News Article | May 22, 2017
Three-dimensional interconnections in electronic devices are now so small and intricate that they cannot be imaged without destroying them in the process. This leaves a ‘metrology’ gap between the design of devices and the actual output of manufacturing. But a technique known as ptychographic X-ray computed tomography (PXCT) that uses X-rays instead of light or electrons to examine samples non-destructively could hold the answer, suggests a team of researchers. Mirko Holler and colleagues from the Paul Scherrer Institute used the synchrotron beam at the Swiss Light Source to examine integrated circuits (ICs) in a new way [Holler et al., Nature (2017), doi: 10.1038/nature21698]. “ICs are typically investigated with focused ion beam scanning electron microscopy (FIB SEM) methods, which are destructive,” explains Holler. “Such imaging causes artifacts, so-called curtaining, because the cutting does not produce a flat surface.” Instead of taking physical slices through a sample, PXCT takes of many (hundreds of thousands) of individual X-ray diffraction patterns, where a coherent beam of X-rays is scattered by the sample. The multiple images are then added together – or reconstructed – into three-dimensional, real-space renditions of density variations in the sample. In contrast to conventional scanning imaging techniques, the resolution is not determined by the diameter of the beam or the optics of the microscope but by noise levels in the signal and the accuracy with which the sample is positioned. “In our case, imaging artifacts do not occur, and the resolution and measurement time is comparable [to FIB SEM],” says Holler. Holler and his colleagues put the technique through its paces using a microchip of known design and then examined a commercial chip of unknown design. The reconstructed three-dimensional images allow the identification of the key parts of the chip, as well as manufacturing artifacts. The technique is sensitive enough to detect regions of silicon doped with n- or p-type impurities. With its resolution of less than 15 nm, PXCT has sufficient contrast and sensitivity to allow imaging of even the smallest circuitry but the time required to acquire images is long. With the advent of more advanced synchrotron systems, Holler believes that the resolution and/or imaging speed of PXCT can be improved dramatically. “X-ray imaging can now produce images of high quality and resolution of ICs that allow seeing the individual transistors,” says Holler. “I believe that the state-of-the-art FIB SEM methods used in chip inspection will be replaced by PXCT imaging.” The researchers believe the technique could support the optimization of production processes, identification of failure mechanisms, and validations of microchips. While the technique currently needs a small sample to be removed from an IC, the team is working on an approach whereby a whole two-dimensional chip could be imaged in three-dimensions. Jianwei (John) Miao of the University of California, Los Angeles believes that the real novelty of the method lies in its ability to non-destructively image heterogeneous features buried in a three-dimensional material at very high spatial resolution. However, he points out that the current work was performed using a third generation synchrotron radiation source. “As a coherent diffractive imaging technique, PXCT requires a coherent X-ray source (i.e. the beam is parallel and the bandwidth is narrow). To make this technique widely accessible to both academia and industry, one needs to implement it using tabletop coherent X-ray sources. Presently, several groups around the world are developing such X-ray sources," he says.
News Article | September 28, 2016
If not stated otherwise, all chemicals and reagents were obtained from Sigma Aldrich. Restriction enzymes were obtained from New England Biolabs. The AquaMet catalyst was purchased from Apeiron Synthesis S.A. All plasmids used in this study are collated in Supplementary Table 1. To construct the periplasmic expression vector for SAV, the gene for T7-tagged SAV was amplified by polymerase chain reaction (PCR) from pET-11b-SAV31 using primers 1 and 2 (Supplementary Table 2) to add the 21 amino acid OmpA signal peptide (MKKTAIAIAVALAGFATVAQA) to the N terminus of SAV. The PCR product was digested with restriction enzymes NdeI and BamHI, gel purified and ligated into the target vector pET-30b(+) (Merck Millipore) pre-treated with the same enzymes. The resulting expression vector, designated pET-30b-SAVperi, carries the gene for the OmpA::SAV fusion protein under the control of a P promoter. To construct a comparable cytoplasmic expression construct, the gene for T7-tagged SAV from pET11b-SAV was PCR-amplified without adding additional amino acids using primers 3 and 2 (Supplementary Table 2) and subsequently cloned into pET-30b(+) by restriction digest (NdeI and BamHI) and ligation, resulting in plasmid pET-30b-SAVcyto. All strains used in this study are summarized in Supplementary Table 3. A strain for periplasmic expression was constructed that combines the ease of library generation with the compatibility with the T7-expression system. Therefore, the gene of the T7 RNA polymerase was integrated into the chromosome of E. coli TOP10 (F− mcrA Δ(mrr-hsdRMS-mcrBC) φ80lacZΔM15 ΔlacX74 nupG recA1 araD139 Δ(ara-leu)7697 galE15 galK16 rpsL(StrR) endA1 λ−, Thermo Fisher Scientific) using the λDE3 Lysogenization Kit (Merck Millipore). The resulting lysogen was designated E. coli TOP10(DE3). E. coli TOP10(DE3) containing the respective expression plasmid for SAV (pET-30b-SAVperi or pET-30b-SAVcyto) was cultivated in a Luria–Bertani (LB) medium (50 ml)32 supplemented with kanamycin (50 mg l−1) in shake flasks (500 ml, 37 °C, 200 r.p.m.). An LB pre-culture was diluted 1:100 in fresh medium and incubation was performed until an optical density at 600 nm (OD ) of about 0.5 was reached. Subsequently, the cultivation temperature was lowered to 20 °C, and the expression of SAV was induced by addition of isopropyl-β-d-thiogalactopyranoside (IPTG, 50 μM) and cells were harvested by centrifugation (6,000 r.c.f., 2 min) after four hours of induction. The cell pellet was further processed either for fractionation to analyse the cellular protein content or for flow cytometry. To separate the periplasmic and the cytoplasmic fraction of cellular proteins, the PeriPreps Periplasting Kit (Epicentre Technologies) was used. For in-gel staining of SAV, biotin-4-fluorescein (10 μM) was added to the respective fractions before sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS–PAGE) analysis. Owing to the binding to the biotinylated fluorescent dye, SAV can be visualized under ultraviolet light in the gel (before staining with Coomassie blue). Owing to its high stability, SAV remains tetrameric and fully functional even under the otherwise denaturing conditions of SDS–PAGE. To stain SAV in whole cells of E. coli, the pellet of a shake-flask expression culture was resuspended in phosphate buffer saline (PBS32) to a final OD = 1 before the addition of Atto-565-biotin (2 μM, Atto-Tec GmbH). Cells were incubated on ice (30 min) and subsequently subjected to three wash cycles by centrifugation (6,000 r.c.f., 2 min) and resuspension of the pellet in PBS. The resulting cell suspensions were analysed on a BD LSR Fortessa SORP (BD Biosciences) using a 561-nm laser for the excitation of Atto-565-biotin and a 610/20BP-600LP filter combination for analysis of the emitted fluorescence signal (peak area). The histograms displayed comprise data of 100,000 ungated events for each sample. For expression of SAV (and mutants thereof) in the 96-deepwell format, LB medium (500 μl) supplemented with kanamycin (50 mg l−1) was inoculated from a single colony and grown until stationary phase (37 °C, 300 r.p.m., 50-mm shaking amplitude). This pre-culture (aliquot of 30 μl) was used to inoculate a main culture (1 ml) in a modified ZYM-5052 medium33 that lacked lactose as auto-induction agent, but contained kanamycin (50 mg l−1). Induction was performed by the addition of IPTG ([IPTG] = 50 μM) and continued for four hours before harvesting. Subsequently, a fraction of the cultures (100 μl) was set aside for OD determination and the remaining suspension was subjected either to ICP-OES quantification or to metathesis with whole cells. The activity of the artificial metalloenzyme biot-Ru–SAV was quantified in the reaction buffer (100 mM sodium acetate, pH 4.0, 0.5 M MgCl , 2.5% DMSO) both in the presence and absence of 10 equiv. of glutathione (GSH or GSSG; 500 μM) relative to the amount of the cofactor biot-Ru (50 μM). The reaction mixture (total volume of 200 μl) was incubated shaking (16 h, 37 °C) and, after the reaction, 800 μl of methanol containing 2-phenylethanol (final concentration of 100 μM) as an internal standard was added. After centrifugation (5 min, 21,000 r.c.f., 4 °C), 500 μl of the supernatant were diluted with 500 μl of de-ionized water and the samples were subjected to UPLC analysis. UPLC analysis was performed using a Waters H-Class Bio using a BEH C18 1.7 μM column and a flow rate of 0.6 ml min−1 (eluent A, 0.1% formic acid in water; eluent B, 0.1% formic acid in acetonitrile; gradient at 0 min: 90% A, 10% B; at 0.5 min: 90% A, 10% B; at 2.5 min: 10% A, 90% B; at 3.5 min: 90% A, 10% B; at 4.5 min: 90% A, 10% B). The ultraviolet signal at 210 nm was used for quantification, and concentrations of the metathesis product umbelliferone 2 (retention time of 1.38 min) were determined on the basis of a standard curve with commercially available umbelliferone (Sigma Aldrich). To quantify the ruthenium content of cells, cultures from 96-deepwell plates were pelleted by centrifugation (3,220 r.c.f., 12 min, 4 °C) and resuspended in ice-cold Tris/HCl-buffer (1 ml, 50 mM, pH 7.4, 0.9% (w/v) NaCl) containing biot-Ru (10 μM). After incubation on ice (30 min), the cells were spun down (1,260 r.c.f., 8 min, 4 °C), the supernatant-containing excess cofactor was discarded, and the pellet was washed by resuspension in ice-cold Tris/HCl-buffer (1 ml), centrifugation and careful removal of the supernatant. Afterwards, the pellet was resuspended in de-ionized water (500 μl). Twelve replicates of this suspension were combined and concentrated nitric acid was added (550 μl, 65%) to fully digest the cellular material (48 h, 110 °C, pressure vials). The resulting clear solutions were diluted to a final volume of 10 ml with de-ionized water (containing 1 p.p.m. yttrium as internal standard) and subjected to ICP-OES quantification. The obtained p.p.b.-values (1 p.p.b. = 1 μg l−1) for ruthenium were transformed into a molar concentration and the average ruthenium atom count per cell was calculated from the OD of the cultures before decomposition assuming an OD -to-cell-number correlation for E. coli in complex medium of 7.8 × 108 ml−1 OD −1, as described in ref. 34. The cellular metathesis activity was quantified using a fluorescent assay. For this purpose, cell cultures from 96-deepwell plate cultivations (see above) were pelleted by centrifugation (3,220 r.c.f., 12 min) and resuspended in ice-cold Tris/HCl-buffer (500 μl, 50 mM, pH 7.4) containing biot-Ru (2.1 μM). This buffer was supplemented with NaCl (0.9% (w/v)) to adjust to a physiological NaCl concentration. To allow cellular uptake of biot-Ru, the suspensions were incubated on ice for 30 min, spun down (1,260 r.c.f., 8 min) and the supernatant with its excess of cofactor was discarded. The pellets were resuspended in the reaction buffer (160 μl, 100 mM sodium acetate, pH 4.0, 0.5 M MgCl ) and the reaction was initiated by addition of these cell suspensions (150 μl) to the substrate solution (50 μl reaction buffer containing 40 mM precursor 1 and 20% DMSO), leading to a final substrate concentration of 10 mM and 5% DMSO. As the metathesis product umbelliferone 2 is fluorescent, the reaction progress was monitored by fluorescence in a microtiter plate reader (excitation wavelength, 322 ± 4.5 nm; emission wavelength, 440 ± 10 nm; Infinite M1000 PRO, Tecan Group AG) at 37 °C and agitation (6-mm amplitude, orbital). Cell-specific metathesis activity is specified as the slope of the increasing fluorescence signal in the linear range of the reaction normalized by the OD of the respective culture. To generate diversity in the scaffold protein SAV, a focused, semi-rational strategy was pursued: the 20 amino acid residues closest to the ruthenium ion (see Supplementary Table 4) in a related artificial metalloenzyme structure8 were selected and individually randomized in SAVperi by site-saturation mutagenesis using NNK codons30. Degeneration was introduced by application of the Quikchange Site Directed Mutagenesis Protocol (Stratagene) using degenerate oligos (see Supplementary Table 4) and, after transformation, the libraries were checked for diversity by Sanger-sequencing of at least four individual clones before screening. To evaluate the performance of the SAVperi variants, the aforementioned fluorescent metathesis assay was carried out with at least 90 clones from the library in one 96-well plate (three replicates of strains producing ‘wild-type’ SAV (parent) and three replicates of strains carrying an empty vector control (pET-30b(+ ), lacking the gene for SAV ). Evaluating 90 members of an NNK library ensures a >94% likelihood of screening all 20 amino acid residues in the respective position29. To compensate for biological variance, promising clones were isolated and subjected to a replicate assay that was identical to the protocol described above, but using eight independent cultures per clone. After this first screening round, promising residues were ordered according to their potential impact on catalysis and mutations were combined using iterative saturation mutagenesis (ISM)30. To perform kinetic experiments on the artificial metathase, the initially used T7-tagged SAV as well as the quintuple mutant SAVmut that was isolated after ISM were cloned into a cytoplasmic expression vector (see Methods section ‘Cloning of SAV expression constructs’) and purified on an iminobiotin sepharose column as described elsewhere35. The biotin binding capacity was determined using a fluorescent quenching assay36. Kinetic measurements were performed in reaction buffer (200 μl total volume, 100 mM sodium acetate, pH 4.0, 0.5 M MgCl , 11.5% (v/v) DMSO) containing biot-Ru (50 μM) both in the presence and absence of purified SAV (100 μM binding sites of either T7-tagged SAV or SAVmut) as well as the substrate 1 (variable concentrations, 0–5 mM) for fluorescent RCM. The reaction was monitored in a microtiter plate reader as described in Methods section ‘Fluorescent metathesis assay with whole cells’. The maximum velocity of the reaction was determined from the fluorescent signal curve by linear regression. To retrieve kinetic parameters, the reaction velocities were plotted over the respective substrate concentrations using the software GraphPad Prism (GraphPad Software, version 6.05) and applying the integrated ‘Michaelis–Menten’ least-squares fit with no constraints for the maximum velocity V and substrate affinity K . Crystals of SAV and variant SAVmut were obtained at 20 °C within two days by the sitting-drop vapour diffusion technique mixing 1 μl crystallization buffer (1.5 M ammonium sulfate, 0.1 M sodium acetate, pH 4.0) and 4 μl protein solution (26 mg ml−1 lyophilized protein in water). The droplet was equilibrated against a reservoir solution of 100 μl crystallization buffer. Subsequently, single crystals were soaked for two days at 20 °C in a soaking buffer, which was prepared by mixing 1 μl of a 10 mM stock solution of complex biot-Ru (in 50% aqueous DMSO) and 9 μl crystallization buffer. After the soaking, crystals were transferred for 30 s into a cryo-protectant solution consisting of 25% (v/v) glycerol in crystallization buffer. Next, crystals were shock-frozen in liquid nitrogen. Additional soaking of the above metathase crystals with substrate surrogate 1 did not lead to fluorescence. We therefore conclude that the multiple catalytic steps (for example, ligand displacement and cross-metathesis) required to ultimately liberate umbelliferone 2 cannot take place within a crystal. X-ray diffraction data were collected at the Swiss Light Source beam line X06DA at a wavelength of 1 Å and processed with the software XDS37 and AIMLESS (CCP4 Suite)38. The structure was solved by molecular replacement using the program PHASER (CCP4 Suite)38 and the structure 2QCB from the PDB as an input model with ligand and water molecules removed. For structure refinement REFMAC5 (CCP4 Suite)39 and PHENIX.REFINE40 were used. Ligand manipulation was carried out with the program REEL using the small-molecule crystal structure ABEJUM from the Cambridge Structural Database as an input model40. For water picking and electron density and structure visualization, the software COOT41 was used. Figures were drawn with PyMOL (the PyMOL Molecular Graphics System, version 188.8.131.52, Schrödinger, LLC). Crystallographic details, processing and refinement statistics are given in Supplementary Table 5. There is one SAVmut monomer in the asymmetric unit from which a tetramer can be generated by application of two orthogonal crystallographic two-fold symmetry axes. The 12 N-terminal residues of the T7-tag and 25 residues at the C terminus are not resolved, probably owing to disorder. Residual electron density in the biotin-binding site and the biotin vestibule as well as two strong anomalous dispersion density peaks in the biotin vestibule (Extended Data Fig. 3) suggested modelling of complex biot-Ru in two conformations I and II (56% and 44% occupancy, respectively). This projects the ruthenium atom in either one of the two densities, in close proximity to a crystallographic two-fold symmetry axis (Fig. 3b, Extended Data Figs 3, 4c, d). Only partial or no electron density was present for the mesityl linker and the terminal mesityl group, probably owing to high flexibility. In conformer I, the lengthy dimesitylimidazolidine (DMI)-Ru head group reaches into the neighbouring cis-related SAVmut monomer. The I conformer is stabilized mostly by hydrophobic interactions between the distal face of the DMI ligand and amino acid side chains within two neighbouring cis-related 7,8-loops (L1101, S1121, T114Q1, K121R1, L1241, S1122, W1202, K121R2, L1242; superscripts refer to monomers 1 or 2 of the tetrameric SAVmut; Extended Data Fig. 4c). Besides the DMI ligand, two chloride ions could be modelled binding to the ruthenium, but no density was found for the alkylidene, presumably owing to high flexibility and/or low occupancy. The orientation of the chlorides was very similar to that in a small-molecule crystal structure of the Grubbs–Hoyveda second-generation catalyst (ABEJUM from the Cambridge Structural Database), placing them nearly in trans position to each other. The ruthenium is largely solvent-exposed, which could facilitate substrate binding and product release. In Fig. 3b, the alkylidene was modelled binding to the ruthenium (magenta stick model) to highlight its orientation in the biotin vestibule. Conformer II is different from I by a rotation (about 60°) of the DMI-Ru moiety around an axis parallel to the cylinder axis of the SAVmut β-barrel (Fig. 3b, Extended Data Fig. 4d). This rotation places the hydrophobic distal side of the DMI-ligand in proximity to amino acid side chains within loop-5,61 (A86, H87), loop-7,81 (S112, T114Q) and loop-4,53 (D67, S69, A65) of a neighbouring trans-related SAV monomer. Atom N49K-Nε is located in close proximity to ruthenium (2.4 Å). No electron density was found to model chloride ions and the alkylidene ligand bound to the ruthenium. Because the cofactor is bound in close proximity to a two-fold crystal symmetry axis, formation of a cis-symmetry-related neighbouring cofactor by application of the crystal symmetry operation results in extensive steric clashes between the two cofactors in the orientation I. In contrast, coexistence of cofactor pairs I–I or II–II orientation is sterically accessible (Extended Data Fig. 3a, c, e). Normalized B factors of residues within cofactor-flanking loop-7,8 are increased by up to about 0.5 when compared to related SAV structures that crystalized in the same space group and in a very similar unit cell (Extended Data Fig. 5b). This suggests increased loop-7,8 flexibility (Fig. 3b, Extended Data Fig. 4c, d). This flexibility is likely to be caused by three factors: (i) mutation T114Q cleaves a hydrogen-bond between threonine-OγH and T115-carbonyl oxygen (green dashed line in Fig. 3b and Extended Data Fig. 4a, b) and leads to a new hydrogen-bond between T114Q-glutamine-Nε and S112-OγH (red dashed line in Fig. 3b and Extended Data Fig. 4c, d); (ii) mutation A119G in the loop leads to increased entropy; and (iii) mutation V47A reduces steric hindrance between V47A in loop-3,42 and W120 in loop-7,81 (Fig. 3b and Extended Data Fig. 4c, d). The overall structure of complex biot-Ru–SAV is virtually identical to that of complex biot-Ru–SAVmut (root-mean-square deviation, r.m.s.d. = 0.25 Å). As in the mutant, two strong residual electron density peaks (F − F cofactor omit map: 12σ (conformer I) and 11σ (conformer II)) and two anomalous dispersion density peaks (9σ (conformer I) and 5σ (conformer II)) were located within the biotin vestibule of a SAV monomer at the interface between two symmetry-related SAV monomers (Extended Data Fig. 3b, d, f). The same two cofactor conformations I and II found in mutant SAVmut were modelled in the biotin binding vestibule of SAV with an occupancy of 50% for each conformer I and II (Extended Data Fig. 4a, b). The side chain of residue L110 adopts two conformations with 50% occupancy each (Extended Data Fig. 4a, b). The close proximity of the terminal methyl group in L110 conformation A to the aromatic mesityl ring (distance between L110-CδH and mesityl of 4.4 Å) of cofactor conformation I suggests a stabilizing σ–π interactions (Extended Data Fig. 4a, red star). The L110 side chain conformation B can coexist only with cofactor conformation II (Extended Data Fig. 4b). This hypothesis is supported by the fact that the same L110 side chain conformation A is found in complex [(Cp*)Ir(Biot-p-L)Cl]-SAV-S112A (PDB, 3PK2), which has an aromatic ring located in the same position as the mesityl linker in structure biot-Ru–SAV, suggesting a similar σ–π interaction. In contrast, the side chain of L110 in apo-SAV (PDB, 2BC3) adopts conformation B. In complex biot-Ru–SAV, a water molecule is bound in proximity to L110 with 50% occupancy (Extended Data Fig. 4b). Steric clashes with L110 side chain in conformation A and the NHC ligand of biot-Ru conformer I suggest that the water is present only with L110 conformation B and biot-Ru conformer II. Additionally, the side chain of L124 adopts two conformations, each with 50% occupancy. Only conformation L124A does not undergo steric clashes with a methyl group of the bridging mesityl moiety of cofactor conformer I (Extended Data Fig. 4a, b). Together, the conformational side chain flexibility of residues L110 and L124 reflects the presence of the two cofactor conformations I and II. In contrast to artificial metalloenzyme biot-Ru–SAVmut, the normalized B factors of residues within the cofactor-flanking loop-7,8 in complex biot-Ru–SAV do not show increased values when compared to those in related crystal structures (Extended Data Fig. 5b). Indeed, a hydrogen-bond is formed between T114-OγH and T115-carbonyl oxygen in complex biot-Ru–SAV that could rigidify the loop (Extended Data Fig. 4a, b). The conversion for the product 4 was quantified by 1H-NMR. For this purpose, a deuterated reaction buffer was prepared from acetic acid-d , dry MgCl and D O with the same concentrations as for the reaction buffer used for substrate 1 (100 mM acetate, 0.5 M MgCl ). The pH was adjusted to 3.6 by addition of 1 M NaOD in D O (with respect to pD = pH + 0.4). For the reaction, 300 μl of a substrate 3 stock solution (100 mM in deuterated reaction buffer) was mixed with 291 μl of either a solution of SAV, SAVmut, or SAVmut2 (200 μM binding sites in deuterated reaction buffer) or plain deuterated reaction buffer (for samples without SAV or any of the SAV variants). Afterwards, 9 μl of a biot-Ru (or HGII/AQM) stock solution (3.34 mM in DMSO-d ) was added to obtain a final concentration of 50 μM and the reaction was performed for 16 h at 37 °C and 200 r.p.m. The mixture was analysed by 1H-NMR and the yield of the reaction product 4 was quantified by comparing integrals (I) of the product 4 peaks at 3.41 p.p.m. and 2.05 p.p.m. and the substrate 3 peaks at 3.33 p.p.m. and 1.91 p.p.m. using: yield = I /(I + I ). To quantify the conversion of substrate 5, a 97 μl aliquot of either SAV solution (200 μM SAV binding sites in reaction buffer) or plain reaction buffer (for samples without SAV variants) was mixed with 100 μl of a stock solution of substrate 5 (20 mM in reaction buffer). Subsequently, 3 μl of the respective catalyst/cofactor stock solution (3.34 mM in DMSO) was added to obtain a final concentration of 50 μM. The reaction was performed for 16 h at 37 °C and 200 r.p.m. Then, an aqueous solution of benzyltriethylammonium chloride (100 μl, 10 mM) was added as an internal standard and 700 μl of methanol was added. The mixture was cleared by centrifugation and 250 μl of the supernatant was mixed with 750 μl of water for the final quantification of product 6 by UPLC-MS. For the kinetic experiment, the reaction mixture was scaled up to a total volume of 1 ml and 50 μl aliquots of this mixture were collected at different time points and immediately injected into 950 μl of a quenching solution (0.5 mM potassium cyanoacetate, 0.25 mM benzyltriethyl-ammonium chloride (internal standard) in 50% aqueous methanol). After removal of precipitated protein by centrifugation, the supernatant was analysed by UPLC-MS. To quantify the cellular metathesis activity for substrate 5, a protocol analogous to that applied for the umbelliferone precursor 1 was applied. The substrate 5 (final concentration 10 mM) was added to whole cells and the samples were incubated at 37 °C and 300 r.p.m. for 16 h. To quantify the conversion for the non-fluorescent product 6, an extraction was performed: 800 μl of methanol was added to each sample and an extraction was carried out (one hour with vigorous shaking, 800 r.p.m. at room temperature). The samples were cleared by centrifugation, the supernatant was diluted with water (factor four) and analysed by UPLC-MS using a calibration curve recorded for product 6. No statistical methods were used to predetermine sample size.
