK4 specifically localizes in the podosome ring, nearer to the actin core than other ring proteins. We propose PAK4 kinase activity intersects with the Akt pathway at the podosome ring:core interface to drive regulation of macrophage podosome turnover.  •PAK4 kinase activity is crucial to macrophage podosome dynamics•Podosome loss due to PAK4 suppression reduces migration speed and degradation•PAK4 localizes to the podosome ring where it interacts with vinculin and paxillin The differential role of PAK family proteins in podosome function has not been previously evaluated. Foxall et al. demonstrate that PAK4 rather than PAK1 plays a pivotal role and use super resolution microscopy to position PAK4 at the podosome ring:core interface. Previously, PAK4 was found to partially colocalize with podosome F-actin cores in primary macrophages, and short hairpin RNA (shRNA)-mediated knockdown of PAK4 reduced podosome numbers . In thisde novo-binding interactions of PAK4Δkinase precipitated podosome formation in a background of endogenous PAK4 activity. Our data now clarify the importance of PAK4 kinase activity for podosome formation.Given that the function of PAK1 and PAK4 kinase activity in podosomes remains unclear  and thatMacrophages require mature, degradative podosomes to migrate efficiently . Using aIn addition to their roles in invadosome dynamics, both PAK1 and PAK4 have been implicated in the regulation of focal adhesions . Thus, wBecause PAK4i may also affect PAK5/PAK6 , which is able to reveal podosome organization at the nanoscale . EGFP-PAUsing the reconstructed localizations, we then calculated the distances of each protein from the podosome center in a given segment of the podosome ring, as described in It remains to be elucidated how PAK4 is being regulated within this process. PAK4 binds preferentially to Cdc42; however, binding to Cdc42 was suggested to be an intermediate step to PAK4 activation, acting to localize PAK4 activity to specific subcellular compartments . Active Our data point to the Akt pathway as a potential target for PAK4 activity within the podosome. However, there may be other alternative or additional targets. PAK4 can promote paxillin phosphorylation at focal adhesions . WhetherIn conclusion, we have shown that PAK4 kinase activity is essential in the regulation of macrophage podosomes. Inhibition of PAK4 results in a dramatic loss of podosomes and the formation of focal adhesions, both in differentiated THP-1 cells and dramatically in primary human macrophages. shRNA-mediated knockdown of PAK4 also reduces podosome number; this phenotype cannot be rescued by a kinase-dead variant of PAK4. Importantly, this is a report of serine/threonine kinase activity as a crucial component of podosome turnover. The switch from podosomes to focal adhesions confers a reduction in cell migration speed and matrix degradative ability, demonstrating a crucial role for PAK4 in promoting macrophage migration by the modulation of adhesion phenotype. Finally, we have shown that PAK4 localizes to the podosome ring by superresolution 3B analysis and that PAK4 is closer to the podosome core than vinculin, paxillin, and talin.claire.wells@kcl.ac.uk). All unique reagents generated in this study are available from the Lead Contact with a completed Materials Transfer Agreement.Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Dr Claire Wells  were stored in 90% FBS and 10% DMSO in liquid nitrogen. Podosome formation was induced by plating cells on surfaces coated with 10μg/ml fibronectin (Sigma) in media containing 2ng/ml recombinant human TGF-β1 (R&D Systems), and incubating for 16 hours.THP-1 cells purchased from ATCC were cultured in suspension in RPMI-1640 medium (GIBCO) containing 10% heat-inactivated fetal bovine serum , 50μM β-mercaptoethanol (Sigma) and 1% penicillin/ streptomycin . Cells were incubated at 37°C with 5% CO2.Adherent HEK293T cells (ATCC) were cultured in RPMI-1640 medium (GIBCO) containing 10% heat-inactivated FBS, 1% penicillin/streptomycin and 2mM Glutamine (Sigma). HEK293T cells were maintained at between 50%–100% confluency by subculturing using 2ml trypsin EDTA  in PBS. Cells were maintained at 37°C with 5% CO6 cells/ml on fibronectin coated coverslips (coated following the same protocol as for THP-1 cell differentiation). Cells were cultured in RPMI-1640 medium containing 10% FBS, 1% penicillin/streptomycin and 2mM Glutamine, in the presence of 50ng/ml M-CSF (Miltenyi Biotec) for 4.5 days.Primary human monocytes were isolated from anonymous healthy donors peripheral blood samples purchased from the London Blood Transfusion service. Density gradient separation by centrifugation was carried out using 15ml Lymphoprep . Peripheral blood was diluted with PBS at a ratio of 1:2, and the cell suspension added to the density gradient before centrifuging at 800 x g at room temperature for 30 minutes. Peripheral blood mononuclear cell (PBMC) fraction was harvested and washed twice with PBS, centrifuging at 200 x g for 10 minutes at room temperature. Cells were resuspended in MACS buffer  and 0.5M EDTA), then CD14+ monocytes were isolated using magnetic bead separation using CD14 MicroBeads (Miltenyi Biotec) following the manufacturer’s protocol. To differentiate monocytes to macrophages, monocytes were seeded at a density of 0.3x10E. coli (Invitrogen).cDNA encoding wild-type human PAK4 was amplified by PCR from pDEST27-PAK4 template plasmid and subcloned into the pCR-BLUNT vector  while incorporating C- and N-terminal restriction sites for subsequent cloning into the pLNT/SffV lentiviral transfer vector. To generate shRNA-resistant and kinase-dead mutants, the QuikChange® XL Site-Directed Mutagenesis Kit (Stratagene) was used following the manufacturer’s instructions using the intermediate vector of PAK4 in pCR-BLUNT. Primers of approximately 20 nucleotides were designed to introduce the desired mutations. Cloned vectors were amplified using TOP10 chemically competent PAK4 shRNA sequences were cloned into the lentiviral transfer vector pLKO.1 (Addgene) following the manufacturer’s protocol. Three shRNA sequences were chosen and are listed in the 5 cells/ml in 12-well plates in 1ml growth medium, and incubated at 37°C with 5% CO2 overnight. The following day, HEK293T cells were transfected with viral plasmids. A 500μl transfection mixture was made containing 1.3μg p8.91 packaging plasmid, 0.42μg pMD.G envelope plasmid and 1.74μg pLNT/SffV or pLKO.1 transfer plasmid and 4.35μM polyethylenimine  in OptiMEM (Invitrogen). This mixture was incubated at room temperature for 15 minutes, then HEK293T cells were washed gently with OptiMEM before the transfection mix was added. Cells were then incubated at 37°C with 5% CO2 for 4 hours, before removing the transfection mix and adding 1ml growth medium. Transfected HEK293T cells were incubated at 37°C with 5% CO2 for 48 hours, before harvesting the virus by collecting the growth medium and centrifuging for 5 minutes at 2000 x g, then filtering through a 0.45μm syringe filter (Thermo Fisher Scientific).HEK293T cells were seeded at a density of 3-6x105 THP-1 cells in 600μl growth media in each well of a 12-well plate and adding 400μl filtered lentivirus solution, with 4μg/ml polybrene (Sigma) to increase infection efficiency. Cells were incubated at 37°C with 5% CO2 for 72 hours before washing twice by centrifuging at 1200rpm for 5 minutes, removing media and adding 5ml PBS before centrifuging again at 1200rpm for 5 minutes. Cells were then resuspended in 3-5ml growth medium and cultured at 37°C with 5% CO2. For cells transduced with pLKO.1 encoding PAK4 shRNAs, cells were selected at this stage by adding 500nM puromycin (Sigma) to growth medium.Viral transduction of THP-1 cells was carried out by seeding 1x102, before being fixed in 3.7% paraformaldehyde  in PBS for 30 minutes. See Table 1 below. For inhibitor wash-out experiments, following 4 hours incubation with inhibitors, cells were washed 3 times with fresh media and then incubated for 1-4 hours in media containing 2ng/ml TGF-β, before being fixed in 3.7% PFA in PBS. Primary human macrophages differentiated for 4.5 days with M-CSF were treated with 1μM or 5μM small molecule PAK inhibitors diluted in DMSO for 4 hours while incubating at 37°C with 5% CO2.THP-1 cells were differentiated toward a macrophage-like phenotype by seeding on fibronectin-coated coverslips in the presence of TGF-β for 16 hours. Cells were then treated with 1μM or 5μM small molecule PAK inhibitors (PAK4i from Cancer Research UK and CRUK Therapeutic Discovery Laboratories) or IPA-3 from Santa Cruz Biotechnology) or 1μM, 5μM or 10μM of Akt inhibitor , diluted in DMSO (Sigma) and added to culture media for 4 hours while incubating at 37°C with 5% COTGF-β differentiated THP-1 cells or M-CSF differentiated primary macrophages seeded on fibronectin coated coverslips were fixed and stained for vinculin and F-actin, and visualized using 100x objectives on LSM510 or Nikon confocal microscopes. For each coverslip, 5 distinct regions were visualized (top/bottom/left/right/center), and the number of podosomes in 20 cells per region was counted, to give a total of 100 cells per coverslip. From these counts, percentage of cells with podosomes was calculated, as well as the number of cells with 0, 1-10, 11-20, 21-30 or >30 podosomes was calculated. For each treatment condition, at least 3 coverslips were counted (> 300 cells per treatment condition) from >3 separate experiments.4 THP-1 cells per treatment condition were taken from suspension culture, centrifuged, and resuspended in 1ml fresh growth medium. Cells were pre-treated with 1μM or 5μM of the PAK inhibitors PAK4 or IPA3 diluted in DMSO, or DMSO as a control, for 1 hour while incubating at 37°C with 5% CO2. Cells were then taken up into 15ml falcons with 5ml PBS and centrifuged at 1200rpm for 3 minutes. Cells were resuspended in 800μl growth media containing 2ng/ml TGF-β and the same concentration of PAK inhibitor used for pre-treatment. For each treatment condition, 4 wells of a 96-well plate were coated with fibronectin as previously described, and these wells were washed twice with PBS before seeding 200μl cells per well. An additional 4 wells were coated with fibronectin and incubated with media containing TGF-β to give blank measurements. Cells were then incubated at 37°C with 5% CO2 for 16 hours, before dumping the plate roughly onto paper towel to remove media and unattached cells. 