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News Article | August 23, 2016
Site: www.sciencedaily.com

Permanent brain damage from a stroke may be reversible thanks to a developing therapeutic technique, a study has found. The novel approach combines transplanted human stem cells with a special protein that the US Food and Drug Administration already approved for clinical studies in new stroke patients. The researchers say they are the first to use 3K3A-APC to produce neurons from human stem cells grafted into the stroke-damaged mouse brain.

The proteasome, a ~2.5-MDa molecular machine, is the focal point for global regulation of ubiquitin pathway output10. Among its major regulators is a deubiquitinating enzyme known in mammals as USP14 (in yeast, Ubp6). USP14 negatively regulates proteasome activity by ubiquitin chain disassembly as well as by a noncatalytic mechanism1, 2, 3, 4, 5, 6. USP14 inhibitors produce selective effects on the turnover of proteasome substrates3, 8, suggesting that the rate at which USP14 disassembles proteasome-bound ubiquitin chains may depend on the nature of the substrate. We test this hypothesis in the present work. USP14 is thought to progressively trim monoubiquitin groups from the substrate-distal tip of a chain9 (Fig. 1a). To assess this model, we employed a canonical substrate of USP14, ubiquitinated cyclin B1 (refs 1, 3). When cyclin B1 is modified by the ubiquitin ligase anaphase-promoting-complex/cyclosome (APC/C) in the presence of E2 enzyme UbcH10, the resulting conjugates typically carry multiple short ubiquitin chains of varied length11, 12. We refer to these as supernumerary chains, insofar as only one chain is in principle required for substrate degradation. The flexible, 88-residue amino-terminal element of cyclin B1 (NCB1) is enriched in lysine residues, nine of which are competent for ubiquitination11. The N-terminal element is necessary for cyclin B1 degradation, and contains an APC/C recognition motif, the destruction box11, 12, 13. Ubiquitinated NCB1 was rapidly degraded by proteasomes lacking USP14, but in the presence of USP14 deubiquitinated species were observed. These were not fully deubiquitinated, but rather carried 2–4 ubiquitin groups and were resistant to both degradation and further deubiquitination (Fig. 1b). This strong stop to USP14-dependent processing might be assumed to reflect that these conjugates retain too few ubiquitin moieties for efficient proteasome binding. However, studies below show that USP14 activity depends on the architecture of ubiquitin chains on the substrate and not simply substrate binding to the proteasome. The behaviour of NCB1 was comparable to that of full-length cyclin B1 (data not shown)3, 12. We used the experimental system described above to test whether the architecture of ubiquitin chains on NCB1 was relevant to its rapid deubiquitination. Cyclin B1 is modified by APC/C and UbcH10 to form mixed-linkage chains11 primarily via residues K11, K48, and K63. Thus, a lysine-free mutant of ubiquitin12 will modify NCB1 via multiple mono-ubiquitination events (Fig. 1c). USP14 deubiquitinated this form of NCB1 at a rate comparable to that of wild-type conjugates, indicating that USP14 does not act obligatorily on ubiquitin–ubiquitin linkages (Fig. 1c). In a complementary experiment, we eliminated lysines from NCB1 rather than ubiquitin, leaving behind only K64 of NCB1 to allow for the synthesis of a single ubiquitin chain on NCB1. This mutant is competent for degradation (Fig. 1d)12. Although single-chain conjugates are canonical proteasome substrates14, USP14 showed no detectable activity on this substrate (Fig. 1d). The lack of deubiquitinating activity on the single-chain substrate was accompanied by a failure to inhibit its degradation. Similar results were obtained with NCB1 ubiquitinated with Ubc4 in place of UbcH10 (Extended Data Fig. 1a, b). A caveat to the experiment above is that, because NCB1 is degraded quickly by the proteasome, USP14 has little time to act. However, the proteasome must be present in the assay because it is required to activate USP14. When we quenched substrate degradation by replacing ATP with ADP (Extended Data Fig. 2), USP14-dependent deubiquitination of WT NCB1 conjugates was preserved (Fig. 1e). In contrast, no deubiquitination was detected with single-chain conjugates (Fig. 1f). Thus, USP14 shows marked specificity for NCB1 conjugates bearing multiple ubiquitin chains, whereas the proteasome does not effectively distinguish between these two classes of substrate in the absence of USP14. We next considered whether the resistance of single-chain conjugates to USP14 was idiosyncratic to the particular modified lysine in