News Article | April 15, 2016
Abstract: EPFL scientists have built a single-atom magnet that is the most stable to-date. The breakthrough paves the way for the scalable production of miniature magnetic storage devices. Magnetic storage devices such as computer hard drives or memory cards are widespread today. But as computer technology grows smaller, there is a need to also miniaturize data storage. This is epitomized by an effort to build magnets the size of a single atom. However, a magnet that small is very hard to keep "magnetized", which means that it would be unable to retain information for a meaningful amount time. In a breakthrough study published in Science, researchers led by EPFL have now built a single-atom magnet that, although working at around 40 Kelvin (-233.15 oC), is the smallest and most stable to date. Magnets work because of electron spin, which is a complicated motion best imagined as a spinning top. Electrons can spin up or down (something like clockwise or anti-clockwise), which creates a tiny magnetic field. In an atom, electrons usually come in pairs with opposite spins, thus cancelling out each other's magnetic field. But in a magnet, atoms have unpaired electrons, and their spins create an overall magnetic field. A challenge today is to build smaller and smaller magnets that can be implemented in data storage devices. The problem is something called "magnetic remanence", which describes the ability of a magnet to remain magnetized. Remanence is very difficult to observe from a single atom, because environmental fluctuations can flip its magnetic fields. In terms of technology, a limited remanence would mean limited information storage for atom-sized magnets. A team of scientists led by Harald Brune at EPFL and his colleagues at ETH Zurich, have built a prototypical single-atom magnet based on atoms of the rare-earth element holmium. The researchers, placed single holmium atoms on ultrathin films of magnesium oxide, which were previously grown on a surface of silver. This method allows the formation of single-atom magnets with robust remanence. The reason is that the electron structure of holmium atoms protects the magnetic field from being flipped. The magnetic remanence of the holmium atoms is stable at temperatures around 40 Kelvin (-233.15 oC), which, though far from room temperature, are the highest achieved ever. The scientists' calculations demonstrate that the remanence of single holmium atoms at these temperatures is much higher than the remanence seen in previous magnets, which were also made up of 3-12 atoms. This makes the new single-atom magnet a worldwide record in terms of both size and stability. ### This project involved a collaboration of EPFL's Institute of Condensed Matter Physics with ETH Zurich, Swiss Light Source (PSI), Vinča Institute of Nuclear Sciences (Belgrade), the Texas A&M University at Qatar and the European Synchrotron Radiation Facility (Grenoble). It was funded by the Swiss National Science Foundation, the Swiss Competence Centre for Materials Science and Technology (CCMX), the ETH Zurich, EPFL and the Marie Curie Institute, and the Serbian Ministry of Education and Science. For more information, please click If you have a comment, please us. Issuers of news releases, not 7th Wave, Inc. or Nanotechnology Now, are solely responsible for the accuracy of the content.
News Article | February 15, 2017
The new method offers unprecedented detail in measuring molecular motion and energy – enabling better control and understanding of chemical reactions in the field of biotechnology research. The technique, which was recently published in Nature Communications, uses X-ray scattering to measure the specific movements of atoms in a molecule with extreme energy resolution. "The idea is based on exciting a molecule to a high-energy, localized state," says Victor Kimberg, a researcher in the department of Theoretical Chemistry and Biology, at KTH Royal Institute of Technology in Stockholm. The X-ray radiation that it emits then scans the energy landscape of the molecule with a level of precision that Kimberg compares to observing the movements of individual insects from on top of a mountain. Every molecule has its own energy "landscape", or the full multidimensional spectrum of motion that atoms undergo when the molecule is energized. These motions include bending and stretching of bonds. Expressed in geometric terms, the relationships of atoms to one another in a single molecule are among the key things scientists need to know in order to determine a molecule's potential energy surface (PES) – an important value in the study of molecular structures, properties and reactivity. "The PES is useful for processes such as catalysis and photochemistry," Kimberg says. For the first time, the technique enables scientists to break down the collective atomic motion in a molecule into elementary components, he says. "We can now go further than examining the collective multidimensional atomic motion in a molecule, and look at specific vibrations along selected reaction coordinates," he says. "We are not aware of any other way to do this, so it looks like our idea is new." Typically when a molecule is excited, the only measurements available show all atomic motion simultaneously. The full PES landscape is complex with all types of vibrations. Kimberg says that with the method they propose, x-ray photon energy can be tuned to excite vibrations of a singled-out type of nuclear motion – which he says can serve as a basis for developing methods of reaction control. "We show clearly that tuning the x-ray photon energy in resonance with one core-excited state induces only symmetric stretching motion; while tuning to another core-excited state excites exclusively the bending motion," he says. The measurements were conducted at the Swiss Light Source (SLS) synchrotron laboratory in Zurich, in collaboration with the KTH group, comprised of Kimberg, Faris Gel'mukhanov and Hans Ågren, who were responsible for the underlying theory and simulations. Explore further: High-energy electrons synced to ultrafast laser pulse to probe how vibrational states of atoms change in time More information: Rafael C. Couto et al. Selective gating to vibrational modes through resonant X-ray scattering, Nature Communications (2017). DOI: 10.1038/ncomms14165
News Article | March 31, 2016
Scientists from the University of Copenhagen and the Paul Scherrer Institute in Switzerland have, for the first time, created a 3D image of food on the nanometer scale. The method the scientists used is called Ptychographic X-ray computed tomography. It has promising prospects as a more detailed knowledge of the structure of complex food systems could potentially save the food industry large sums of money and reduce food waste that occurs because of faulty production. The researchers found that 98 percent of the fat globules in the cream cheese-like food system are cemented together in a continuous 3D network. They have visualized the network using the same techniques that are used in computer animation. The coherent network of fat globules (approximately 25 percent of the volume) is seen in the video as the cohesive yellow structure, while the coherent structure of water fills out the area between the fat and is not highlighted by any color for clarity. The small areas of fat or water, which are not connected to the remaining fat or water structures, are red and blue respectively. The grey areas are the food ingredient microcrystalline cellulose. The network shown in the video is about 20 microns in diameter and made of fat globules of about 1 micron in size. "There is still a lot we don’t know about the structure of food, but this is a good step on the way to understanding and finding solutions to a number of problems dealing with food consistency, and which cost the food industry a lot of money," says Associate Professor Jens Risbo from the Department of Food Science at the University of Copenhagen. He is one of the authors of a recently-published scientific paper in Food Structure, which deals with the new groundbreaking insight into the 3D structure of food. The researchers used a cream based on vegetable fat for the research. The cream system is a good test material, since it can represent the structures of a large group of food systems, for example cheese, yogurt, ice cream, spreads, but also the more solid chocolate. All the aforementioned products contain liquid water or fat as well as small particles of solid materials, which stick together and form three-dimensional structures — i.e. a network that provides the consistency that we like about cheese, yogurt, or chocolate. In cheese and yogurt, the casein particles form the network. In chocolate it is the fat crystals and in ice cream and whipped cream it is the fat globules. "It's about understanding the food structure and texture. If you understand the structure, you can change it and obtain exactly the texture you want," says Risbo. To create a three-dimensional model of the food and convert it into images and video, the scientists have been in Switzerland, where they have used the Swiss Light Source (SLS) synchrotron at the Paul Scherrer Institute. In the synchrotron electrons are accelerated to near speed of light. The synchrotron is used for research in materials science in areas such as biology and chemistry. The method the researchers used is called "Ptychographic X-ray computed tomography." This is a groundbreaking new method for creating images on the nanometer scale, which also provides a high contrast in biological systems. The synchrotron in Switzerland is one of the leading places in the world in this area, and it was the first time ever that it was used within food science. "We have been using the tomography principle, also known from an X-ray CT (computed tomography) scanner. The sample of the food system is rotated and moved sideways back and forth with nanometer precision, while we send a very strong and focused X-ray beam through it. The X-rays are deflected by colliding with electrons in the food, and we shoot a lot of pictures of the patterns that the defleted X-rays form. The patterns are combined in a powerful computer, which reconstructs a 3D image of the sample. The Swiss scientists of the team have created a device that can move and rotate the sample with ultra-high precision, allowing us to see the small details," says Research Assistant Mikkel Schou Nielsen, who has recently completed his Ph.D. in tomographic methods applied to food at the Niels Bohr Institute in Copenhagen. The reconstructed 3D image can be described as a three-dimensional table of numbers describing the electron density (the number of electrons per volume) through the entire sample. The various food components, such as water and fat, have different densities and hence different electron density. Water is heavier than fat, which is known from oil that settles on top of water when you try to mix them, and it is this contrast in electron density which causes X-rays to deflect to different degrees and eventually to form 3D-images of the sample. Water thus appears light grey, while fat appears dark grey, and the glass around the sample with a high density is seen as a white ring. One may now use the electron density (greyscale) to identify the various food components and study their location and structure. The vegetable-based cream which the method is used on consists of several ingredients. In addition to water and vegetable fat, it contains milk protein, stabilizers and emulsifiers. By adjusting the addition of emulsifiers, it is possible to achieve a state in which the cream continues to be fluid until you whip it to foam, whereby all the fat globules are reorganized and sticking together on the outside of the air bubbles in a three-dimensional system. "It is a difficult balance, because you only want the fat globules to stick together when the cream is whipped — not if it is simply being exposed to vibration or high temperatures. When the fat globules nevertheless begin to stick together prematurely - for example due to too many shocks during transport — the cream will get a consistency reminiscent of cream cheese. It becomes a relatively hard lump that can be cut," says Postdoctoral Researcher Merete Bøgelund Munk, Department of Food Science, University of Copenhagen. Munk's Ph.D. project, “The physical stability of whippable oil-in-water emulsions. Effects of monoglyceride-based emulsifiers and other ingredients,” was fundamental for the research. The Ph.D. project is made as a collaboration between the Department of Food