0.5mg/ml MTT reagent (Sigma) in PBS was sterile filtered before adding 100μl to each well and incubating at 37°C with 5% CO2 for 4 hours. MTT reagent was removed by needle aspiration and 100μl DMSO added to each well, pipetting up and down to mix, before incubating for a further 10 minutes at 37°C. Absorbance at 540nm was measured per well using a FLUOstar® Omega microplate reader (BMG Labtech) and blank-corrected values used. Readings from 4 wells per treatment condition x 3 separate experiments were used to give mean absorbance levels.6x105 cells/ml with 2ng/ml TGF-β for 16 hours. Cells were then fixed and stained.Fibronectin-488  coating of coverslips was carried out following the same protocol as used with unlabeled fibronectin: coverslips were inverted onto 200μl of 10μg/ml fibronectin-488 diluted in PBS, and incubated at 37°C for >3 hours. Coverslips were washed twice with PBS before seeding THP-1 cells at a density of 2x102) or degraded area (μm2)/cell was calculated.Images were taken using a 100x objective on a Nikon confocal microscope and analyzed using ImageJ. Fibronectin-488 images were converted to binary images with fibronectin-488 appearing white with black degradation spots. A threshold was applied  and the ImageJ Analyze Particles tool used to measure the total degradation area for each image. The same process was carried out using the F-actin channel to give a measure of total cell area in each image. The degradation area was divided by total cell area. For each treatment condition, >20 images were measured, and the mean degraded area/cell area . Just before filming, 25mM HEPES  was added to each well. Cells were imaged at 37°C in phase contrast using a 10x objective on an Olympus IX-71 microscope, taking images of 2 regions per well every 2.5 minutes for 2 hours.Cells were tracked using the ImageJ Manual Tracking tool . All cells in the starting frame were tracked, except cells that migrated outside the field of view during the duration of the film. Tracks were saved and analyzed using Mathematica notebooks developed by Professor Graham Dunn, to calculate mean cell speed for each treatment condition. To plot cell tracks, DiPer™ was used following the developers’ instructions . ApproxiAdherent THP-1 cell lysates were made by seeding THP-1 cells on fibronectin-coated 10cm plates with 2ng/ml TGF-β for 16 hours, before washing twice with ice cold PBS and adding 1ml lysis buffer , then incubating on ice for 10 minutes. Cells were then scraped into 1.5ml Eppendorf tubes, before incubating on ice for a further 10 minutes. For non-adherent cells, THP-1 cells were counted and centrifuged at 1200rpm for 3 minutes, culture medium removed and pelleted cells washed twice with ice cold PBS before resuspending in 1ml lysis buffer and incubating on ice for 20 minutes while vortexing every 5 minutes. Lysates were then spun down at 4°C for 10 minutes at 13,000 rpm, and the supernatant was transferred to new tubes. Before running on SDS polyacrylamide gels, 5x SDS sample buffer was added to lysates before heating at 95°C for 5 minutes. SDS-PAGE was carried out using 7.5%–12% gels and transferring onto nitrocellulose membranes. To immunoblot for protein levels, membranes were blocked with 5% milk in TTBS before probing with primary antibodies followed by secondary antibodies listed in the 7 TGF-β differentiated THP-1 cells were lysed in 1ml lysis buffer. 30μl lysate with 20μl 5x SDS sample buffer was kept at −20°C as input. The remaining lysate was incubated with 1-2μg antibody for immunoprecipitation overnight at 4°C while rotating. 60μl protein A/G agarose beads were washed 3 x in lysis buffer, and lysate/antibody samples added to the beads, before rotating at 4°C for 1 hour. Samples were centrifuged at 4000rpm for 3 minutes at 4°C and the supernatant removed, before washing beads 3x by incubating with 1ml lysis buffer for 5 minutes while rotating at 4°C. Lysis buffer was removed before adding 30μl 5x SDS sample buffer and boiling for 10 minutes to precipitate proteins from the beads. Immunoprecipitates were loaded on polyacrylamide gels and western blots carried out.For immunoprecipitation, 6x1020 before mounting on glass slides on 15μl mounting media  and incubating at 37°C for 20 minutes before storing at room temperature in the dark. For all antibodies used, see the Cells were stained for immunofluorescence imaging at room temperature as follows: cells were fixed with 3.7% PFA for 30 minutes, then washed 3x with PBS before permeabilization with 0.1% Triton X-100 (Sigma) in PBS for 5 minutes. Cells were again washed 3x with PBS, then blocked with 3% BSA in PBS for 30 minutes, before incubating with primary antibody diluted in 3% BSA in PBS for 1 hour. Cells were then washed 4x for 5 minutes per wash with PBS, before incubating with secondary antibody diluted in 3% BSA in PBS for 30 minutes. Cells were again washed 4x for 5 minutes per wash with PBS, before mounting coverslips. Coverslips were washed with ddHImages of fixed cells were acquired using 100x oil Plan Fluor immersion objectives  on Zeiss LSM510, Nikon TIRF or Nikon Spectral confocal microscopes. Excitation wavelengths of 488nm, 543nm and 633nm were used. Images were converted to PNG files from database files.mdb or ND2 files using ImageJ, and merged or single channel images generated using ImageJ. For live cell imaging, a 10x objective on an Olympus IX-71 wide-field inverted microscope equipped with a 37°C environment chamber was used. Image acquisition was controlled by Metamorph imaging software , and images were taken in phase contrast every 2.5 minutes for 2 hours. Combined STORM and 3B imaging was carried out with Nikon N-STORM microscope. For STORM imaging, 10,000 frames per sample were processed with ThunderSTORM software  using deFocal adhesions were quantified using ImageJ as follows: vinculin images were converted to binary images, a threshold applied and the ImageJ Analyze Particles tool used to measure the mean number of adhesions per cell. Statistical analyses were performed using SPSS Statistics software (IBM). Checks for normality were performed before the appropriate statistical test for the data was chosen. Student’s t test or one-way analysis of variance (ANOVA) was used; for each set of data, the test used and n values are indicated in the figure legend. Results were defined as significantly different with a p value of < 0.05.This study did not generate any unique datasets or code. All data are contained within the paper.


Mouse models exhibit increased Ca2+ buffering and arrhythmias, and we hypothesized that these changes are primary effects of the mutations (independent of compensatory changes) and that increased Ca2+ buffering and altered Ca2+ handling contribute to HCM pathogenesis via activation of Ca2+-dependent signaling. Here, we determined the primary effects of HCM mutations on intracellular Ca2+ handling and Ca2+-dependent signaling in a model system possessing Ca2+-handling mechanisms and contractile protein isoforms closely mirroring the human environment in the absence of potentially confounding remodeling. Using adenovirus, we expressed HCM-causing variants of human troponin-T, troponin-I, and α-tropomyosin  in isolated guinea pig left ventricular cardiomyocytes. After 48 h, each variant had localized to the I-band and comprised ∼50% of the total protein. HCM mutations significantly lowered the dK of Ca2+ binding, resulting in higher Ca2+ buffering of mutant cardiomyocytes. We observed increased diastolic [Ca2+] and slowed Ca2+ reuptake, coupled with a significant decrease in basal sarcomere length and slowed relaxation. HCM mutant cells had higher sodium/calcium exchanger activity, sarcoplasmic reticulum Ca2+ load, and sarcoplasmic/endoplasmic reticulum calcium ATPase 2 (SERCA2) activity driven by Ca2+/calmodulin-dependent protein kinase II (CaMKII) phosphorylation of phospholamban. The ryanodine receptor (RyR) leak/load relationship was also increased, driven by CaMKII-mediated RyR phosphorylation. Altered Ca2+ homeostasis also increased signaling via both calcineurin/NFAT and extracellular signal–regulated kinase pathways. Altered myofilament Ca2+ buffering is the primary initiator of signaling cascades, indicating that directly targeting myofilament Ca2+ sensitivity provides an attractive therapeutic approach in HCM.Mutations in thin filament regulatory proteins that cause hypertrophic cardiomyopathy (HCM) increase myofilament Ca For example, reduction of SERCA levels e fibers –7. Furth of cTnC , 9. Othe in MYH7  and MYBP (MYBPC3 ) are als systole . We preduffering , 14, wheuffering , 16. In A levels  and myofA levels , both weinea pig . The gen is ∼30% . Cardiac is ∼30% , indicat2+ handling in HCM. Furthermore, we present for the first time a novel analytical paradigm to fully incorporate the consequences of Ca2+ buffering on cardiomyocyte Ca2+ transients and Ca2+ handling protein activities. We consider the dynamics of total intracellular Ca2+  derived from NCX integral measurements rather than free Ca2+ ([Ca2+]i) from fura2 fluorescence when assessing SR load, fractional SR release, and SERCA activity. When applied, our data show that the primary effects of HCM mutant gene expression include altered Ca2+ transients and increased myofilament Ca2+ buffering, increased [Ca2+] in both the SR and cytoplasm, and activation of Ca2+-dependent signaling mediated by Ca2+/calmodulin-dependent protein kinase II (CaMKII), calcineurin/nuclear factor of activated T-cells (NFAT), and mitogen-activated protein kinase/ERK.We have engineered adenoviruses to express WT and R92Q cTnT, WT, and R145G cTnI, and WT and D175N α-TM. By measuring the impact of mutations affecting three different sarcomeric proteins, we aimed to identify hallmark changes in CaA and C). The relative expression of