NCB1. On the contrary, conjugates generated using a K36-only form of NCB1 behaved equivalently to the K64-anchored conjugates (Extended Data Fig. 1c). The supernumerary chain effect was seen with the substrate PY-Sic1 as well as NCB1 (Extended Data Fig. 1d, e). We identified a third preferred substrate of USP14, securin, which is likewise modified at multiple lysines (Extended Data Fig. 1f)13, 15. Ubp6, the Saccharomyces cerevisiae orthologue of USP14, showed the same preference for supernumerary chains, indicating that this property is evolutionarily conserved (Extended Data Fig. 3). It has been hypothesized that USP14 or Ubp6 may preferentially cleave K63-linked chains16, 17. This model could account for our data if single-chain NCB1 conjugates were devoid of K63 linkages. To test this model, we prepared single-chain NCB1 with homogeneous K63 linkages. These conjugates were also cleaved slowly if at all by USP14 (Extended Data Fig. 4), regardless of length. One explanation for the specificity of USP14 for substrates modified at multiple sites is that cleavage of one chain is promoted allosterically by binding of a second chain to USP14 or another site on the proteasome. However, adding free chains over a wide concentration range failed to stimulate deubiquitination of single-chain NCB1 (Extended Data Fig. 5). Another measure that failed to stimulate removal of singleton chains was phosphomimetic substitution of Ser432, a site subject to modification by Akt18 (Extended Data Fig. 6). If singleton chains are as a rule poor substrates for USP14, then all free ubiquitin chains—those not attached to an acceptor protein—must be poor USP14 substrates, regardless of length or linkage type. Extended Data Fig. 7 shows that free chains of diverse length and linkage type are all cleaved minimally by proteasome-activated USP14, even upon long incubation. By ubiquitinating NCB1 with preformed tetraubiquitin chains, we prepared conjugates carrying multiple chains of homogeneous linkage type and length. When these conjugates were incubated with USP14, proteasomes, and either ATP or ADP, a limit digest product was formed consisting of NCB1 carrying four ubiquitin groups (Fig. 2a, b). Thus, USP14 seemed to completely remove each supernumerary chain, and stop when a single chain remained. Accordingly, ubiquitin was released from this substrate in the form of tetraubiquitin chains (Fig. 2c). Using mass spectrometry, we confirmed that K48 linkages were not consumed in this reaction (Fig. 2d). Studies with other linkage types (Extended Data Fig. 8) further indicated that USP14 removes chains preferentially en bloc (Fig. 2e), rather than trimming them from the distal tip, as previously thought. Why does USP14 discriminate against cleavage within chains? The presumptive active form of Ubp6 sits with its catalytic domain directly contacting the OB domain of subunit Rpt1 (refs 5,6). Modelling studies with diubiquitin as substrate indicated that, regardless of chain linkage type, the proximal ubiquitin may have restricted access to the active site of the enzyme (Fig. 2f, Extended Data Fig. 9). The coiled-coils of Rpt1 and Rpt2 are responsible for this occlusion (Fig. 2f). This effect may account for the dominant en bloc cleavage mode of Ubp6 and USP14. Cleavage within ubiquitin chains by free forms of USP14 and Ubp6, as observed by many researchers, may reflect that free USP14 is not subject to occlusion from proteasome subunits. In a recent study, we established a single-molecule TIRF (total internal reflection fluorescence) microscopy method to monitor deubiquitination events mediated by the en bloc zinc-dependent activity of RPN1119, 20, using USP14-free proteasomes15. Deubiquitination was identified as a stepwise reduction of signals from fluorescently-labelled ubiquitin on substrates (Fig. 3a, Extended Data Fig. 10). To study USP14-mediated deubiquitination, we inhibited RPN11 activity (as well as substrate degradation) using ADP and a zinc chelator15. ‘Stepped’ events representing deubiquitination increased sevenfold upon USP14 addition (Fig. 3a). Single-molecule traces with USP14-reconstituted proteasome in the presence of ADP showed remarkable similarity to those produced by RPN11 (Fig. 3b, Extended Data Fig. 10). The step size of ubiquitin removal by RPN11 and USP14 followed a similar distribution, supporting en bloc cleavage by USP14 (Fig. 3c). USP14-mediated deubiquitination reduced the dwell time of ubiquitin conjugates on the proteasome, suggesting that rapid deubiquitination can suppress degradation of the substrate by contracting the duration of its encounter with the proteasome (Fig. 3d). This dwell time effect suggests that, to suppress degradation, USP14 must