Science at the University of Copenhagen and the food ingredient company Palsgaard A/S. This undesirable cream cheese-like state of the vegetable cream system is nevertheless extremely interesting for researchers. "The organization of the fat globules and the network structure after the cream has been converted into a ‘cream cheese-like’ product is exciting because the mass is now sliceable, even though the system consists of 65 percent water and only 25 percent fat and some other ingredients and sugars. That means we have a network structure that captures a lot of water. There are many foods with similar network systems of something solid in something liquid, where the liquid is typically, but not always, water. This applies to all semi-solid and solid products such as chocolate, butter, cheese and spreads. The network of the cream cheese-like system is thus a model for something general in our food," says Risbo. It is the structure of the networks which forms a texture that makes you want to bite into a piece of chocolate and cut yourself a piece of cheese. But the structure and the networks are something of a mystery, because until now you could only see the surface or only slightly underneath the surface of the food material on the microns scale and the images you could see have only been two-dimensional. "If we eventually come to understand the structure of chocolate, we can change it and obtain exactly the consistency that we want. A lot of money is wasted because the consistency of chocolate is really hard to control, so the end product is not good enough and must be discarded. A possible future understanding of the crystal network in chocolate might mean that we will be able to develop components that prevent the chocolate from becoming grey and crumbly, and thus unsaleable. It is certainly a possibility that tomographic methods could be developed so we would be able to understand the mysteries of chocolate," says Risbo. "Ptychographic X-ray computed tomography can be compared with a CT scanner in a hospital. Instead of getting an image of a patient's organs, we are looking into food. But, unlike a CT scanner, we can go down to the nanometer scale," says Risbo. The sample with the cream cheese-like system that the scientists X-rayed was about 20 microns thick. "It would take too much time and too many calculations to develop a nanometer resolution of the cream system for a whole package of cream cheese from the fridge. The amount of information and calculations would simply be too great. Although X-rays can almost go through everything, you lose the intensity of the beams, the more they have to shoot through," says Risbo.
News Article | March 31, 2016
Home > Press > For the first time scientists can observe the nano structure of food in 3-D Abstract: Scientists have, for the first time, created a 3-D image of food on the nanometer scale. It has promising prospects as a more detailed knowledge of the structure of complex food systems could potentially save the food industry large sums of money. Computer animation/video: Liborius ApS (0:33) "There is still a lot we don't know about the structure of food, but this is a good step on the way to understanding and finding solutions to a number of problems dealing with food consistency, and which cost the food industry a lot of money," says Associate Professor Jens Risbo, Department of Food Science at the University of Copenhagen, Denmark. He is one of the authors of a recently-published scientific paper in Food Structure, which deals with the new groundbreaking insight into the 3D structure of food. The researchers used a cream based on vegetable fat for the research. The cream system is a good test material, since it can represent the structures of a large group of food systems, for example cheese, yogurt, ice cream, spreads, but also the more solid chocolate. All the aforementioned products contain liquid water or fat as well as small particles of solid materials, which stick together and form three-dimensional structures - i.e. a network that provides the consistency that we like about cheese, yogurt or chocolate. In cheese and yoghurt the casein particles form the network. In chocolate it is the fat crystals and in ice cream and whipped cream it is the fat globules. "If you understand the structure, you can change it and obtain exactly the texture you want," says Jens Risbo. Electrons with close to speed of light generate intense X-rays To create a three-dimensional model of the food and convert it into images and video, the scientists have been in Switzerland, where they have used the Swiss Light Source (SLS) synchrotron at the Paul Scherrer Institute. In the synchrotron electrons are accelerated to near speed of light. The synchrotron is used for research in materials science in areas such as biology and chemistry. The method the researchers used is called "Ptychographic X-ray computed tomography." This is a new method for creating images on the nanometer scale, which also provides a high contrast in biological systems. The synchrotron in Switzerland is one of the leading places in the world in this area, and it was the first time ever that it was used within food science. "We have been using the tomography principle, also known from an X-ray CT (computed tomography) scanner. The sample of the food system is rotated and moved sideways back and forth with nanometer precision, while we send a very strong and focused X-ray beam through it. The X-rays are deflected by colliding with electrons in the food, and we shoot a lot of pictures of the patterns that the defleted X-rays form. The patterns are combined in a powerful computer, which reconstructs a 3D image of the sample. The Swiss scientists of the team have created a device that can move and rotate the sample with ultra-high precision, allowing us to see the small details," says Research Assistant Mikkel Schou Nielsen, who has recently completed his Ph.D. in tomographic methods applied to food at the Niels Bohr Institute in Copenhagen. The number of electrons reveals the various food components The reconstructed 3D image can be described as a three-dimensional table of numbers describing the electron density (the number of electrons per volume) through the entire sample. The various food components, such as water and fat, have different densities and hence different electron density. Water is heavier than fat, which is known from oil that settles on top of water when you try to mix them, and it is this contrast in electron density which causes X-rays to deflect to different degrees and eventually to form 3D-images of the sample. Figure 1 shows a 2D slice of the three dimensional structure. Areas with higher electron density appear lighter on the figure. Water thus appears light grey, while fat appears dark grey, and the glass around the sample with a high density is seen as a white ring. One may now use the electron density (greyscale) to identify the various food components and study their location and structure. A complicated food system The vegetable-based cream which the method is used on consists of several ingredients. In addition to water and vegetable fat, it contains milk protein, stabilizers and emulsifiers. By adjusting the addition of emulsifiers, it is possible to achieve a state in which the cream continues to be fluid until you whip it to foam, whereby all the fat globules are reorganized and sticking together on the outside of the air bubbles in a three-dimensional system (see Figure 2). "It is a difficult balance, because you only want the fat globules to stick together when the cream is whipped - not if it is simply being exposed to vibration or high temperatures. When the fat globules nevertheless begin to stick together prematurely - for example due to too many shocks during transport - the cream will get a consistency reminiscent of cream cheese. It becomes a relatively hard lump that can be cut," says Postdoctoral Researcher Merete Boegelund Munk, Department of Food Science, University of Copenhagen. Merete Boegelund Munk's Ph.D. project, "The physical stability of whippable oil-in-water emulsions. Effects of monoglyceride-based emulsifiers and other ingredients", was fundamental for the research. The Ph.D. project was made as a collaboration between the Department of Food Science at the University of Copenhagen and the food ingredient company Palsgaard A / S. This undesirable cream cheese-like state of the vegetable cream system is nevertheless extremely interesting for researchers. "The organization of the fat globules and the network structure after the cream has been converted into a 'cream cheese-like' product is exciting because the mass is now sliceable, even though the system consists of 65% water and only 25% fat and some other ingredients and sugars. That means we have a network structure that captures a lot of water. There are many foods with similar network systems of something solid in something liquid, where the liquid is typically, but not always, water. This applies to all semi-solid and solid products such as chocolate, butter, cheese and spreads. The network of the cream cheese-like system is thus a model for something general in our food," says Associate Professor Jens Risbo, Department of Food Science, University of Copenhagen. It is the structure of the networks which forms a texture that makes you want to bite into a piece of chocolate and cut yourself a piece of cheese. But the structure and the networks are something of a mystery, because until now you could only see the surface and slightly underneath the surface of the food material on the microns scale and the images you could see have only been two-dimensional. "If we eventually come to understand the structure of chocolate, we can change it and obtain exactly the consistency that we want. A lot of money is wasted because the consistency of chocolate is really hard to control, so the end product is not good enough and must be discarded. A possible future understanding of the crystal network in chocolate might mean that we will be able to develop components that prevent the chocolate from becoming grey and crumbly, and thus unsaleable. It is certainly a possibility that tomographic methods could be developed so we would be able to understand the mysteries of chocolate," says Associate Professor, Jens Risbo, Department of Food Science, University of Copenhagen. How the tomography works "Ptychographic X-ray computed tomography can be compared with a CT scanner in a hospital. Instead of getting an image of a patient's organs, we are looking into food. But, unlike a CT scanner, we can go down to the nanometer scale," says Jens Risbo. The sample with the cream cheese-like system that the scientists X-rayed was about 20 microns thick. "It would take too much time and too many calculations to develop a nanometer resolution of the cream system for a whole package of cream cheese from the fridge. The amount of information and calculations would simply be too great. Although X-rays can almost go through everything, you lose the intensity of the beams, the more they have to shoot through," says Jens Risbo. Swiss Light Source synchrotron Basically, you can make X-rays in two different ways. If you go to the dentist and have an X-ray done, this is done using an X-ray tube, which can be compared with a cathode ray tube showing the pictures in an old type of television, where electrons are not accelerated to very high speeds. In the X-ray tube the electrons collide with a metal, such as copper, which now emits X-rays. The X-ray tube is not so powerful, but you can make medical photos and also do some research work with this type of X-rays. But if you want to examine very small samples, things that are changing rapidly, or make tomography at the nanometer scale, you will use facilities like Swiss Light Source or the Swedish synchrotron MAX IV which opens in Lund, Sweden, this year. "Technically, it is electrons that are accelerated to nearly the speed of light and circulates in a ring controlled by electromagnets. The electron beams are then deflected and will then emit intense and energetic X-rays," says Associate Professor Jens Risbo, Department of Food Science, University of Copenhagen. The Swiss Light Source, SLS, is funded by the Swiss government and scientists from around the world can apply to use syncroton X-rays and related scientific equipment under the guidance of local scientists. For more information, please click If you have a comment, please us. Issuers of news releases, not 7th Wave, Inc. or Nanotechnology Now, are solely responsible for the accuracy of the content.