thin filament regulatory protein subunits was determined by Western blotting and subsequently adjusted by the infection levels for each cell preparation tested. Conjugation of an N-terminal (cTnT and cTnI) or C-terminal (α-TM) FLAG tag increased the molecular weight of the recombinant proteins sufficiently to allow differentiation from the endogenous subunit (E). The relative expression level of recombinant protein in purified cardiomyocytes coexpressing human recombinant GFP was found to be 54.3 ± 9.5% (cTnT R92Q), 49.3 ± 8.5% (cTnI R145G), and 57.6 ± 7.5% (TM D175N) (B and D), thus likely to closely reflect the levels of mutant protein found in patients with autosomal dominant cardiomyopathy. The recombinant FLAG-tagged thin filament proteins were localized to the I-band of the myofilaments of infected cardiomyocytes . To assess the impact of culture time on cardiomyocyte function, we compared the contractility and Ca2+ transients of cardiomyocytes cultured for 1, 24, and 48 h along with the extent of t-tubular dedifferentiation . We observed a small but significant reduction in the number of t-tubules and a prolongation in the relaxation and Ca2+ reuptake after 48 h .Isolated guinea pig left ventricular cardiomyocytes were infected with ∼1000 MOI of recombinant adenovirus. The level of infection was between 32.1 ± 7.2 (WT TnI) and 87.6 ± 9.1% (WT TM), calculated from the coexpression of human recombinant GFP , A and C subunit E. The reM D175N) , B and D2+]total and [Ca2+]i were measured upon the application of 10 mm caffeine using simultaneous measurement of fura2 fluorescence ratio and whole-cell voltage-clamp current in a method adapted from Trafford et al. . We postulated that increasing the Ca2+ affinity of myofilaments by introducing a Ca2+ sensitizing HCM mutation will in turn increase total Ca2+ buffering in the intact cardiomyocytes. We fitted the calibrated total versus free [Ca2+]i data to a Michaelis–Menten type equation (A). The dK of Ca2+ buffering was significantly decreased in the presence of each HCM-causing mutation . However, the total Ca2+ occupancy of the myofilaments (Bmax) was not significantly altered between groups (Table S2). The relative buffering at low [Ca2+] is described by Bmax/dK, and this ratio was increased for all three mutants compared with WT . The calculated relationship between [Ca2+]total and [Ca2+]i over the full range of [Ca2+] is shown in Fig. 2D. We have used these fits to convert observed [Ca2+]free to [Ca2+]total for each cell and calculate changes in SR load and SERCA activity. This incorporates the effect of altered Ca2+ buffering when considering these key Ca2+-handling parameters.i (Δ[Ca2+]i = 0.175 ± 0.007 to 0.307 ± 0.011 μm)  and significantly prolonged rates of Ca2+ reuptake (Δτ = 0.027 ± 0.003 to 0.041 ± 0.002 s−1)  compared with cardiomyocytes expressing the corresponding WT protein. The time to 50% Ca2+ release from the SR and the Ca2+ transient amplitude were not significantly different between groups. In measurements of sarcomere shortening at 1-Hz pacing, all three HCM mutations were found to decrease basal sarcomere length compared with WT (ΔSL = −0.030 ± 0.007 to −0.094 ± 0.007 μm) in line with the higher levels of diastolic [Ca2+]i. Cardiomyocytes expressing HCM mutations also had significantly prolonged relaxation (ΔT50 = 0.011 ± 0.002 to 0.037 ± 0.003 s) . Cardiomyocytes expressing the R145G mutation in cTnI displayed reduced fractional shortening compared with WT; these cells also had the shortest diastolic sarcomere length (1.776 ± 0.007 versus 1.865 ± 0.009 μm in WT), and this may impose some limit on the extent of shortening compared with the other two mutations, because of fura2 accumulation .In analyses of cytosolic i transients, SR load, fractional SR Ca2+ release, and SERCA activity were calculated and found to be decreased, increased, and unchanged, respectively . Furthermore, we found that these alterations were independent of pause duration prior to caffeine spritz (Table S6). However, we also calculated caffeine and Ca2+ transients expressed as [Ca2+]total derived from the NCX current integral, thus incorporating any alterations to Ca2+ buffering, and used this approach to recalculate SR load, fractional SR Ca2+ release and SERCA activity . HCM mutant cells consistently displayed a higher SR load  . The preceding [Ca2+]i transient amplitude (ATable S5) was also converted to [Ca2+]total (BTable S5) to calculate the fractional SR Ca2+ release and SERCA2 activity. HCM cells had unchanged SR fractional release compared with WT  and had increased SERCA2 activity (Δrate = 1.53 ± 0.30 to 3.87 ± 0.42 s−1) .Using a standard method  that utiE), whereas the PKA site at serine 16 of PLN and absolute levels of NCX and SERCA2 remained unaltered . This result also confirms the need to consider the [Ca2+]total transient amplitudes when investigating SERCA2 activity and SR load in cardiomyocytes with altered Ca2+ buffering.The increases in SR load and SERCA2 activity appear to be driven by increased CaMKII-dependent phosphorylation of phospholamban (PLN). Relative phosphothreonine 17 PLN levels were increased in HCM cells (between 87 ± 27 and 220 ± 72% more than WT) E, wherea2+]i in HCM cardiomyocytes suggest there may be Ca2+ leak from the RyR. To test this, we used a method adapted from Shannon et al. (A) and found that RyR leak is profoundly increased in cells expressing HCM mutations (B). It was also observed that RyR blockade with tetracaine increased the [Ca2+]i caffeine transient amplitude by 12–25% in HCM cells (C) but had no effect on WT-infected or uninfected control myocytes. Despite the combination of increased leak (B) and [Ca2+]i SR load  (C), the relative leak load relationship was still higher in HCM cardiomyocytes (Δμm/s/μm = 5.24 × 10−5 ± 1.65 × 10−5 to 2.22 × 10−4 ± 3.75 × 10−5) (D), suggesting that other factors may be driving RyR channel open probability. We measured the phosphorylation status of each of the known regulatory sites on the RyR  and found that (consistent with changes in PLN phosphorylation), there was higher phosphorylation at the RyR CaMKII site serine 2814 (20 ± 11 to 87 ± 27% greater than WT) (E); phosphorylation at the PKA/PKG sites (serine 2030 and 2808) did not differ between WT and HCM mutant cardiomyocytes .Increased SR load and diastolic i and [Ca2+]total to be calculated. To exclude the other flux mechanism's contribution to the extrusion of Ca, the measured NCX currents were integrated and corrected as previously described  the change of integral of NCX current. This gave the Δ[Ca2+]total, whereas the change of intracellular Ca2+ transient measured by fura2 gave Δ[Ca2+]i during caffeine application. Thus, the relationship between Δ[Ca2+]i and Δ[Ca2+]total represents the Ca2+ buffering regardless of differential bound myofilament Ca2+ in control and mutant cells at baseline. Plots of [Ca2+]i versus [Ca2+]total were fitted to a Michaelis–Menten equation to give estimates of buffering dK and Bmax. NCX amplitudes were taken from currents obtained during the buffering measurement protocol. Relative changes in SR load were calculated using [Ca2+]total transients upon caffeine spritz to account for the Ca2+ buffering of each cell. SERCA2 activity was determined from the decay constants from the caffeine transient and the preceding Ca2+ transient using standard methods . Sarcomere shortening was captured by Fourier transform of the cardiomyocyte striations under phase contrast microscopy using a switching rate of 100 Hz. Ca2+ transients were captured simultaneously, using the ratio of fura2 fluorescence emission at 365/380 nm at a switching rate of 1000 Hz. All contracting cardiomyocytes were measured for contractility and fura2 Ca2+; any cells displaying asynchronous contractility or excessive blebbing/dysmorphology were ignored for acquisition. fura2 fluorescence ratio was subsequently calibrated to [Ca2+]i by selective membrane permeabilization using ionomycin .fura2-loaded guinea pig left ventricular cardiomyocytes were then allowed to settle to the bottom of a perfusion chamber with a 0 thickness coverslip base, which was mounted on an inverted fluorescence microscope. The cells were perfused with buffer containing 150 mm CaCl2. RyR leak was measured for 50 s in the absence of field stimulation in a Na+- and Ca2+-free solution. Field stimulation and perfusion of 1.8 mm CaCl2 were restarted to allow verification that basal functional conditions were as at the start of the experiment. Perfusion was again switched to Na+- and Ca2+-free solution containing 1 μm of tetracaine, and field stimulation was stopped to make a baseline reading that was subtracted from the preceding RyR leak measurement. Leak/load relationships were measured by 10 mm caffeine spritz following each perfusion switch.RyR leak was calculated using the RyR channel blocker tetracaine as previously reported . Brieflyversus α-actinin. NFAT and ERK nuclear localization was also measured versus nucleoplasmic marker 4′,6′-diamino-2-phenylindole.Western blots were performed to detect cTnT, cTnI α-TM and FLAG tag, PLN, phosphoserine 16 PLN, phosphothreonine 17 PLN, RyR, phosphoserine 2030 RyR, phosphoserine 2808 RyR, phosphoserine 2814 RyR, phosphoserine 22/24 cTnI, NCX, SERCA2A, NFAT, phosphoserine 165 NFAT, ERK, or phosphothreonine 202/tyrosine 204 ERK in WT and mutant-infected cardiomyocytes. In immunolocalization experiments, recombinant FLAG-tagged protein was detected n experiments ± S.E. throughout. Statistically significance was determined using unpaired Student–Newman–Keuls analysis for nonnormally distributed data , with significance values defined as p < 0.05.The data are expressed as the average of P. R., Y.-H. Z., B. C., H. W., and C. R. conceptualization; P. R., X. L., A. S., S. P., and C. R. data curation; P. R., X. L., and S. P. formal analysis; P. R., B. C., H. W., and C. R. supervision; P. R., X. L., A. S., S. P., and Y.-H. Z. investigation; P. R. and X. L. methodology; P. R. and C. R. writing-original draft; P. R. project administration; P. R., X. L., Y.-H. Z., B. C., H. W., and C. R. writing-review and editing.