remove chains at least as rapidly as the substrate is degraded in the absence of USP14. To test this scenario we used quench-flow methods, which allow reactions to be followed on a millisecond time scale. An excess of proteasome over substrate was employed to approximate single-turnover conditions. USP14-free proteasomes degraded NCB1 conjugates efficiently, with most degradation occurring between 9 and 30 s (Fig. 4a, b). USP14 delayed degradation (Fig. 4b), as high molecular weight conjugates were instead converted into partially deubiquitinated species. The production of lower molecular weight species through deubiquitination was initiated before degradation was evident with USP14-free proteasomes (Fig. 4a–c). Thus, USP14 is sufficiently fast that deubiquitination can be accomplished before the proteasome would otherwise degrade NCB1. Similar results were obtained using full-length cyclin B1 (data not shown), and we confirmed that deubiquitination was mediated by proteasome-bound rather than free USP14 (Extended Data Fig. 6b). The ability of USP14 to prevail over the proteasome in a kinetic competition may explain the suppression of protein degradation by USP14’s deubiquitinating activity3. Thus, USP14-loaded proteasomes may discriminate against substrates bearing multiple ubiquitin chains, whereas USP14-free proteasomes do not. Because USP14 and Ubp6 are modulated by growth factors and stress conditions2, 18, the ability of the proteasome to discriminate between single-chain and multi-chain substrates is likely to be under biological control. We expect that the supernumerary chain rule should bias USP14 action towards the products of ubiquitin ligases that are weak in chain-extending activity, thus adding ubiquitin to multiple sites on a target protein. Many ubiquitinated proteins can be modified at multiple lysines11, 12, 21, 22, as originally described for cell surface receptors23. In most cases however, it is unclear whether simultaneous modification of lysines occurs on endogenous substrates, because proteomics methods for analysis of ubiquitination generally rely on proteolytic digestion. Therefore, the development of new methodologies may be critical for a better understanding of how USP14 regulates proteasome function. We suggest that the last ubiquitin chain resists cleavage by USP14 because its site of attachment to substrate is not docked in proximity to USP14, regardless of the ubiquitin receptor to which the chain is bound. If the chain that is spared by USP14 is short, degradation will be suppressed; if it is long, then proteasome–substrate interaction will be preserved and substrate degradation will proceed. This chain will be removed by RPN11 once substrate translocation is initiated. Because RPN11 also cleaves chains proximally19, 20, our data reveal an unexpected point of similarity between these enzymes, which may act sequentially. However, RPN11 activity differs from that of USP14 in requiring ATP hydrolysis19, 20, presumably to drive translocation of the conjugate to its active site24, 25. That RPN11 promotes substrate degradation whereas USP14 suppresses degradation may reflect that RPN11 removes chains following the commitment step in the degradation pathway. It will be interesting to determine whether supernumerary chain removal is the principal role of USP14 in cells, or whether other specificity rules exist for this enzyme. Substrate flexibility, a feature of the cyclin B and securin degrons26, but not of ubiquitin itself, may be important for USP14 to act on these substrates. Indeed, the occlusion that we propose to impair docking of proximal ubiquitin in a free chain near the USP14 active site may apply as well to many folded globular domains that are ubiquitinated, which would be required to access the same site. This substrate docking constraint may further limit the substrate range of USP14. In contrast to USP14, RPN11 is expected to have a robust activity against chains on domains that are normally folded, because such chains may be presented to RPN11 only after the domain has been unfolded by mechanical force applied by the proteasomal ATPases.