News Article | February 9, 2017
EPFL (École polytechnique fédérale de Lausanne) scientists have been able to measure the ultrashort time delay in electron photoemission without using a clock. The discovery has important implications for fundamental research and cutting-edge technology. When light shines on certain materials, it causes them to emit electrons. This is called “photoemission” and it was discovered by Albert Einstein in 1905, winning him the Nobel Prize. But only in the last few years, with advancements in laser technology, have scientists been able to approach the incredibly short timescales of photoemission. Researchers at EPFL have now determined a delay of one billionth of one billionth of a second in photoemission by measuring the spin of photoemitted electrons without the need of ultrashort laser pulses. The discovery is published in Physical Review Letters. Photoemission has proven to be an important phenomenon, forming a platform for cutting-edge spectroscopy techniques that allow scientists to study the properties of electrons in a solid. One such property is spin, an intrinsic quantum property of particles that makes them look like as if they were rotating around their axis. The degree to which this axis is aligned towards a particular direction is referred to as spin polarization, which is what gives some materials, like iron, magnetic properties. Although there has been great progress in using photoemission and spin polarization of photo-emitted electrons, the time scale in which this entire process takes places have not been explored in great detail. The common assumption is that, once light reaches the material, electrons are instantaneously excited and emitted. But more recent studies using advanced laser technology have challenged this, showing that there is actually a time delay on the scale of attoseconds. The lab of Hugo Dil at EPFL, with colleagues in Germany, showed that during photoemission, the spin polarization of emitted electrons can be related to the attosecond time delays of photoemission. More importantly, they have shown this without the need for any experimental time resolution or measurement — essentially, without the need for a clock. To do this, the scientists used a type of photoemission spectroscopy (SARPES) to measure the spin of electrons photo-emitted from a crystal of copper. “With lasers you can directly measure the time delay between different processes, but it is difficult to determine when a process starts — time zero,” says Mauro Fanciulli, a PhD student of Dil’s group and first author on the paper. “But in our experiment we measure time indirectly, so we don’t have that problem — we could access one of the shortest timescales ever measured. The two techniques [spin and lasers], are complementary, and together they can yield a whole new realm of information.” The information about the timescale of photoemission is included in the wavefunction of the emitted electrons. This is a quantum description of the probability of where any given electron can be found at any given time. By using SAPRES, the scientists were able to measure the spin of the electrons, which in turn allowed them to access their wavefunction properties. “The work is a proof of principle that can trigger further fundamental and applied research,” says Dil. “It deals with the fundamental nature of time itself and will help understand the details of the photoemission process, but it can also be used in photoemission spectroscopy on materials of interest.” Some of these materials include graphene and high-temperature superconductors, which Dil and his colleagues will be studying next. This work involved a collaboration between EPFL Institute of Physics, the Paul Scherrer Institut (Swiss Light Source), the Ludwig Maximillian University, the University of West Bohemia, and the University of Bielefeld. It was funded by the Swiss National Science Foundation (SNSF), the Bundesministerium für Bildung und Forschung (BMBF), the Deutsche Forschunsgsgemeinschaft (DFG), and the Ministry of Education of the Czech Republic.
News Article | March 23, 2016
No statistical methods were used to predetermine sample size. The experiments were not randomized. The investigators were not blinded to allocation during experiments and outcome assessment. M. smegmatis (ATCC 700084) was cultured according to ATCC recommendations. Cells were pelleted and washed with TE buffer. A cell pellet of gammaproteobacterium HdN1 was provided by J. Zedelius. Cells were resuspended in lysis buffer (0.1 M Tris pH 8.0, 0.2 M NaCl, 5 mM EDTA, 0.2 mg ml−1 lysozyme), incubated for 6 h at 37 °C, subsequently supplemented with 0.5% SDS and 0.2 mg ml−1 proteinase K, and incubated at 65 °C for 24 h. DNA was purified by phenol–chloroform extraction and dissolved in TE buffer. The MAS KS–AT constructs (Uniprot: A0R1E8, aa 1–884, 1–887, 1–892), MAS DH (A0R1E8, 884–1186), MAS DH–ΨKR–ER–KR (A0R1E8, 884-2020), and ‘Pks’ DH–ΨKR–ER–KR (Q3L885, 2450–3580) were cloned into pNIC28a-Bsa vectors; GpEryA DH–ΨKR–ER–KR (E1VID6, 2420–3575) constructs were cloned into a Gateway-compatible pETG-10A destination vector (provided by EMBL Heidelberg). MAS DH–ΨKR–ER–KR–ACP (A0R1E8, 884–2111) was cloned by codon-optimized gene synthesis of ACP (GenScript) and restriction cloning (BsrGI/HindIII) into pNIC28a-Bsa-MAS DH–ΨKR–ER–KR (884-2020). All constructs were designed as N-terminal tobacco etch virus (TEV) protease cleavable hexa-histidine (His6) fusion constructs and co-expressed with Streptomyces chaperonins31 (pETcoco-2A-L1SL2 plasmid) in BL21(DE3) pRIL cells or for GpEryA in Rosetta(DE3) pLysS cells with a kanamycin-resistant version of pETcoco-2A-L1SL2. Cells were cultured in 2×YT media, supplemented with 0.5% glycerol, NPS (25 mM (NH4) SO , 50 mM KH PO , 50 mM Na HPO ), kanamycin (100 μg ml−1), chloramphenicol (34 μg ml−1), and ampicillin (100 μg ml−1). An expression culture (1.5 l) was inoculated (1:20), grown at 37 °C for 2 h, cooled to 20 °C, and induced with isopropyl-β-d-thiogalactopyranosid (0.1 mM) at an absorbance at 600 nm of 1.0. Cells were collected after 12 h by centrifugation (7,000g) and resuspended in lysis buffer (50 mM HEPES pH 7.4, 20 mM imidazole, 0.5 M NaCl, 5 mM MgCl , 10% glycerol (v/v), 2.5 mM β-mercaptoethanol), supplemented with protease inhibitors (200 μM phenylmethylsulfonyl fluoride, 20 μM bestatin, 4 μM E64, 2 μM pepstatin A, 20 μM phenantrolin, 2 μM phosphoramidon) as well as DNase, RNase, and lysozyme. Cells were placed on ice and lysed by sonication. The lysate was cleared by centrifugation (100,000g, 30 min) and the supernatant was loaded onto a 5 ml Ni-affinity column (GenScript) pre-equilibrated with lysis buffer. Unbound protein was eluted with four alternating wash cycles of five column volumes (CV) lysis buffer and HisA buffer (50 mM HEPES pH 7.4, 20 mM imidazole, 50 mM NaCl, 5 mM MgCl , 10% glycerol (v/v), 2.5 mM β-mercaptoethanol, inhibitors), until a stable baseline (A280) was reached. The sample was eluted with 2 CV HisB buffer (50 mM HEPES pH 7.4, 250 mM imidazole, 50 mM NaCl, 10% glycerol (v/v), 2.5 mM β-mercapto ethanol, inhibitors) and diluted (1:10) with AIC-A buffer (50 mM Tris-HCl pH 7.4, 20 mM KCl, 10% (v/v) glycerol, 2.5 mM β-mercaptoethanol). The sample was loaded on a 6.5 ml anion exchange column (PL-SAX 4,000 Å, 10 μm) and washed with 20 CV. The samples were eluted with a stepped gradient to 100% AIC-B (50 mM Tris-HCl pH 7.4, 1 M NaCl, 10% (v/v) glycerol, 2.5 mM β-mercaptoethanol). For DH–ΨKR–ER–KR the gradient was held at a conductivity of 15 mS cm−1 until a stable baseline (A280) was obtained to elute Streptomyces chaperonins. DH–ΨKR–ER–KR eluted at 17–20 mS cm−1. Pure fractions were pooled, supplemented with TEV protease (1 mg protease per 100 mg tagged protein), and incubated for 10 h at 4 °C. Uncleaved protein, as well as the cleaved His6-tag, was removed by passing the solution through a 5 ml orthogonal Ni-affinity column (GenScript). The flow-through was pooled, concentrated, and subjected to gel permeation chromatography (Superdex 200 16/60, GE Healthcare) using GPC buffer (20 mM HEPES pH 7.4, 250 mM NaCl, 5% glycerol (v/v), 5 mM dithiothreitol). Pure fractions were pooled, and monodispersity was monitored by dynamic light scattering at 1 mg ml−1. Related purification protocols were applied to MAS DH–ΨKR–ER–KR–ACP, ‘Pks’ DH–ΨKR–ER–KR (HiTrap CaptoQ column), GpEryA DH–ΨKR–ER–KR (both no TEV protease cleavage and orthogonal Ni-affinity column), and MAS DH (no anion exchange chromatography). All crystallization experiments were performed using a robotic setup applying the sitting drop vapour diffusion method. MAS KS–AT bi-pyramidal crystals were grown at 4 °C by mixing 0.2 μl of protein in GPC buffer (38 mg ml−1) with 0.2 μl reservoir solution (0.1 M MES/imidazole pH 6.5, 0.1 M MgCl , 0.1 M CaCl , 12.5% (v/v) polyethylene glycol 1000 (v/v), 7.5% (w/v) polyethylene glycol (PEG) 3350, 12.5% 2-methyl-2,4-pentanediol (MPD)). Crystals grew to a final size of 0.8 mm × 0.4 mm × 0.2 mm within 1 week and were flash-frozen in liquid nitrogen. The MAS DH domain was crystallized in space group P2 at 18 °C by mixing 0.2 μl of protein in GPC buffer (38 mg ml−1) with 0.1 μl reservoir solution (0.1 M bis-Tris pH 6.5, 0.2 M MgCl , 25% (v/v) PEG 3350) and grew to a final size of 0.4 mm × 0.2 mm × 0.1 mm within one week. Crystals in space group P2 2 2 appeared after 30 days at 18 °C by mixing 1 μl of protein in GPC buffer (38 mg ml−1) with 2 μl reservoir solution (0.25 M di-sodium malonate, 24% (w/v) PEG 3350) and grew to a final size of 1 mm × 0.4 mm × 0.3 mm. Before harvesting, all crystals of MAS DH were cryoprotected (25% (v/v) ethylene glycol) and flash-frozen in liquid nitrogen. Needle-shaped crystals of MAS DH–ΨKR–ER–KR were obtained by mixing protein solution at 18.4 mg ml−1 (GPC buffer, 1.5 mM NADP+) and reservoir solution (0.03 M MgCl , 0.03 M CaCl , 20% ethylene glycol, 10% PEG 8000, 0.1 M MES/imidazole pH 6.5) at 4 °C. Crystallization was optimized by exchanging PEG 8000 by PEG 3350, decreasing the PEG 3350 concentration to 7–13% (w/v), and by carefully monitored microseeding. Subsequent optimization was performed using automated robotic setup and seeding at 4 °C. Final crystals (1.0 mm × 0.3 mm × 0.2 mm) were obtained after mixing 1 μl protein (20.3 mg ml−1 in GPC buffer incl. 1.5 mM NADP+) with 1 μl of reservoir solution (5.25% (w/v) PEG 3350, 20% (v/v) ethylene glycol, 0.1 M MES pH 7.0, 52 mM MgCl , 52 mM CaCl ) and 0.2 μl seed stock. Diffraction properties were optimized by crystal dehydration: over a period of 4 h, crystals were transferred to a dehydration solution (0.05 M MES pH 7.0, 25% ethylene glycol, 25% PEG 3350, 56 mM MgCl , 56 mM CaCl , 1.5 mM NADP+) by a step-wise exchange of the drop solution. All crystals were harvested and plunge-frozen in liquid nitrogen. Integrity of the protein in final crystals was examined by denaturing polyacrylamide gel electrophoresis. All data sets were collected at the Swiss Light Source (Villigen, Switzerland) at a temperature of 100 K. Data sets of DH crystals were collected at beamline X06DA (P2 , λ = 0.999870 Å, T = 100 K; P2 2 2, λ = 0.97626 Å). All other data sets were collected at beamline X06SA (KS–AT, λ = 0.97940 Å; DH–ΨKR–ER–KR, λ = 0.97626 Å). Data reduction was performed using XDS32 and XSCALE32, and data sets were analysed with phenix.xtriage33. All structures were solved with PHASER34 using molecular replacement. Crystals of all KS–AT di-domain variants of MAS are isomorphic in space group P4 2 2. The KS and AT domains of DEBS KS -AT (ref. 11) were used as molecular replacement templates and initial rebuilding was achieved by BUCCANEER35. All three crystal structures were virtually identical except for the identity of the last ordered C-terminal residue. The construct with the most extended C terminus (1–892) revealed aa 887 as the last ordered residue, overlapping in sequence with the modifying region. Thus we continued refinement only for crystals of this variant (aa 1–892) with unit cell constants of a = 77.5 Å, b = 77.5 Å, c = 371.2 Å, and a solvent content of 56%. A final model was obtained after iterative cycles of real space model building in COOT36 and TLS refinement in Phenix33, and was refined to R /R values of 0.21/0.23 at 2.3 Å resolution with excellent geometry (Ramachandran favoured/outliers: 97.8%/0.2%) (Extended Data Table 1a). Crystals of the DH domain of MAS belong to space group P2 (a = 59.7 Å, b = 162.4 Å, c = 66.6 Å, β = 91.4°) and P2 2 2 (a = 67.1 Å, b = 162.2 Å, c = 59.5 Å) with a solvent content of 49% and 51%, respectively. A molecular replacement search model was based on CurK DH15. Initial maps were improved by density modification and NCS averaging with PARROT37, followed by automated rebuilding with BUCCANEER35. Final models were obtained after iterative cycles of model building in COOT36, and refinement in BUSTER38 (P2 ) and Phenix33 (P2 2 2), yielding excellent geometry (Ramachandran favoured/outliers: P2 = 98.2%/0.0%; P2 2 2 = 98.2%/0.2%) and R /R values of 0.18/0.20 (P2 ) and 0.15/0.18 (P2 2 2) (Extended Data Table 1a). Crystals of MAS DH–ΨKR–ER–KR in space group P1 (a = 151.4 Å, b = 190.4 Å, c = 270.8 Å, α = 95.6°, β = 91.9°, γ = 103.7°) diffracted to a maximum resolution of 3.75 Å. The asymmetric unit contained 18 protomers in 9 dimers with 20,502 amino acids and a molecular mass