Human carboxylesterases (hCES) are key serine hydrolases responsible for the hydrolysis of a wide range of endogenous and xenobiotic esters. Although it has been reported that some ginsenosides can modulate the activities of various enzymes, the inhibitory effects of ginsenosides on hCES have not been well-investigated.In this study, more than 20 ginsenosides were collected and their inhibitory effects on hCES1A and hCES2A were assayed using the highly specific fluorescent probe substrates for each isoenzyme. Molecular docking simulations were also performed to investigate the interactions between ginsenosides and hCES.O-β-(d-glucosyl)-dammarenediol II (DMG) displayed potent inhibition against both hCES1A and hCES2A, while protopanaxadiol (PPD) and protopanaxatriol (PPT) exhibited strong inhibition on hCES2A and high selectivity over hCES1A. Introduction of O-glycosyl groups at the core skeleton decreased hCES inhibition activity, while the hydroxyl groups at different sites might also effect hCES inhibition. Inhibition kinetic analyses demonstrated that DM and DMG functioned as competitive inhibitors against hCES1A-mediated d-luciferin methyl ester (DME) hydrolysis. In contrast, DM, DMG, PPD and PPT inhibit hCES2A-mediated fluorescein diacetate (FD) hydrolysis via a mixed manner.Among all tested ginsenosides, Dammarenediol II (DM) and 20S-The structure–inhibition relationships of ginsenosides as hCES inhibitors was investigated for the first time. Our results revealed that DM and DMG were potent inhibitors against both hCES1A and hCES2A, while PPD and PPT were selective and strong inhibitors against hCES2A. Panax ginseng Meyer, one of the most popular edible herbs used in both eastern and western countries, has been found with many beneficial effects for human health. Modern pharmacological and pharmacodynamic researches have demonstrated that both ginseng extract and its major constituents (ginsenosides) can enhance memory, improve immunity, improve cardiovascular functions, delay aging and anti-tumor, etc. [Panax ginseng, and most of them are 20(S)-protopanaxadiol (PPD)-type and 20(S)-protopanaxatriol (PPT)-type ginsenosides [Ginseng, the root or rhizome of or, etc. –6. Over or, etc. –10. Ginsor, etc. , anti-tuor, etc. –14 and aor, etc. , 16. Untenosides  In many Asia countries, ginseng has been widely used as a key material for preparing dietary supplements, herbal medicines and cosmetics , 19. CurMammalian carboxylesterases (CES) are key hydrolases in most organs, which catalyze the hydrolysis of various esters (including endogenous and xenobiotic ones) into the corresponding alcohols and carboxylic acids –24. In tOver the past decade, ginseng-related products have been widely used in combination with a variety of therapeutic drugs to treat various types of cancer in clinical settings. In addition to enhancing efficacy and reducing drug toxicity, the accumulating evidence has indicated that ginseng products may bring benefits to the cancer patients, including increasing immunity and tolerance of cancer patient , 39. NotN--2-chloroacetamide (NCEN) and d-luciferin methyl ester (DME) was synthesized by us and the synthetic schemes have been reported previously [Twenty-five ginsenosides were purchased from Chengdu Pufei Biotech Co., Ltd.  and used in this study. Luciferin Detection Reagent (LDR) was obtained from Promega Biotech . Fluorescein diacetate (FD) was from TCI , eviously , 42. FD d-luciferin (the hydrolytic metabolite of DME) was measured (excitation wavelength 580 nm) by a multi-mode microplate reader . The calculation of residual activities of different inhibitors against hCES1A was performed as described previously [DME hydrolysis was used as the probe reaction to assay the inhibitory effects of 25 ginsenosides against hCES1A, oleanic acid was used as a positive control of inhibition assays . The inceviously .® iD3 multi-mode microplate reader. The kinetic parameters were set at 30 reads, with an interval of 30 s (30 min) and shake during 5 s before the first read at 37 °C. The calculation method for residual activities of different inhibitors against hCES2A have been reported previously [To determine the inhibitory effects of ginsenosides against hCES2A, FD was used as the probe substrate, while loperamide (LPA) was used as a positive control of hCES2A inhibition assays. The incubation system contains 2 μL HLM , 194 μL PBS (0.1 M pH 7.4) and 2 μL corresponding inhibitor. After pre-incubating the incubation mixture for 3 min at 37 °C, the reaction was started by adding FD  with a total volume of 200 μL, in which the final content of DMSO was limited to 2% (v/v). The hydrolytic metabolite of FD was quantified  by a SpectraMaxKi) and inhibition kinetic types of ginsenosides were determined. The experimental details used to determine the inhibition constants have been reported previously [According to the inhibition kinetics of both hCES1A and hCES2A, which were investigated by using varying concentrations of probe substrates in the presence of different concentrations of ginseng compounds (inhibitors), the inhibition constant  on hCES2A were investigated in living cells. The detailed processes for hCES2A inhibition assays in HepG2 cells have been depicted previously .Docking simulations were executed to simulate the interactions between ginseng constituents and hCES by using Discovery Studio . The detailed processes for docking simulation were depicted previously , 43.50 and Ki values with GraphPad Prism 7.0 software .All inhibition assays were performed in triplicate, nonlinear regression was used to determine the ICO-glycosyl groups on the core skeleton , were hardly inhibit the hCES1A-mediated DME hydrolysis and hCES2A-mediated FD hydrolysis in HLM even at high dosage (100 μM). By contrast, the ginsenosides without O-glycosyl groups (such as Dammarenediol II (DM), panaxadiol, panaxatriol, PPD and PPT), exhibited strong or moderate inhibition on both hCES1A and hCES2A. Of note, 20S-O-β-(d-glucosyl)-dammarenediol II (DMG), a ginsenoside with a O-glycosyl group at the Rʹ site, also displayed strong inhibition on both hCES1A and hCES2A.First of all, the inhibitory potentials of 25 ginsenosides against hCES1A and hCES2A were preliminarily screened. As shown in Fig. 50 values of each ginsenoside against hCES1A and hCES2A were depicted using varying inhibitor concentrations  against hCES1A in HLM was also determined as 0.10 μM  against hCES2A in HLM was determined as 1.46 μM  on the skeleton  will lead to the loss of hCES1A inhibition, with the exception of DMG (IC50 = 1.76 μM).For the ginsenosides with skeleton A or skeleton C, introduction of any O-glycosyl group or introduction of a O-glycosyl group at the R2 site will abolish the hCES2A inhibition, such as ginsenoside C-K (IC50 = 15.46 μM) VS ginsenoside Re (IC50 > 100 μM), ginsenoside Rg1 (IC50 > 100 μM) and ginsenoside R1 (IC50 > 100 μM), PPD (IC50 = 1.34 μM) VS ginsenoside Rg2 (IC50 > 100 μM) and ginsenoside Rh1 (IC50 > 100 μM).For the ginsenosides with skeleton A, replacing the hydroxyl groupby a 3 site of the ginsenosides with skeleton A significantly reduces the inhibitory effects on hCES1A, such as DM (IC50 = 1.99 μM) VS PPD (IC50 > 100 μM), and DMG (IC50 = 1.76 μM) VS ginsenoside C-K (IC50 > 100 μM).Addition of a hydroxyl group at the R2 site reduces inhibitory effects on both hCES1A and hCES2A (about 3.8–5.0-folds), such as panaxadiol (IC50 = 6.95 μM) VS panaxatriol (IC50 = 34.93 μM) for hCES1A, and panaxadiol (IC50 = 3.78 μM) VS panaxatriol (IC50 = 14.59 μM) for hCES2A.For the ginsenosides with skeleton B, introduction of a hydroxyl group at the RO-glycosyl group does not affect hCES1A inhibition but reduces hCES2A inhibition, such as DM (IC50 = 0.69 μM) VS DMG (IC50 = 1.06 μM), PPT (IC50 = 0.86 μM) VS ginsenoside F1 (IC50 = 37.08 μM), Rh2 (IC50 = 5.35 μM) VS F2 (IC50 = 14.37 μM), as well as PPD (IC50 = 1.34 μM) VS C-K (IC50 = 15.46 μM).For the ginsenosides with skeleton A, replacing the hydroxyl group at the Rʹ site by a 1 site is essential for hCES2A inhibition, the presence of a monosaccharide or polysaccharide group at this site will decrease hCES2A inhibition activity, such as PPD (IC50 = 1.34 μM) VS Rg3 (IC50 = 10.37 μM), PPD (IC50 = 1.34 μM) VS Rh2 (IC50 = 5.35 μM).For the ginsenosides with skeleton A and skeleton B, the hydroxyl group at the R3 site will decrease hCES2A inhibition, such as DM (IC50 = 0.69 μM) VS PPD (IC50 = 1.34 μM), and DMG (IC50 = 1.06 μM) VS C-K (IC50 = 15.46 μM).For the ginsenosides with skeleton A, introduction of a hydroxyl group at the RAfter comprehensively analyzing the inhibitory effects of all tested ginsenosides on both hCES1A and hCES2A, the structure–inhibition relationships could be summarized as follows,O-glycosyl group(s) on the ginsenoside basic skeleton  is unbeneficial for hCES1A inhibition, while introduction of the O-glycosyl group(s) at the R2 site on skeleton A will lead to the loss of hCES2A inhibition. Replacing the O-glycosyl group at the Rʹ site by a hydroxyl group will enhance the inhibition potency of hCES2A and the specificity over hCES1A, but introduction of a hydroxyl group at the R3 site will decrease hCES2A inhibition. These findings will be of advantage to the design and development of more potent and specific ginsenoside-type inhibitors against hCES2A.Collectively, introduction of the Ki values of 2.10 μM and 2.40 μM, respectively  and a modeling hCES2A structure (accession number O00748) as basic models , 47. As 309 via hydrogen bonding in the active site, while the hydroxyl groups at C-2 site and C-7 site of PPT could create strong interactions with several key residuals located in the active site or the Z site of hCES2A, such as MET309, PRO260 or GLN408, via hydrogen bonding. But the similar effects were not shown between hCES1A and hydroxyl groups at C-2 site and C-7 site. As a result, the LibDockScores of these four ginsenosides on hCES2A binding on two distinct ligand-binding sites are similar to each other  on the skeleton A or skeleton C (including the C-2 and C-7 sites) may lead to the loss of hCES1A inhibition, suggesting that the O-glycosyl group at the C-2 and C-7 sites should be replaced by other groups that may create strong interactions with hCES1A. Furthermore, in view of the naturally occurring ginsenosides could be easily available via isolation from herbs or using synthetic biology techniques [O-β-carboxypropionyl or ethyl ester on these ginsenosides, should be used for the development of next generation hCES2A inhibitors bearing the ginsenoside skeleton [From the viewpoint of chemical structure, the ginsenosides have several hydroxyl groups that could be easily modified to generate a series of derivatives. On the basis of SAR of ginsenosides as hCES2A inhibitors, the hydroxyl group at the C-13 site is beneficial for the substrate specificity on hCES2A over hCES1A, which can be used for the development of novel ginsenoside-type hCES2A inhibitors with excellent specificity. Meanwhile, our findings show that the introduction of any chniques , 68, thechniques , 69, whiskeleton . In addiskeleton , 71, 72.O-glycosyl group(s) on the skeleton A or skeleton C (including the C-2 and the C-7 sites) would lead to the loss of hCES1A inhibition. Among all tested ginsenosides, DM and DMG displayed relatively strong inhibitions on both hCES1A and hCES2A. Moreover, the gut metabolites of naturally occurring ginsenosides (such as PPD and PPT) exhibited potent and highly specific inhibition on hCES2A, suggesting that these two compounds could serve as ideal lead compounds to develop novel hCES2A inhibitors. Molecular docking simulations demonstrated that PPD and PPT could be well-docked into both the catalytic cavity and the Z site of hCES2A. In short, our results demonstrated that some ginsenosides exhibit strong to moderate inhibition on hCES2A, which hold great promise in modulating the pharmacokinetic profiles and treatment outcomes of hCES2A-substrate drugs, such as irinotecan. Our findings provided novel insights into the interactions between ginsenosides and CES, which were very helpful for the rational use of traditional herb ginseng in clinic, in order to avoid the potential risks of HDI and HEI via inhibition of hCES.In summary, the inhibitory effects of 25 ginsenosides on hCES as well as the structure–inhibition relationships of ginsenosides as hCES inhibitors, were revealed for the first time. The results clearly demonstrated that the hydroxyl group at the C-13 site is beneficial for the substrate specificity on hCES2A, while the introduction of any Additional file 1: Table S1. The Docking simulations of DM, DMG, PPD and PPT on hCES2A. Fig S1. The dose-inhibition curves of DM, DMG, panaxadiol and panaxotriol against hCES1A-mediated DME hydrolysis in HLM. Fig S2. The dose-inhibition curves of oleanic acid and LPA. Fig S3. The dose-inhibition curves of ginsenosides against hCES2A-mediated FD hydrolysis in HLM. Fig S4. The inhibitory effect of PPT on hCES2A in living HepG2 cells. Fig S5. Docking simulations of DME into hCES1A in the active site. Fig S6. 2D representation of the interactions between DM, DMG and the residuals in the active site of hCES1A. Fig S7. The docking simulations of FD into hCES2A in the active site. Fig S8. 2D representation of the interactions between DM and the residuals in the active site or the Z site of hCES2A. Fig S9. 2D representation of the interactions between DMG and the residuals in the active site or the Z site of hCES2A. Fig S10. 2D representation of the interactions between PPD and the residuals in the active site or the Z site of hCES2A. Fig S11. 2D representation of the interactions between PPT and the residuals in the active site or the Z site of hCES2A. Fig S12. 2D representation of the interactions between PPD and the residuals in the active site or the Z site of hCES1A. Fig S13. 2D representation of the interactions between PPT and the residuals in the active site or the Z site of hCES1A.