of 2.2 MDa at 65% solvent content. Data were collected at four different positions of a single crystal and combined to obtain a complete high-quality data set. The resolution cutoff was determined by CC criterion39. Self-rotation functions revealed rotational NCS and the native Patterson function indicated translational NCS. Initially, a partial molecular replacement solution was obtained for the ER dimer using the ER domain of porcine FAS (pFAS)6. Other known structures of homologous domains did not provide efficient search models. The structure of the isolated MAS DH domain, determined here independently, yielded equivalent solutions in agreement with the pFAS ER based solution. For final structure determination, both models where used in subsequent rounds of molecular replacement. Start models for building further regions were generated by homology modelling using Swiss Model40. To allow unbiased refinement in real and reciprocal space, phenix.reflection_tools33 was used to define a thin-resolution shell-based test set41, and test set reflections were excluded from calculating maps, which were used for real-space refinement. Initial refinement cycles included rigid body refinement and restrained refinement. The impact of various low-resolution restraint formulations on refinement was tested carefully. Local NCS is particularly well suited for MAS DH–ΨKR–ER–KR intermediate resolution refinement because of the high degree of NCS and the fact that using local NCS restrains avoids any external standard restraints based on assumptions on secondary structure or homologous peptide structures. Thus local structural similarity restraints42 were combined only with reference model restraints to the authentic DH domain structure using autopruning in BUSTER38. After every round of refinement, bias-reduced, solvent flattened, and NCS-averaged maps were calculated using DM43 without applying phase combination. Sharpened NCS-average maps were generated by applying a sharpening B-factor to the structure factor amplitudes before averaging. Initially, real-space rigid body fitting of individual secondary structure elements was applied for instances of every domain type (DH, ΨKR, ER , ER , KR) followed by symmetry expansion and rigid body fitting for entire domains. Best-defined regions of the electron density maps were used for rebuilding of every domain type using Coot36 and O44, respectively, symmetry expanded, and recombined into 18 chains. At this point, unambiguous difference electron density indicated the connecting linkers, which were manually built into the maps and refined without symmetry expansion (Extended Data Fig. 2d, e). Later refinement cycles included TLS refinement, using one group per domain and linker, individual B-factor refinement, and automated weight factor determination. During rebuilding, B-sharpening, NCS average, and density modification, as well as feature enhanced maps33 were used. Overall, the use of 18-fold-domain-wise NCS averaging results in highly accurate and unbiased phase determination, irrespective of details of the atomic model. The combined use of NCS-averaging and B-factor sharpening led to an exceptional map quality typical for maps at considerably higher resolution (Extended Data Fig. 2f). Bound NADP+ cofactors were added for final refinement cycles. NADP+ is well ordered in the ER domain, while the nicotinamide moieties are disordered in the KR domains and were not included in the final model. A total of five KR and four ΨKR domains, which lack stabilization by crystal contacts, were either disordered or present in multiple orientations, and not included in the final model, despite substantial positive difference density. The KR domain in chain L shows a considerably more tilted orientation as observed in all other instances of the KR domains, which, however, agrees with the identified hinge regions. A single model was placed for this domain, which achieved the largest improvement of R-factors and was characterized by the lowest B-factors after refinement, although a secondary alternative conformation might be present. The refinement of the final model (excluding disordered regions (chains): 883–895 (E–R), 1206–1213, 1283–1287, 1948–1960, ΨKR(I/L/O/Q–R), KR(I/O/Q–R)) was completed with R /R values of 0.23/0.24 and very good geometry for the resolution range (Ramachandran favoured/outliers: 91.6%/1.8%). To determine oligomeric states in solution, sedimentation equilibrium analytical ultracentrifugation experiments were performed for MAS DH–ΨKR–ER–KR and MAS KS–AT. Columns (140 μl) containing proteins at concentrations of 3.5–4.5 mg ml−1 in GPC buffer were subjected to centrifugation at 4,800 and 7,800 r.p.m. in the Beckman An-50 Ti rotor, corresponding to 1,800g and 4,760g at the radial midpoint of the solution column, at 12 °C, with detection by radial absorbance scanning at 305 nm. At each speed, centrifugation was allowed to proceed until sedimentation equilibrium was attained, as judged by pairwise comparison of scans using the approach to equilibrium function in SEDFIT ( https://sedfitsedphat.nibib.nih.gov). Buffer density (1.0277 g ml−1) and viscosity (1.5306 centipoise) were measured at 12 °C using an Anton Paar DMA4500M densitometer and an AMVn viscometer, respectively. Molar extinction coefficients at 305 nm were calculated for each protein from the ratio of observed absorbance at various wavelengths in spectra at different dilutions and calculated molar extinction coefficients. The partial specific volume for each protein was calculated from sequence in SEDFIT. The radial absorbance scans at equilibrium for the two speeds were globally fitted to the ‘single species of interacting system’ mode in SEDPHAT45 to determine the apparent molecular mass of the protein in solution. If the obtained molecular mass was intermediate between the value expected for a monomer and a dimer, the data were globally fitted to the monomer–dimer association model in SEDPHAT, with the molecular mass of the monomer fixed to the value calculated from the sequence. In both cases data were fitted using a fixed meniscus position, a floating bottom position, mass conservation constraints, a floating baseline and fitting radially independent noise components. Confidence intervals on single-species masses or dissociation constants were obtained by the Monte-Carlo method implemented in SEDPHAT. SAXS data were collected at beamline X12SA of Swiss Light Source. Samples were dialysed into GPC buffer, diluted to concentrations between 3 and 10 mg ml−1and centrifuged at 13,000g and 8 °C until measurement. Glass capillaries (1 mm inner diameter) were mounted on a temperature-controlled holder at 12 °C. Data collection was performed using a Pilatus 2M detector at a distance of 2.14 m and a wavelength of 1.000 Å. Data were collected in eight repetitive scans each including ten 40 ms acquisitions at ten capillary positions yielding a total of 800 frames per buffer and protein, respectively. Frames with artefacts for example, from air bubbles, were identified using Swiss Light Source/PSI software (SAXS_inspect2) and excluded from the data sets. Radial averages were calculated and exported using beamline software for scattering vectors from 0.005 to 0.7 Å−1 defined as q = 4π/λsinθ. Scattering curves were averaged using DATAVER46; buffer profiles were subtracted using DATOP46. Scaling factors and P values of a Student’s t-test were analysed using DATMERGE46 and DATCMP46, respectively. Later frames were affected by increasing radiation damage and were excluded from further processing. Final scattering curves for each sample concentration were thus obtained from 300 individual profiles. The radius of gyration (R ) and zero angle intensity (I(0)) was calculated from the Guinier approximation using AUTORG46 and is consistent with values obtained from atomic distance distributions p(r) using DATGNOM46 (Extended Data Table 1b). Scattering profiles at different concentrations were only combined if a noise reduction at medium and high scattering vectors could be obtained. Modifying regions bear an intrinsic flexibility, which requires a flexible fitting approach to sample the full conformational space of the structures. Some approaches for flexible SAXS fitting have been described47, 48, but none was able to refine an individual structure while maintaining two-fold symmetry. Therefore, we combined dynamic elastic network restraints from CNS49 with SAXS-target refinement and two-fold symmetry averaging in XPLOR-NIH50 for the refinement of individual structures by simulated annealing. SAXS scattering curves of atomic models, fits with experimental data, and distance distributions were calculated using CRYSOL46 and DATGNOM46. All SAXS curves were plotted using Python Matplotlib. To compare calculated and experimental SAXS scattering curves, three models for the architecture of modifying regions were generated on the basis of the crystal structure of the domain-swapped SpnB fragment (ER–KR/ΨKR). The first model was obtained according to the original publication19 by superposing the monomeric ER–KR/ΨKR domain on the KR domain of pFAS6. A linear homology model of SpnB DH40 was placed into the position of pFAS DH and the domain swap in SpnB ER–KR/ΨKR was replaced with the corresponding region from DEBS KR (ref. 51). The second model was constructed in the same way via a superposition on MAS KR. The relative domain arrangement of SpnB ER–KR/ΨKR was not altered in these two models, only the domain swap was corrected. The third, more generalized modPKS model was constructed to verify if shorter ER–KR linkers are in contradiction with the architecture of MAS. As a representative for short ER–KR linkers, the structure of SpnB ER–KR/ΨKR (6 aa) and the corresponding DH homology model were modelled as individual domains on MAS, while the linear DH dimer was maintained. ΨKR–ER linkers could be readily reconnected and regularized, whereas the ER–KR linker required a tilt of the ΨKR/KR domain. The tilt maintained a reasonable distance between the C terminus of the DH and the N terminus of the ΨKR domain, and yielded a linker architecture of a modPKS in agreement with MAS without stable direct interdomain contacts. SAXS curves and distance distributions were calculated for all models and compared with experimental SAXS scattering curves of MAS and two modPKS modifying regions with short ER–KR linkers (GpEryA, 9 aa; MsPks, 8 aa). Related structures were identified using PDBeFold52 and interfaces were analysed using QtPISA53. Transformations and coordinate manipulations were performed using CCP4 (ref. 54) tools, MODTRAFO (T. Schirmer; http://www.biozentrum.unibas.ch), and MOLEMAN55. The automated Oligo algorithm56 as implemented in Swiss Model unambiguously detected and predicted a single mode of dimerization of MAS KS–AT based on sequence homology. Initially, the dimeric form of KS–AT was assembled by least-squares fitting of secondary structure elements on DEBS KS (ref. 11). Then, all residues in a radius of 7.5 Å to the dimer interface were deleted and multi-template homology modelling using modeller 9.15 (ref. 57) was used to construct a full-length dimeric homology model based on 20 homodimeric PKSs/FASs KS structures and the interface deleted MAS KS–AT structure. Remodelled regions (excluding all crystallographically defined regions beyond the radial cutoff) were geometry minimized using phenix.geometry_minimization33. The position where the post-AT linker becomes disordered was located by crystallization of KS–AT di-domains with three different linker lengths (1–884, 1–887, 1–892). Normal mode analyses were performed using the Bio3D58 library in ‘R’. Hinge bending analysis was performed by pre-aligning all structures to a reference substructure using LSQKAB59, followed by a MODTRAFO (T. Schirmer; http://www.biozentrum.unibas.ch) analysis of the moving substructure. Principle screw axes were determined by averaging the direction vectors of the screw axes using Python Numpy and locating a central hinge point from the position of all screw axes. Active-site distances were calculated using BIOPYTHON60. All axes were visualized using PYMOL61. Interdomain angles of DH dimers were calculated by pre-aligning all DH dimers to one DH domain of MAS DH, followed by calculating the angle between the first principle component vector of the secondary structure elements of both domains. The angles were visualized using PYMOL61. Bias-removal for F − F omit maps was achieved by applying a random perturbation to coordinates (Δ0.2 Å) and B-factors (Δ20% of the mean overall B-factor) using MOLEMAN2 (ref. 55) before refinement. Figures, videos, and active-site tunnels were generated using PYMOL61, LSQMAN62, and CAVER 3.0 (ref. 63). Fifty-five sequences containing fully reducing modifying regions were selected from FASs, fiPKSs, Msl-, one trans-AT, and 36 modPKSs modules. Structure-based sequence alignments of all PKSs/FASs type I domain structures were generated using PDBefold52 and used as reference for the alignment of individual domains using ClustalW2 (ref. 64). Linkers were aligned without reference, assembled with the individual domain alignments, and manually corrected in Geneious version 7.1.7 (ref. 65). Phylogenetic trees were generated using the neighbouring joining algorithm in Geneious version 7.1.7 (ref. 65).
News Article | April 6, 2016
No statistical methods were used to predetermine sample size. The experiments were not randomized, and the investigators were not blinded to allocation during experiments and outcome assessment. Wild-type and mutant versions of human DDB1 (Q16531), human CRBN (Q96SW2), human CK1α (P48729) and human IKZF1 (Q13422) were subcloned into pAC-derived vectors29 and recombinant proteins expressed as N-terminal His , StrepII or StrepII-Avi fusions in Trichoplusia ni High-Five insect cells using the baculovirus expression system (Invitrogen). For purification of His -DDB1-His -CRBN, His -DDB1∆BPB-StrepII-CRBN∆1–40 and His -DDB1∆BPB-His -CRBN, cells were resuspended in buffer containing 50 mM tris(hydroxymethyl)aminomethane hydrochloride (Tris-HCl) pH 8.0, 200 mM NaCl, 0.25 mM tris(2-carboxyethyl)phosphine (TCEP), 1 mM phenylmethylsulfonyl fluoride (PMSF), 1× protease inhibitor cocktail (Sigma-Aldrich) and lysed by sonication. Cells expressing StrepII-Avi-CK1α or truncated versions of StrepII-Avi-IKZF1 (Δ256–519, Δ197–238/Δ256–519 and Δ1–82/Δ197–238/Δ256–519; full-length IKZF1 forms aggregates during purification) were lysed in the presence of 50 mM Tris-HCl pH 8.0, 500 mM NaCl, 0.25 mM TCEP, 1 mM PMSF and 1× protease inhibitor cocktail (Sigma-Aldrich). Following ultracentrifugation, the soluble fraction was passed over Strep-Tactin Sepharose (IBA) or His-Select nickel affinity resin (Sigma-Aldrich) and following elution, affinity tags were removed from CK1α and IKZF1 by overnight TEV protease treatment as indicated. The affinity-purified protein was further purified via ion exchange chromatography (Poros 50HQ and 50HS) and subjected to size-exclusion chromatography in 50 mM HEPES pH 7.4, 200 mM NaCl and 0.25 mM TCEP. The protein-containing fractions were concentrated using ultrafiltration (Millipore) and flash frozen (DDB1–CRBN constructs at 40–120 μM, TEV cleaved CK1α at ~280 μM, StrepII-Avi-CK1α at ~50 μM, TEV cleaved IKZF1 at ~350 μM and StrepII-Avi-IKZF1 at ~150 μM). Proteins were stored at −80 °C. Attempts to crystallize CK1α, lenalidomide and CRBN with the full-length DDB1 adaptor protein were unsuccessful. Because the WD40 β-propeller B (BPB) of DDB1 is not involved in CRBN binding, we generated a human DDB1 construct in which a GNGNSG-linker replaced the BPB domain (residues 396–705; DDB1∆BPB). For crystallization of the DDB1∆BPB–CRBN–lenalidomide–CK1α complex, His DDB1∆BPB–StrepII–CRBN∆1–40 at 70 μM was mixed with lenalidomide at 80 μM before the addition of TEV cleaved full-length CK1α at 80 μM. The mixture was incubated on ice for 1 h and subsequently centrifuged at 20,000g for 30 min at 4 °C. Crystallization plates were set up and stored at room temperature. Crystals appeared within 3 days after mixing the protein solution 1:1 with the reservoir containing 70 mM Tris pH 7.0, 140 mM MgCl and 7% (w/v) PEG 8000 and continued growing until day 13 in MRC 2 Well Crystallization format vapour diffusion plates (Swissci). Crystals were cryo-protected in reservoir solution supplemented with 20% ethylene glycol and flash-cooled in liquid nitrogen. Diffraction data were collected at the Swiss Light Source (beamline PXII) with a Pilatus 6M detector at the wavelength of the Zn-edge (1.28162 Å) and a temperature of 100 K. Data were indexed and integrated using XDS30 and scaled using AIMLESS supported by other programs of the CCP4 suite31. The optimal high-resolution cut-off (2.45 Å) was determined based on the half-set correlation criterion32 (CC = 0.45 for the highest resolution shell). Data processing statistics, refinement statistics and model quality parameters are provided in Extended Data Table 1. The DDB1∆BPB–CRBN–lenalidomide–CK1α quaternary complex crystallized in space group P1 with two complexes in the unit cell. PHASER33 was used to determine the structure by molecular replacement using a crystallographic model of DDB1 omitting the BPB domain, a CK1α homology model generated with Modeller34 based on a CK1δ crystal structure (PDB entry 4TWC), and human CRBN (PDB entry 4TZ4) as search models. The initial model was iteratively improved with COOT and refined using PHENIX.REFINE35 and autoBUSTER36, with ligand restraints generated by Grade server (Global Phasing). Figures were generated with PyMOL (The PyMOL Molecular Graphics System, Version 184.108.40.206 Schrödinger, LLC) and model quality was assessed with MOLPROBITY37. Interaction surfaces were determined with PISA38. The IKZF1 homology model was calculated using Modeller based on a multiple sequence alignment with the experimental zinc-finger structures of Aart, YY1 and Kasio (PDB entries 2I13, 1UBD, 2LT7). Purified human NEDD8(M1C) (NEDD8) was incubated with DTT (8 mM) at 4 °C for 1 h. DTT was removed using a S200 16/60 gel filtration column in a buffer containing 50 mM Tris pH 7.3 and 150 mM NaCl. Alexa-488–C5-maleimide (Invitrogen) was dissolved in 100% DMSO and mixed with NEDD8 to achieve fourfold molar excess of Alexa-488–C5-maleimide. NEDD8 labelling was carried out at room temperature for 3 h in a vacuum desiccator and stored overnight at 4 °C. Labelled NEDD8 was purified on a S200 16/60 gel filtration column in 50 mM Tris pH 7.5, 150 mM NaCl, 0.25 mM TCEP and 10% (v/v) glycerol, concentrated by ultrafiltration (Millipore), flash frozen (~40–80 μM) in liquid nitrogen and stored at −80 °C. In vitro CRL4CRBN reconstitution and CUL4A neddylation was performed as described4, 19, 21. His –CUL4A–His –RBX1 at 3.5 μM was incubated with His –DDB1–His –CRBN at 1.5 μM (wild-type or mutant forms) in a reaction mixture containing 3.8 μM Alexa-488–NEDD8, 50 nM NAE1/UBA3 (E1), 150 nM UBC12 (E2), 1 mM ATP, 50 mM Tris pH 7.5, 100 mM NaCl, 2.5 mM MgCl , 0.5 mM DTT and 5% (v/v) glycerol for 2 h at room temperature. Gel filtration purified neddylated CRL4CRBN ( CRL4CRBN) was concentrated to 2–4 μM, flash frozen and stored at −80 °C. Alexa-488–NEDD8 labelling efficiency of different CRL4CRBN mutant complexes was determined by measuring total fluorescent intensity at equal concentration (excitation at 490 nm, emission at 540 nm) using a Safire2 microplate reader from Tecan (Extended Data Fig. 4f). Purified StrepII-Avi-tagged CK1α or IKZF1 were biotinylated in vitro at a concentration of 25–50 μM by incubation with final concentrations of 2.5 μM BirA enzyme and 0.2 mM D-Biotin in 50 mM HEPES pH 7.4, 200 mM NaCl, 10 mM MgCl , 0.25 mM TCEP and 20 mM ATP. The reaction was incubated for 1 h at room temperature and stored at 4 °C for 14–16 h. Biotinylated proteins were purified by gel filtration chromatography and stored at −80 °C (StrepII-Avi-CK1α at ~25 μM, StrepII-Avi-IKZF1 at ~40 μM). Increasing concentrations of Alexa-488–NEDD8-labelled CRL4CRBN ( CRL4CRBN) were added to pre-mixed biotinylated IKZF1 at 80 nM or CK1α at 100 nM, terbium-coupled streptavidin at 4 nM (Invitrogen) and IMiDs or 2′-deoxyuridine at 5 μM (final concentrations) in 384-well microplates (Greiner, 784076) in a buffer-containing 50 mM Tris pH 7.5, 100 mM NaCl, 0.1% pluronic acid and 1% DMSO (see also figure legends). Before TR-FRET measurements were conducted, the reactions were incubated for 15 min at room temperature. After excitation of terbium (Tb) fluorescence at 337 nm, emission at 490 nm (Tb) and 520 nm (Alexa 488) were recorded with a 70 μs delay to reduce background fluorescence and the reaction was followed over 1 h by recording 60 technical replicates of each data point using a PHERAstar FS microplate reader (BMG Labtech). The TR-FRET signal of each data point was extracted by calculating the 520/490 nm ratio. Data were analysed with GraphPad Prism 6 assuming equimolar binding of the probe to the receptor ( CRL4CRBN) using the following equations: The concentration of the receptor in the bound state, [C ], can be calculated for that setting by the law of mass action: K is the equilibrium constant for the dissociation, and [C ] and [C ] are the total concentrations of the probe and the receptor, respectively. The K value can be calculated from the change in the fluorescence intensity, FI, observed by a titration of the receptor at constant probe concentrations according to: FI is the observed fluorescence intensity, and FI and FI the fluorescence intensities of the probe in its free and its bound states, respectively. The assay window is described by the overall change in the fluorescence intensity, (FI − FI ). Counter titrations with unlabelled proteins were carried out by mixing CRL4CRBN at 0.5–1 μM with 200 nM biotinylated CK1α or 160 nM biotinylated IKZF1 in the presence of 8 nM terbium-coupled streptavidin and IMiDs at 10–20 μM. After 15 min incubation on ice, increasing amounts of unlabelled DDB1∆BPB–CRBN (0.04–40 μM) or IKZF1 (0.04–20 μM; wild-type or mutant forms or pre-incubated with consensus or control DNA as stated in the EMSA section) were added to the pre-assembled CRL4CRBN–CK1α/IKZF1 complexes in a 1:1 volume ratio and incubated for 5 min at room temperature. IMiD titrations were carried out by premixing CRL4CRBN, CK1α/IKZF1 and 8 nM terbium-coupled streptavidin before addition of increasing concentrations of each IMiD (0.005–5 μM) in a 1:1 volume ratio (see figure legends for final concentrations). The 520/490 nm ratios were plotted to calculate the half maximal effective concentrations (EC ) or half maximal inhibitory concentrations (IC ) assuming a single binding site using GraphPad Prism 6. IC values were converted to the respective K as described39. Three biological