Aldh1a2, RA synthesis enzyme expressed in MDs, in other tissues was detected in MDs. When the transcription factor genes were overexpressed in a uterine mesenchymal cell line, C/ebpδ overexpression stimulated Aldh1a2 expression. Furthermore, C/EBPδ protein was strongly expressed in the proximal and middle regions of the MDs and bound to the Aldh1a2 promoter in vivo. Since C/ebpδ mRNA expression was maintained at the same level in proximal, middle and caudal MDs, we hypothesize that a high frequency of mitosis induces a low level protein expression in MD mesenchyme. In fact, the mitotic activity was significantly high in caudal mesenchyme, and a mathematical model showed that a gradient of protein was induced by cell proliferation. Therefore, morphogenesis of MDs controls the fate of mesenchyme via RA degradation in urogenital sinus and a gradient of proteins involved in RA synthesis.In female mice, proximal, middle and caudal Müllerian ducts (MDs) differentiate into oviduct, uterus and vagina, respectively. The fates of female reproductive tract epithelia are determined by the mesenchyme. However, the mesenchymal fate determination system is still unclear. It is reported that presence or absence of retinoic acid (RA) signaling in MD mesenchyme induced uterine or vaginal mesenchyme, respectively. To analyze determination of the borderline, RA signal switching factors were found to play critical roles. Expression of a RA metabolizing enzyme, CYP26A1, was high in the epithelium of caudal MD and urogenital sinus, indicating that the enzyme causes the absence of RA signaling in the region. mRNA expression of some transcription factors regulating   •Degradation of RA by CYP26A1 is involved in fate determination of vaginal mesenchyme.•Aldh1a2 promoter and stimulates the expression.C/EBPδ can directly bind to •Müllerian ductal cell proliferation causes protein gradient involved in RA synthesis. Hoxa9 is expressed at high levels in regions that will become oviducts, Hoxa10 is expressed in the development of uterus, Hoxa11 is expressed in the primordial lower uterus and cervix, and Hoxa13 is expressed in the cervix and upper vagina Hoxa10 deficiency induces an oviduct-like structure only in anterior part of uterus Hoxa13 deficiency causes hypoplastic urogenital genital sinus and agenesis of posterior part of MD, but not affects the differentiation Hox genes are also involved in adult functions of female reproductive tracts (e.g. implantation) Hox genes are mainly responsible for adult function in female reproductive tract, but they are not main factors that determined regional fate of MD mesenchyme.Homeobox (Retinoic acid (RA) is an essential component of cell-cell signaling during organogenesis Rdh10 or Aldh1a2 in MDs were searched. Roles of the candidates in Rdh10 and Aldh1a2 expression are then demonstrated.In the present study, regulation mechanisms of RA metabolism and synthesis were analyzed in MDs. The expression profile of CYP26 in MDs was investigated, and candidates of transcription factors that could be upstream of 22.1ad labium and kept at 22–24°C under 12 h light/12 h darkness by artificial illumination (lights on 08:00–20:00). Animals were maintained in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were approved by our institutional Animal Care Committee. The presence of vaginal plugs indicated embryonic day 0.5 (E0.5). MDs at E12.5 were not selected for male or female. Spleens were dissected from adult mice.Female CD-1 mice  were given a commercial diet and tap water 2.2p53-/- mice at P3 and cultured in 1:1 mixture of Dulbecco modified Eagle medium and Ham nutrient mixture F-12 without phenol red  containing heat-inactivated FBS  at 10%, penicillin  and streptomycin  (10% FBS medium) in a humidified atmosphere of 5% CO2 at 37°C. The cells were passaged using 0.05% trypsin-0.02% EDTA (Sigma-Aldrich).P3US cells were established from uterine mesenchyme of 2.3β-actin was chosen as an internal standard. Ten-thirteen MDs or UGS were pooled in one sample.Total RNA was isolated from MDs and UGS at E12.5 and P0 A or P3US2.4RNA isolation and RT were performed from the middle of MDs at E16.5 as described above. ORFs were amplified with TksGflex DNA Polymerase  with specific primers , and insP3US cells in 60 mm dishes were transiently transfected with 1 μg of these plasmids by Screen Fect A  according to the manufacturer's instructions. Control cells were P3US cells transfected with pcDNA 3.1 Hygro (+) vector. After 48 h, gene expressions in these cell lines were analyzed by RT-PCR.2.5Ten MDs at E14.5 were sampled in 1% Protease Inhibitor Cooktail /PBS, and crosslinked with 1.5% PFA. Fixation was stopped by addition of glycine. The samples were washed with PBS and homogenized. The pellets were lysed in 0.5% NP40 buffer (10 mM Tris (pH8.0), 10 mM NaCl), and then lysed in 1% SDS buffer (50 mM Tris (pH8.0), 10 mM EDTA). The samples were added with ChIP dilution buffer (50 mM Tris (pH8.0), 167 mM NaCl, 1.1% Triton X-100 and 0.11% Sodium Deoxycholate). The chromatins were sheared with a sonicator to an average length of 200 bp.Aldh1a2 TSS (−89 to +11), Aldh1a2 C/EBPδ predicted binding site (+2872 to +3987) and Aldh1a2 conserved region primer  were derived from Gene ID 19,378  and 0.5 μg anti-RNA polymerase II CTD repeat antibody , anti-CEBPδ antibody (Santa Cruz Biotechnology) or normal rabbit IgG (Santa Cruz Biotechnology) were mixed in RIPA buffer and incubated at 4°C overnight. After washing with RIPA buffer, the mixtures and the fragmented chromatins were mixed and incubated at 4°C overnight. Samples were then sequentially washed in RIPA buffer, RIPA buffer containing 500 mM NaCl, LiCl wash solution and TE buffer, and eluted in 0.5% SDS buffer (10 mM Tris (pH 8.0), 300 mM NaCl, 5 mM EDTA) at 65°C for 8 h, then added with protease K at 500 μg/ml and incubated for 1 h at 55°C. Proteins were removed by phenol and chloroform and DNAs were precipitated by ethanol. The DNAs were compared with inputs by quantitative PCR using KOD SYBR qPCR Mix (TOYOBO). D 19,378 .2.6To detect CYP26A1, female mice at E14.5, E16.5 and P0 were fixed in 4% PFA at 4°C for 3 h. The samples were embedded in paraffin and cut into 6 µm sections. To detect C/EBPδ, female mice at E14.5 and spleens for positive control were embedded in O.C.T. Compound and frozen by liquid nitrogen. The cryosections (10 µm) were fixed in 4% PFA at room temperature for 5 min. To detect CYP26A1, the sections were microwaved in 10 mM sodium citrate buffer (pH 6.0) for 10 min. Nonspecific binding was blocked in PBS containing 5% goat serum and 1% BSA for 30 min. The sections were incubated at 4°C overnight with primary antibody for CYP26A1  in PBS containing 0.1% Triton X-100, 1% BSA and 0.1% nonfat dried milk, C/EBPδ  in PBS containing 5% goat serum and 1% BSA, and then incubated with Alexa488-conjugated anti-rabbit IgG goat antibody . For negative control, normal rabbit IgG (Santa Cruz Blotechnology) was used. DAPI was used to stain nucleic acids.To detect cell proliferation, pregnant mice at day of post coitus 14.5 were injected with 100 μg/0.1 ml/20 gBW EdU . After administration for 1 h, female embryos were fixed in 4% PFA at 4°C overnight (n = 5–7). The samples were embedded in paraffin and cut into 6 µm sections. The sections were stained with Click-iT EdU Alexa Fluor 594 Imaging Kit (Thermo Fisher), according to manufacturer’s instructions. The number of EdU-positive cells and Hoechst-stained cells were counted manually. All of epithelial cells in the 1–3 pictures and 200 mesenchymal cells were counted in a section.2.7C”), and frequency of cell proliferation is set as 18, 24 or 30% in proximal , middle (2/5 following region) or caudal region (1/5 last region). mRNA or protein degradation levels are indicated as “d”, and the synthesis levels are indicated as “s”, and d and s are hypothesized at constant.A one dimensional cell-based MD ductal elongation model was generated. In this model, cells line one dimensionally and cell proliferation occurs from proximal to caudal region. C. First,C is averaged each in 100 cells and shows as dots graphically (n = 10).When the number of cells reaches more than 5000, 2.8t-test or Welch’s t-test was used for single comparisons. For multiple comparisons, differences were estimated using ANOVA with appropriate post hoc tests. A statistically significant difference was defined as p ≤ 0.05.Data were expressed as mean ± standard error. Two-tailed Student’s 33.1Cyp26, was investigated in proximal, middle and caudal MDs and UGS at E16.5 and P0 .The expression of RA metabolizing enzyme gene, 5 and P0 B. Cyp26aCYP26A1 protein expression was investigated in cross, sagittal or horizontal sections of MDs and UGS at E14.5, E16.5 and P0 C. CYP26A3.2Rd10 and Aldh1a2, and RA metabolizing enzyme Cyp26a1 was analyzed in MDs from E12.5 to P0. At E12.5, MDs have grown to the caudal region of the developing gonad mesonephros .The effects of the transcription factors on US cells B. OverexAldh1a2 promoter sequence Aldh1a2 promoter sequence in vivo, ChIP assays were performed in MDs at E14.5 (Aldh1a2 transcription start site (TSS). C/EBPδ bound to Aldh1a2 TSS and the promoter region at + 2872 bp, indicating that C/EBPδ directly binds to Aldh1a2 promoter in vivo.Huang et al. reported that C/EBPδ binding sites were on conserved region of at E14.5 B. RNA poLocalization of C/EBPδ protein was investigated in cross sections of MDs at E14.5 by immunohistochemistry C. C/EBPδ3.4C/ebpδ mRNA expression was maintained at the same level in proximal, middle and caudal regions , frequency of mitosis induces a gradient of protein level, but not the due to the value of s, or d = 5.6% . These results suggested that constant synthesized and degraded protein represent gradient expression in MDs.Although  regions A, C/EBPδ regions C. Half-l regions