replicates were carried out per experiment. The standard deviation was derived from the sum of the mean absolute error of 10 technical replicates (per data point and replicate) and the standard deviation of the biological replicates. Cy5-conjugated thalidomide4 (10 nM) was mixed with increasing concentrations of either wild-type or mutant forms of purified DDB1–CRBN (0.004–2 μM) in a 384-well microplate (Greiner, 784076) and incubated for 30 min at room temperature. CRBN–IMiD interactions were measured in 50 mM Tris pH 7.5, 100 mM NaCl, 0.1% pluronic acid and 1% DMSO by change in fluorescence polarization using a PHERAstar FS microplate reader (BMG Labtech). The Cy5-thalidomide bound fraction was calculated as described40. Data were plotted and analysed using GraphPad Prism 6 assuming a single IMiD binding site on CRBN. Hct116 cells were purchased from The European Collection of Cell Cultures (ECACC, Sigma-Aldrich), immediately used for experiments and regularly tested for mycoplasma contamination. Cells were cultured in L-lysine and L-arginine free DMEM supplemented with unlabelled L-lysine, L-arginine, 10% FBS and 2 mM L-glutamine. Cells were grown to approximately 50% confluency and the medium was exchanged for DMEM supplemented with 13C,15N-labelled L-lysine (Lys8) and 13C,15N-labelled L-arginine (Arg10) containing lenalidomide at 30 μM or equivalent amounts of DMSO as control. Cells were incubated for 16 h and harvested for mass spectrometry analysis in 0.5 M Tris-HCl pH 8.6, 6 M guanidine hydrochloride, reduced in 16 mM TCEP for 30 min, and alkylated in 35 mM iodoacetamide for 30 min in the dark. The proteins were digested at 37 °C with lysyl endopeptidase (Wako) after dilution to ~2 M guanidine hydrochloride (with 50 mM Tris-HCl pH 7.3, 5 mM CaCl buffer) for 6 h, and after dilution to <1 M guanidine hydrochloride with trypsin (Promega) at 37 °C overnight. The resulting peptides were desalted using C solid state extraction cartridges and offline fractionated into 36 fractions by basic reverse phase chromatography. The 36 fractions were recombined to a final of 12 samples. Generated peptides were separated on an EASY n-LC 1000 liquid chromatography system equipped with a C EASY-Spray column coupled to an Orbitrap Fusion mass spectrometer (all from Thermo Scientific). Maxquant41 was used for .RAW file processing and controlling peptide and protein level false-discovery rates, assembling proteins from peptides, and protein quantification from peptides. Peptides were searched against a human Uniprot database with both forward and reverse sequences. For analysis, we did not utilize the SILAC component of the experiment, but instead used the Maxquant LFQ algorithm to quantify the relative abundance of casein kinase isoforms across three independent replicates. In vitro ubiquitination was performed by mixing wild-type or mutant CRL4CRBN at 70 nM with a reaction mixture containing IMiDs at 350 nM or 10 μM, CK1α at 500 nM, E1 (UBA1, BostonBiochem) at 40 nM, E2 (UBCH5a, BostonBiochem) at 1 μM, wild-type (20 μM) or lysine-free (10 μM) ubiquitin as indicated. Reactions were carried out in 50 mM Tris pH 7.5, 30 mM NaCl, 5 mM MgCl2, 0.2 mM CaCl , 1 mM ATP, 0.1% Triton X-100 and 0.1 mg ml−1 BSA, incubated for 15–30 min at 30 °C and analysed by western blot using anti-CK1α (abcam, ab108296, 1:20,000) and anti-rabbit IRDye 800CW antibodies (LI-COR, 926-32211, 1:10,000). Blots were scanned on a LI-COR Odyssey infrared imaging system. To identify target lysines following in vitro ubiquitination of CK1α in the presence or absence of lenalidomide, samples were precipitated with 20% (v/v) trichloroacetic acid followed by several acetone washes. The precipitated protein was dissolved in 10 μl of 500 mM Tris pH 8.6, 6 M guanidine hydrochloride and 8 mM TCEP and incubated with 18 mM iodoacetamide for 30 min at room temperature in the dark. After addition of 50 μl 50 mM Tris pH 7.4, 5 mM CaCl , samples were incubated with 200 ng trypsin at 37 °C for 12–14 h. Mass spec was carried out on a FUSION Orbitrap, the data was searched with MASCOT, site probabilities were calculated with ASCOR and peak integration for relative quantification was performed using ProgenesisLC. Equimolar ratios of complementary DNA single strands at 200 mM were mixed in 10 mM Tris pH 8.0, 50 mM NaCl and 1 mM MgCl , incubated at 95 °C for 2 min and annealed by slowly lowing temperature using a thermal cycler. IKZF1 was mixed with duplex DNA (consensus DNA: 5′-TCAGAAAAAGGGAATTCCGTCAC-3′; control DNA: 5′-TCAGACACTTTTGGTACTGTCAC-3′) in 50 mM HEPES pH 7.4, 200 mM NaCl, 0.25 mM TCEP and 10% (v/v) glycerol and incubated on ice for 15 min before the addition of DDB1–CRBN and pomalidomide, as indicated. Binding reactions were incubated for 30 min at room temperature, applied to a 4–16% NativePAGE Novex Bis-Tris gel (Invitrogen) and separated in 1× NativePAGE buffer at 150 V for 75 min at room temperature. Gels were stained with 1 μg ml−1 ethidium bromide in 1× NativePAGE buffer followed by Coomassie staining.
News Article | March 18, 2016
A team from Helmholtz-Zentrum Berlin has been able to measure how new bonds influence molecules for the first time: they have reconstructed the energy landscape of acetone molecules using measurement data from the Swiss Light Source (SLS) of the Paul Scherrer Institut, and, thereby, empirically established the formation of hydrogen bonds between acetone and chloroform molecules. The results have been published in Nature Scientific Reports and assist in understanding fundamental phenomena of chemistry. Molecules are composed of atoms that maintain specific intervals and angles between one another. However, the shape of a molecule can change, for example, through proximity to other molecules, external forces and excitations, and also when a molecule makes a chemical connection with another molecule, for instance in a chemical reaction. A very useful concept in describing the changes that are possible in molecules is the use of what are called “potential surfaces” or energy landscapes. However, these are not actual surfaces in real space. They are more viewed as parameters defining the molecule, which can then be portrayed as a surface. An example would be the stretching of a carbon-oxygen bond, or the angle between various molecular groups. You can imagine such surfaces as being like hilly landscapes. If light excites part of the molecule into oscillation, the state of the molecule moves upward, energetically speaking, perhaps even up over a pass or a peak. It either returns finally to its previous energy minimum, or lands in a different energy dip that corresponds to altered angles or bond lengths. Some of these changes allow us to draw conclusions about hydrogen bonding with neighboring molecules. Response after excitation of the double bond C=O analyzed The team headed by Annette Pietzsch and Alexander Föhlisch has now for the first time succeeded in precisely measuring these extremely subtle surfaces surrounding a small molecule named acetone (C H O). They used the resonant inelastic X-ray scattering (RIXS) method at the Swiss Light Source of the Paul Scherrer Institut (PSI) in Switzerland for this work. “We chose to selectively excite the double bond between the carbon and oxygen atom of acetone into oscillation and analyzed the responses in detail,” explains Annette Pietzsch. Thanks to the extremely high resolution of the measurement data, they were successful in mapping the potential surface along this C=O double bond. In the second part of the experiment, they investigated a mixture of acetone and chloroform. A liquid mixture like this is denoted as azeotropic, meaning that the two ingredients can no longer be separated from one another through distillation. The scientists were now able for the first time to empirically observe how the acetone molecules linked tightly to the chloroform molecules via hydrogen bonding. They were able to identify in the measurement data the fingerprint of the hydrogen bonds that form between the C=O group of the acetone molecules and hydrogen groups of the chloroform molecules. “In conclusion, we demonstrated how sub-natural line width vibrational resolved RIXS gives direct experimental access to the ground state potential energy surface around selected atomic sites and moieties, not accessible with other techniques. Our approach to the local ground state potential energy surface (...) resembles finding a needle in a haystack,” writes the team in its contribution published in the periodical Nature Scientific Reports. The performance of this approach will benefit strongly from upcoming high-brilliance synchrotrons and free-electron lasers in combination with upcoming high resolution RIXS instruments. Therefore, they foresee wide applicability of this technique to all thermal, collective and impurity driven chemistry and materials issues in the near future. Annette Pietzsch works at the BESSY II synchrotron source in Berlin, setting up METRIXS—an instrument for resonant inelastic X-ray scattering that will be able to achieve considerably higher resolution in the future. In addition, the meV-RIXS experiment will make high-resolution X-ray scattering in low-energy regions feasible. Alexander Föhlisch heads the HZB Institute for Methods and Instrumentation for Research with Synchrotron Radiation and is spokesperson of Helmholtz Virtual Institute for Dynamic Pathways in Multidimensional Landscapes (Helmholtz Virtual Institute 419).