A, B. We  regions C. When t4The present study demonstrated that CYP26A1 was expressed not only in caudal MDs but also UGS connected with MDs, and in MD elongation, high proliferative activity at caudal region caused dilution of proteins involving in RA synthesis (e.g. ALDH1A2 and C/EBPδ). Point mutation of lipoma HMGIC fusion partner-like 2 gene leads failure of the connection to UGS and vaginal agenesis Cyp26a1 mRNA was expressed in UGS, and CYP26A1 protein was expressed in all regions of MD mesenchyme and the epithelium of caudal MD and UGS. In contrast, the expression of ALDH1A2 and RA signaling was high in proximal and middle MDs Cyp26a1 expression levels were 15.5 ± 1.7 at diestrus and 48.8 ± 12.9 folds at estrus compared with that in middle MD Cyp26a1 expression level was 168.7 ± 66.9 folds in UGS compared with middle MD, suggesting that CYP26A1 level was sufficiently high for degradation of RA in UGS as well as in adult uterus. Thus, CYP26A1 was involved in the fate determination of vaginal mesenchyme. In caudal MD, ALDH1A2 protein is slightly detected in the mesenchyme Aldh1a2 expression, and it was directly bound to Aldh1a2 promoter in vivo, indicating that C/EBPδ directly regulates Aldh1a2 expression in MDs. However, the expression of C/ebpδ and Aldh1a2 showed reverse correlation in MD development (Aldh1a2 expression. Aldh1a2 was already expressed in mesonephros at E7.5-E10.5 Osr1-expressing cells Aldh1a2 expression is essential for mesonephric development; therefore, in MDs, the system may be diverted to the mesenchymal differentiation.C/EBPδ stimulated elopment A, and feIn human, some disorders are reported for atresia and agenesis of vagina and cervix


E. coli using standard molecular biology techniques with refined and rapid one-step analytical purification techniques. The peptide is isolated and purified from transformed cells using an optimized reverse-phase high-performance liquid chromatography (HPLC) protocol with commonly available C18 columns, yielding high amounts of peptide (~15–20 mg per 1 L culture) within a short period of time. The recombinant human Aβ(M1-42) forms characteristic aggregates similar to synthetic Aβ42 aggregates as verified by western blotting and atomic force microscopy to warrant future biological use. Our rapid, refined, and robust technique produces pure recombinant human Aβ(M1-42) that may be used to synthesize chemical probes and in several downstream in vitro and in vivo assays to facilitate Alzheimer’s disease research.Amyloid plaques found in the brains of Alzheimer’s disease patients primarily consists of amyloid beta 1-42 (Aβ42). Commercially, Aβ42 is synthesized using high-throughput peptide synthesizers resulting in the presence of impurities and the racemization of amino acids that affects its aggregation properties. Furthermore, the repeated purchase of even a small quantity (~1 mg) of commercial Aβ42 can be expensive for academic researchers. Here, we describe a detailed methodology for robust expression of recombinant human Aβ(M1-42) in Rosetta(DE3)pLysS and BL21(DE3)pLysS competent  Amyloid beta 1-42 (Aβ42) is a small ~4 kDa peptide produced when the amyloid precursor protein expressed on neuronal membranes is sequentially cleaved by β-secretase and γ-secretase . The extSeveral therapeutics currently in preclinical and clinical trials for AD are focused on targeting the cellular and molecular mechanisms related to Aβ42 ,6. Thus,E. coli cells Mica sheet Aluminum coated silicon probes with resonant frequency ~300 kHz and 40 N/m force constant Phosphate-buffered saline (PBS), 10x at pH 7.4 Benchtop incubator shaker (New Brunswick™ Excella® E24) Sonicator Ultrasonic Homogenizer (125W) with 1/4" Probe 2 Incubator  COCentrifuge  with a Swinging-Bucket Rotor (BIOFlex™ HC) UV-Visible Spectrophotometer Microplate reader for nucleic acid quantification (Take3™ Micro-Volume plate) with Gen5 Software Combiflash EZ prep UV/ELSD RediSep Prep 10 × 250 mm C18 100A, 5 µm column RediSep Prep Guard 20 × 30 mm, C18Aq, 100A, 5 µm GenPure UV/UF × CAD plus Ultrapure Water Purification System Rotary evaporator and water bath Dry Bath with heating block Labconco Freezone 12 Liter Console Freeze Dry System (Lyophilizer) Voyager-DE PRO (MALDI-TOF mass spectrometer) For AFM: Veeco Multimode instrument with NanoScope V controllerUse ultrapure water to prepare all the solutions in the protocol. Make liquid Luria Broth (LB) by dissolving 10 g tryptone, 10 g NaCl, and 5 g yeast extract in 1 L water. Autoclave the liquid LB and bring to room temperature before adding the antibiotics. Prepare solid LB as 3.2 g of LB agar mix in 100 mL water. Autoclave the broth and allowed to cool before adding the antibiotics as follows: (i) 100 mg/L ampicillin for antibiotic selection during amplification of the pET-Sac-Aβ(M1-42) plasmid in LB media and (ii) 100 mg/L ampicillin and 34 mg/L chloramphenicol for maintenance and expression of the Aβ(M1-42) peptide in Rosetta(DE3)pLysS and BL21(DE3)pLysS strains . For solid LB preparation, pour the media with the antibiotics on to the Petri dishes and keep in the biosafety cabinet with lids open to allow the media to solidify. Prepare these plates in bulk and refrigerate at 4 °C for future use to save time during the protocol. For cell lysis steps, make two buffers: (i) Buffer A containing 10 mM Tris/HCl and 1 mM EDTA in water (pH 8.0), (ii) Buffer B containing 8 M urea, 10 mM Tris/HCl, and 1 mM EDTA in water (pH 8.0). Prepare fresh buffers each time. For peptide purification with preparative HPLC, make: (i) Solvent A as 0.1% trifluoroacetic acid (TFA) in water and (ii) Solvent B as 0.1% TFA in acetonitrile . TFA is Note: The bacterial growth on solid agar as well as in liquid culture are performed at 37 °C temperature which is the optimal growth temperature for E. coli cells.Streak the bacteria onto a solid LB agar plate containing 100 mg/L ampicillin using a sterile loop.Note: The pET-Sac-Aβ(M1-42) plasmid arrives as a bacterial agar stab culture.Keep the plate overnight for ~ 16 h in the incubator at 37 °C for the colonies to grow.The next day, pick a single colony from the plate A using aKeep the culture in a shaking incubator at 220 rpm and 37 °C overnight for ~ 16 h.The next day, isolate the plasmid , Part 1 Note: Typical plasmid yield is around 45-55 ng/µL from 5 mL liquid LB culture.Measure the concentration of the plasmid at 260 nm absorbance using a spectrophotometer for nucleic acid quantification. 1.E. coli (To transform the Aβ(M1-42) plasmid into E. coli , Part 2,2.Once thawed, add around 1–2 μL of the isolated plasmid  to the cells and gently flick the tube a few times to mix the plasmid with the cells.3.Incubate the cells and the plasmid mixture on ice for 20 min and then place the tube in a 42 °C water bath for 45 s to facilitate the transformation of plasmid into the cells via heat shock method.4.After heat shock, immediately place the tubes on ice for 2 mins.Inoculate the transformed bacteria into 500 μL of liquid LB media without any antibiotics and keep the tube in the 37 °C shaker for 1 h at 220 rpm. CRITICAL STEP: The transformed bacteria is first inoculated in liquid LB media without any antibiotics for 1 h to allow the bacteria to express the antibiotic resistance proteins necessary for future steps.1.After 1 h, spread around 25–30 μL (out of 500 μL) of the transformed cells onto solid LB agar plates containing 100 mg/L ampicillin and 34 mg/L chloramphenicol using a sterile glass spreader.2.Let the plates sit in the biosafety cabinet for 5 min to allow the cells to absorb on the solid LB.3.Next, keep the plates overnight in the incubator at 37 °C for the transformed colonies to grow. CRITICAL STEP: It is best to use a lower volume of the transformed cells (25–30 μL) as this results in more single colonies that are easier to pick and prevents overcrowding on the plate.4.The next day, pick a single colony from the plate using a sterile loop or a sterile 10 μL pipet tip and inoculate into 5 mL of liquid LB containing 100 mg/L ampicillin and 34 mg/L chloramphenicol (first culture).5.OPTIONAL STEP: At the same time, pick another single colony of the transformed bacteria and inoculate into a second 5 mL liquid LB media containing 100 mg/L ampicillin and 34 mg/L chloramphenicol for overnight growth (second culture). This culture will be used to make frozen glycerol stocks for future use.6.Place both the cultures in the shaking incubator at 220 rpm at 37 °C overnight (16 h) B.7.The following day, inoculate the first 5 mL culture into a large 2 L Erlenmeyer flask containing 1 L liquid LB media with 100 mg/L ampicillin and 34 mg/L chloramphenicol C.8.600).Keep this 1 L culture at 220 rpm and 37 °C until the cell density reaches an optical density (OD) value between 0.40 to 0.45 at 600 nm (OD CRITICAL STEP: The BL21(DE3)pLysS cells reach an OD600 between 0.40 to 0.45 in 3 h time, whereas the Rosetta(DE3)pLysS cells requires around 3.5–4.0 h to reach an OD600 between 0.40 to 0.45. It is best to measure the OD of the culture at regular 20 min intervals starting from the 3-h time point before proceeding to the next step.9.OPTIONAL STEP: The second 5 mL culture of transformed bacteria is used to make 25% glycerol stocks by adding 500 μL of 50% glycerol to 500 μL bacterial culture and is frozen at −80 °C for future use. A −80 °C frozen glycerol stock of the transformed bacteria is thawed for use in the future for inoculating 5 mL liquid LB containing 100 mg/L ampicillin and 34 mg/L chloramphenicol to grow additional cultures of the transformed bacteria containing the Aβ(M1-42) plasmid. Note: Frozen aliquots of 250–500 µL glycerol stocks may be stored for future use. It is recommended that the stored aliquots of glycerol stock be thawed 1–2 times only. Additionally, some of the frozen stock may be scrapped and thawed for inoculation. CRITICAL STEP: This step serves as a starting point for all future experiments performed for the isolation of the Aβ(M1-42) peptide  to obtain a final concentration of 0.1 mM in 1 L of the liquid LB media. Note: Add 1 mL of 0.1 M IPTG stock solution prepared in water to obtain a final concentration of 0.1 mM in 1 L LB media.Once the ODNote: IPTG induction must occur early in the exponential growth phase to form insoluble inclusion bodies, which are essential for the isolation procedure.11.Keep the culture again on the shaking incubator at 220 rpm and 37 °C for additional an 4 h in the presence of IPTG to allow the cells to express the Aβ(M1-42) peptide.12.After 4 h, centrifuge the 1 L culture at 7068× g at 4 °C for 25 min.Note: The cultures are centrifuged at 4 °C temperature to arrest cell growth and metabolism.Note: Our centrifuge allowed for a maximum speed of 7068× g with the corresponding swinging bucket rotor  three more times.6.Resuspend the pellet in 20 mL of freshly prepared Buffer B and sonicate as above until the solution appears clear. CRITICAL STEP: Purification should be performed immediately after the peptide is dissolved in 8 M urea solution (Buffer B). Extended exposure of the peptide to urea is known to cause carbamylation of lysine residues. CRITICAL STEP: Due to the inconsistencies of mechanical lysis using the sonicator, the solution may appear to be cloudy. Note: Prepare fresh Buffer B every time. The pH of the buffers is critical for the complete dissolution of the peptide fraction.7.Centrifuge the solution at 7068× g for 25 min at 4 °C to remove any foam that may have appeared during the sonication step and to pellet unwanted insoluble cell debris, if any. Finally, filter the supernatant through a 0.22 μm non-sterile hydrophilic PVDF syringe filter using a 30 mL syringe to obtain a clear solution. 8.OPTIONAL STEP: Prior to purification, MALDI-TOF MS may be performed on the urea-solubilized recombinant peptide to confirm the presence of Aβ(M1-42) in the solution (data not shown). Since the solution contains additional salts, the solution must be passed through a C18 Zip Tip resin to remove the salts in order to obtain a clear spectrum.lubility .1.To lyEquipment set-up for peptide purification: Heat a water bath containing reverse osmosis water using a commercially-available sous vide between 60 to 80 °C  typically elutes at 26 min.Run the solvent gradient described in Upon completion of the purification protocol, clean the column with the solvent gradient described in Repeat the cleaning and purification steps three more times or until all the solution is used.Combine the collected Aβ(M1-42) fractions eluted at 26 min .Evaporate off the acetonitrile under reduced pressure at 65 °C using a rotary evaporator until a cloudy aqueous solution remains.Freeze the solution at −80 °C and then submerge in liquid nitrogen for 5 min.Perform overnight lyophilization at −90 °C at 0.003 mbar pressure to obtain the white Aβ(M1-42) powder .PAUSE STEP Store the lyophilized peptide at −80 °C until further characterization.Place the solvent lines from the CombiFlash HPLC instrument into the Solvent A and Solvent B bottles. Next, prime the system with Solvent B followed by Solvent A to clear the solvent lines of any previous residual solvents. Then clean the column by injecting 4 mL of Buffer B into the 5 mL injection loop C and runDissolve a small quantity of Aβ(M1-42) in water and dilute in water until the sample is approximately 100 μg/mL.Thoroughly mix 1 μL of the Aβ(M1-42) solution  with 1 μL of α-cyano-4-hydroxycinnamic acid (CHCA) matrix solution (10 mg/mL CHCA in 0.1% TFA).Spot the analyte/matrix mixture on a MALDI target plate and allow to dry.Obtain the MALDI-TOF mass spectra on a Voyager-DE PRO from 1000–22000 Da in the positive ion mode with an accelerating voltage of 25,000 V .Further dilute the analyte solution prepared for the MALDI-TOF MS and inject into the Agilent 6550 iFunnel Q-TOF LC-MS in positive mode and fragment using electrospray ionization (ESI) with a fragmentor voltage of 175V .To perform western blotting, the lyophilized peptide from HPLC was dissolved in 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP) to prepare monomers of Aβ(M1-42) as per the previously established protocols ,17.When the peptide is dried overnight in the chemical hood, dissolve 1 mg of the peptide in 221 μL DMSO to obtain a final concentration of 1.0 mM.Load the peptide on 12% Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE) at different concentrations  and run the gel at 115 V for 1 h and 20 min or until the loading buffer line reaches the bottom of the gel.Transfer the gel to a nitrocellulose membrane at 10 V for 35 min.After transfer, boil the membrane in PBS for 5 min, then incubate on a rocker with blocking buffer ) for 1 h.After blocking, incubate the membrane in the blocking buffer containing the 6E10 monoclonal antibody (with target specificity to the human Aβ peptide) overnight on a rocker at 4 °C.The next morning, wash the membrane 3 times for 10 min each with TBST and incubate with the secondary antibody (HRP-conjugated goat anti-mouse antibody) on the rocker for 1 h.Washed again three times for 10 min each with TBST and develop in the dark room using chemiluminescence reagents as per the manufacturer’s protocol.Run the synthetic Aβ(1-42) in the same gel with the same protocol for comparison .Preparing Aβ(M1-42) monomers: The synthetic Aβ(1-42) and the lyophilized Aβ(M1-42) powder are used to prepare 1 mM solution with 215 μL of 1,1,1,3,3,3-Hexafluoro-2-propanol (HFIP). Handle the HFIP carefully using a 1 mL glass Hamilton syringe with a Teflon plunger and a sharp needle. Start by incubating the clear Aβ(M1-42) and synthetic Aβ(1-42) solution at room temperature for 30 min. Next, transfer 100 μL aliquots (~0.45 mg) to microcentrifuge tubes and leave the tubes open in the fume hood overnight for the HFIP to evaporate. The next morning, dry the sample under high vacuum for 1 h without heating to remove any remaining traces of HFIP and moisture. This results in thin clear films of monomeric peptides at the bottom of the tubes which are stored at −80 °C until further use.1.To evaluate the aggregation property of the recombinant Aβ(M1-42) with respect to the synthetic Aβ(1-42), the protocol established by Stine et al. is used to prepare Aβ oligomers and fibrils ,17.2.In brief, allow the tubes containing monomeric peptide films to equilibrate at room temperature for a few minutes and prepare 5 mM of synthetic Aβ(1-42) and recombinant Aβ(M1-42) DMSO stocks by adding 20 μL of cell-grade DMSO to each tube containing ~0.45 mg of the peptide.3.Pipet the solution thoroughly by scraping down the sides of the tube and vortexed for ~30 s followed by bath sonication for 10 min to ensure complete resuspension of the peptide film. CRITICAL STEP: This stock solution is used immediately as the starting material for oligomeric Aβ preparation.4.To prepare Aβ oligomers, mix 2 μL of the freshly resuspended 5 mM of synthetic Aβ(1-42) and recombinant Aβ(M1-42) in DMSO with 98 μL of 1× PBS (filtered) to make 100 μM solution. To prepare Aβ fibrils, mix 98 μL of 10 mM HCl (filtered) to 2 μL of freshly resuspended 5 mM of recombinant Aβ(M1-42) in DMSO.5.Vortex the solution thoroughly for 15 s and incubate at 4 °C (for oligomers) and at 37 °C (for fibrils) for 24 h.6.After 24 h, prepare the samples for AFM with proper sterile technique in the hood as follows: Dilute the 100 μM samples to a concentration of 30 μM in filtered water.7.Mount the mica sheet on 15 mm stainless steel pucks.8.Immediately before sample plating, remove a few layers of the mica sheet using adhesive tape to reveal a featureless surface for the absorption of the peptide.9.Next, pretreat the mica surface with ~5–8 μL of filtered 1M HCl for 30 s and rinse with 2–3 drops of ultrapure water (filtered) using the 1 mL syringe.10.CRITICAL STEP: Hold the mica plate at a 45° angle to wash with drops of water. Immediately after cleaning, spot the peptide sample onto mica and incubate for 3 min.11.Gently rinse the mica with 2–3 drops of water using the 1 mL syringe and dry with several gentle pulses of clean compressed air or nitrogen gas.12.Incubate the samples at room temperature for a few hours until imaging.13.Perform the AFM imaging using a multimode AFM equipped with aluminum-coated silicon probes with ~300 kHz resonant frequency and 40 N/m force constant under the tapping mode.14.Finally, perform image analysis using the NanoScope Analysis software.E. coli using standard molecular biology techniques, such as transformation and bacterial cell culture. The major differences among these methodologies have been the isolation and purification methods used to obtain purified peptide samples. The purification of the peptides expressed from cells is challenging due to the many different techniques proposed combined with limited resources available in a traditional biochemistry or chemical biology laboratory. For example, previously, nickel affinity chromatography was used to purify Aβ42 fusion proteins with N-terminal His-affinity tag [E. coli [A handful of methodologies have been published in recent years that show expression of Aβ42 in nity tag  or as annity tag . Both th[E. coli . This pu[E. coli  used ion[E. coli  used pre+  and synthetic Aβ42, we demonstrated that the purified peptide mixture contains large quantities of monomers as seen in the 4 kDa region . A higheThus, the Aβ(M1-42) peptide could be used for several downstream in vitro and in vivo applications such as cell-based drug screening, neuroinflammation cell culture, animal models, etc., and for the synthesis and development of novel Aβ-related biorthogonal chemical and fluorescent probes to facilitate the advancement of neurological disease research.


ABI5 are still present.Basic region/leucine zippers (bZIPs) are transcription factors (TFs) encoded by a large gene family in plants. ABF3 and ABI5 are Group A bZIP TFs that are known to be important in abscisic acid (ABA) signaling. However, questions of whether ABF3 regulates ABI5 activation by either ABF3 alone or by co-expression of ABF3 and 14-3-3ω. Moreover, chromatin immunoprecipitation-qPCR results showed that in Arabidopsis, ABI5 ABA-responsive element binding proteins (ABREs) of the promoter region (between − 1376 and − 455) were enriched by ABF3 binding under normal and 150 mM NaCl salt stress conditions.In vitro kinase assay results showed that Thr-128, Ser-134, and Thr-451 of ABF3 are calcium-dependent protein kinase phosphorylation sites. Bimolecular fluorescence complementation (BiFC) analysis results showed a physical interaction between ABF3 and 14-3-3ω. A Thr-451 to Ala point mutation abolished the interaction but did not change the subcellular localization. In addition, the Arabidopsis protoplast transactivation assay using a luciferase reporter exhibited ABI5 expression is regulated by ABF3, which could contribute to salt stress tolerance in Arabidopsis thaliana.Taken together, our results demonstrated that The online version of this article (10.1186/s40529-019-0264-z) contains supplementary material, which is available to authorized users. RD29B (Shinozaki and Yamaguchi-Shinozaki DREB2. DREB2 belongs to the DRE BINDING PROTEIN/C-REPEAT BINDING FACTOR family that binds to the dehydration-responsive element/C-repeat (DRE/CRT) element. RD29A gene activation is both ABA-dependent and ABA-independent. DREB transcription factor binds to the DRE/CRT element of a downstream gene, such as the RD29A gene, which results in up-regulation of this gene under drought stress, which eventually leads to enhanced salt or osmotic stress tolerance in plants.Salt stress has recently become a serious problem that causes decreased crop yields worldwide and is caused by global climate change. The components of salt stress signaling in plants have, therefore, become important topics in recent years. The Salt Overly Sensitive (SOS) pathway was activated to confer salt tolerance in plants under conditions of salt stress Zhu . In addicis-elements , constitutive OsbZIP46 activation conferred drought stress tolerance  GmbZIP1 gene improved high salt stress tolerance in transgenic plants  ZmbZIP72 gene conferred salt stress tolerance in Arabidopsis transgenic plants  SIAREB gene improved drought and high salt stress tolerance in the respective transgenic plants  plants, the LrbZIP gene was shown to be important in the salt resistance of roots   assay was used to confirm the interaction between ABF3 and 14-3-3ω, and the transactivation assay was used to investigate if ABF3 regulates ABI5. Promoter deletion was also tested in the transactivation assay. Moreover, chromatin immunoprecipitation-qPCR was also introduced. Our results showed regulation of ABI5 expression by ABF3 in response to salt stress in Arabidopsis thaliana.The abscisic acid responsive element-binding factor 3 (ABF3) is a member of the group A bZIP TFs. Arabidopsis thaliana ecotype Columbia was used in the present study. A T-DNA insertion mutant line, abi5, was obtained from Dr. Hsu-Liang Hsieh‘s lab at the National Taiwan University. A T-DNA insertion mutant line, abf3 (SALK_096965) of ABF3 gene (At4g34000), was ordered from the Arabidopsis Biological Resource Center (ABRC)  agar medium containing 0.8% sucrose . Seven-day-old seedlings were treated with 100 mM or 150 mM NaCl by transferring the seedlings to plates containing 1/2 MS medium and NaCl, and the plants were incubated for either 0 h, 0.5 h, 1 h, or 3 h before RNA extraction.g  for 15 min and moved to a new 1.5 ml tube. Five hundred microliters of isopropanol were added, and the samples were centrifuged at 12,000×g  for 10 min. The pellets were washed with 75% EtOH and resolved using DEPC-H2O. The contaminating DNA was removed using TURBO DNA-free™ DNase according to the manufacturer’s instructions . Isolated RNA was used for cDNA synthesis using the iScript™ cDNA Synthesis Kit . Real-time PCR was performed using CFX and CFX manager software . SsoFast™ EvaGreen Supermix  was used for amplifications. Arabidopsis ACTIN2 was used as a quantitative control.Seedlings (10–100 mg) were ground into a powder with liquid nitrogen, and 1 ml RezolTM C&T  with 200 μl chloroform was added. The samples were centrifuged at 12,000×2, 5 mM β-mercaptoethanol, and 0.1% BSA) by dipping both sides of the strips. The tissues were then placed under a vacuum for 30 min. The digestion reaction continued for at least 3 h at room temperature with shaking. The enzyme solution should turn green after a gentle swirling motion, which indicates the release of protoplasts. A clean filter paper was washed with a W5 solution  before protoplast filtration. The enzyme solution containing protoplasts were filtered after wetting the filter paper. The flow-through was centrifuged at 100g  to pellet the protoplasts in a 15 ml round-bottomed tube for 3 min. The supernatants were removed, and protoplast pellets were washed by gentle swirling with the W5 solution. Protoplasts were resuspended in the W5 solution after counting cells under the microscope using a hemocytometer . The protoplasts were kept on ice for 30 min and then resuspended in an MMG solution  to a concentration of 2.5 × 105 protoplasts/ml.Arabidopsis protoplast extraction was carried out as previously described and modified , driven by the 35S promoter and fused to the YFP-N or YFP-C in frame. The YN and YC fragments of YFP were fused to the C-terminus of the full-length ABF3 cDNA, and 14-3-3ω, respectively. The paired plasmids were transfected into Arabidopsis protoplasts . ACS7 was used as a positive control for the 14-3-3 interaction   for 3 min to pellet the protoplasts. The supernatants were removed, and the protoplasts were washed twice with the W5 solution. The protoplasts were resuspended with 100 μl of the W5 solution in an Eppendorf tube at room temperature. After 12–16 h, YFP fluorescence was detected using a confocal microscope .BiFC analyses were carried out by a modified method, as previously described , ABF3 (T128A), ABF3 (S126A), and ABF3 (S134A) that were constructed into the pRTL2 vector using the Gateway LR Clonase™ II Enzyme Mix . The PRL plasmid containing the Renilla Luc gene driven by the CaMV35S promoter was used as an internal control for the transactivation assay. Arabidopsis protoplasts were isolated and transfected by a modified polyethylene glycol method, as previously described . Cells were assayed for luciferase activity using the Dual-Glo™ Luciferase Assay System (Promega) following the manufacturer’s instructions.For the reporter gene construct, the 5× GAL4, 4X GCC, and TATA box in the 5× GAL4-4X GCC-TATA-LUC-Nos M13Fprimer vector was replaced by the 32P] ATP) was added into the reaction mixture, which consisted of a 0.25 μg purified recombinant calcium-dependent protein kinase, 5 μg fusion protein substrate, and standard kinase reaction buffer to start the kinase reaction. The reaction tubes were incubated at RT for 10 min, and the reactions were stopped by adding 5 μl of a 5× sodium dodecyl sulfate (SDS) sample buffer. All samples were loaded into the 10% SDS-PAGE loading well for electrophoresis, and the gel was air-dried with acetate sheets. The γ-32P-labeled signals were normalized with the amount of protein as determined from Coomassie Brilliant Blue stained gels after running SDS-PAGE. The γ-32P-labeled signals were detected with an image analyzer (Typhoon 9400).Purification of calcium-dependent protein kinasea (CDPKs)  and substrates (wild type and mutant variants of GST-GFP-Strep-tagged fusion peptides as shown in Table −ΔCt(= 2−[Ct(ChIP)−Ct(Input)]). ACTIN2 and UBQ10 were used as the negative control.Two-week-old seedlings grown on vertically oriented plates with MS medium were collected (~ 2 g) for the ChIP assays  was used to analyze ABI5 gene expression in the abf3 mutant. The stress marker gene, DREB1A, was up-regulated in 100 mM and 150 mM NaCl salt stress conditions. Under 100 mM or 150 mM NaCl salt stress condition, ABI5 was up-regulated in the abf3 mutant. However, the expression levels were still lower than wild type under salt stress conditions  in ABF3 was tested to see if there was an interaction with 14-3-3. No interaction between ABF3 (T451A) and 14-3-3 was detected Fig. , who shoArabidopsis protoplasts. The YFP fluorescent signal was still located in the nucleus for both the ABF3 and mutated ABF3 (T451A) , which lacked the conserved 14-3-3 binding sequence, was YFP-fused and transfected into 1A, whichABI5 gene promoter (started from the translation start site) was analyzed by PLANTCARE (http://bioinformatics.psb.ugent.be/webtools/plantcare/html/) or AGRIS (http://arabidopsis.med.ohio-state.edu). Four ABREs were predicted from − 958 to − 2295 of the ABI5 promoter region. To investigate the relationship between ABI5 and ABF3, a transactivation assay was performed. The reporter plasmid harboring the ABI5 promoter (2055 bp) sequence fused with a firefly luciferase reporter gene was constructed. Effector plasmids were constructed with the ABF3 and 14-3-3ω genes each driven by the CaMV35S promoter. The PRL plasmid containing the Renilla luciferase gene driven by the CaMV35S promoter was used as an internal control. In the transactivation assay, the reporter, effector, and internal control plasmid were co-transfected into Arabidopsis protoplasts, and luciferase signal detection was used to determine transcriptional regulation. Transient luciferase expression indicated that ABI5 was activated when ABF3 alone or ABF3 combined with 14-3-3ω were expressed  was used as an effector and luciferase expression was observed. Compared with the wild type ABF3, mutated ABF3 (T451A) combined with 14-3-3ω did not activate ABI5  analyses, five predicted ABRE cis-elements (CACGTG) -tagged ABF3 driven by the 35S promoter (35S:ABF3-YFP). To study which ABRE cis-elements can be bound by ABF3, the ABI5 promoter was divided into BOX 1 to BOX 5, as shown in Fig. ABI5 is a direct target gene of ABF3.Since ant Fig. , we hypoSolanum tuberosum), StABF1 is phosphorylated in response to ABA and salt stress. Specifically, a potato CDPK isoform (StCDPK2) phosphorylated StABF1 in vitro  and SnRK2 phosphorylation motifs (LXRXX(S/T)  has been shown to be phosphorylated in an ABA-dependent manner  in their promoter regions .