C57BL/6J mice (CD45.2) and B6.SJL-PtprcaPep3b/BoyJ (CD45.1) were purchased from The Jackson Laboratory (Bar Harbour, ME). Prdm16Gt(OST67423)Lex knockout mice31 were obtained from Lexicon Genetics. Conditional Mito-Dendra2 transgenic (Pham) mice32 (B6;129S-Gt(ROSA)26Sortm1(CAG-COX8A/Dendra2)Dcc/J) and E2A-Cre mice33 (B6.FVB-Tg(EIIa-cre)C5379Lmgd/J) were purchased from Jackson Laboratory. Pham mice contain a mitochondrially targeted Dendra preceded by a stoplox sequence in the Rosa locus. These mice were crossed with E2A-Cre mice to effect ubiquitous induction of the MitoDendra2 reporter. Conditional Mfn2 knockout mice34 (B6/129SF1Mfn2tm3Dcc/Mmucd) were obtained from MMRRC and crossed to Vav-Cre transgenic mice35 (B6.Cg-Tg(Vav1-Cre)A2Kio/J) to obtain a homozygous floxed allele Mfn2 allele which generated a B6.Cg-Tg(Vav1-Cre)A2Kio/J;B6/126SF1 Mfn2tm3Dcc/Mmucd mixed mouse strain. All mouse strains were rederived by in vitro fertilization at the Jackson Laboratory. Animals were housed in a specific pathogen-free facility. Experiments and animal care were performed in accordance with the Columbia University Institutional Animal Care and Use Committee. All mice were used at age 8–12 weeks, except in experiments that involved fetal liver cells, when E14.4 embryos were used. Both sexes were used for experiments. Results were analysed in non-blinded fashion. In all experiments, randomly chosen wild type and littermates were used. MEFs were established from approximately 14.5 days post coitum embryos as previously described36 from Prdm16+/− breeder pairs. Briefly, dissected embryo trunks were minced into 1–2 mm fragments, resuspended in 3 ml 0.25% trypsin/EDTA (Gibco, Carlsbad, CA) and passed 20–30 times through a 16 gauge needle. Cell suspensions were incubated at 37 °C for 1 h with frequent agitation. Erythrocytes were lysed with ACK buffer, washed and cells were plated for 3 h in 10% FBS/DMEM. Cells remaining in suspension were aspirated and adherent cells were cultured with fresh media. MEFs were passaged 1:3 every 3 days and cells between passage 2 and 5 were used for all experiments. 293 cells and NIH-3T3 cells were purchased from ATCC (Manassas, VA) and sub-cultured in 10% FBS/DMEM or 10% calf serum/DMEM, respectively. WT and Mfn2−/− MEFs were a kind gift from E. Schon (Columbia University). All lines are tested yearly for mycoplasma contamination and found negative. Prdm16 constructs were generated by subcloning the murine full length (flPrdm16) or truncated (sPrdm16) cDNA into the XhoI/EcoRI sites of the pMSCV-IRES-GFP retroviral expression plasmid. The Mito-dsRed construct was purchased from Addgene (Cambridge, MA) (plasmid 11151). Mfn2 constructs were generated by subcloning the murine Mfn2 cDNA into the EcoRI/BamHI sites of the pLVX-EF1α-IRES-GFP or pLVX-EF1α-IRES-mCherry lentiviral expression plasmid (Clontech). The pGreenFire-Nfat and pGreenFire-CMV gene reporter constructs were purchased from System Biosciences (San Jose, CA) and contained three canonical Nfat response elements (5′- -3′) driving the expression of copGFP and luciferase reporters. The DNDrp1-pcDNA3.1 construct was purchased from Addgene (#45161) and subcloned using the BamHI/EcoRI restriction sites into the pLVX-IRES-GFP vector. Lentiviral 2nd generation packaging construct ΔR8.2 (8455) and pDM2.6 (12259) were purchased from Addgene. The −950/+22 murine MFN2 promoter was constructed by PCR amplification of the RP23-458J18 BAC clone (CHORI, Oakland, CA) and subcloned into the pGL4 luciferase reporter vector (Promega, Madison, WI). All cloning was carried out using KOD hot-start polymerase (Novagen, Billerica, MA) and subcloned for screening and sequencing into the pCR2.1 shuttle vector (Invitrogen, Carlsbad, CA). For peripheral blood analyses, erythrocytes were lysed twice with ACK lysis buffer and nucleated cells were stained with antibody cocktail (Supplementary Table 1) in FACS buffer for 15 min on ice, washed and analysed on a BD FACSCantoII flow cytometer (Becton Dickinson, Mountain View, CA). For bone marrow analyses, cells were isolated using the crushing method and erythrocytes were lysed with ACK lysis buffer followed by 40 μm filtration. bone marrow cells were stained with antibody cocktail in FACS buffer for 30 min on ice, washed and analysed on a BD LSRII flow cytometer (Becton Dickinson, Mountain View, CA). Dead cells were excluded from analyses by gating out 7AAD-positive cells. To isolate purified haematopoietic populations, bone marrow cells were isolated, stained and sorted using a BD Influx cell sorter (Becton Dickinson, Mountain View, CA) into complete media. Data were analysed using FlowJo9.6 (TreeStar Inc., Ashland, OR). Mfn2fl/fl-Vav-Cre fetal liver cells, bone marrow cells or purified LT-HSCs (Lin−cKit+Sca1+CD48−Flt3−CD150+) were transplanted into lethally irradiated (two doses of 478 cGy over 3 h using a Rad Source RS-2000 X-ray irradiator (Brentwood, TN)) recipients together with 2 × 105 competitor cells. As Mfn2fl/fl-Vav-Cre mice were not fully backcrossed onto the C57BL/6 background, recipient mice and competitor bone marrow cells were from the B6.Cg-Tg(Vav1-Cre)A2Kio/J;B6/126SF1 Mfn2tm3Dcc/Mmucd mixed background mouse strain crossed to B6.SJL-Ptprca Pep3b/BoyJ (CD45.1) to generate a CD45.1+CD45.2+ mixed background mouse. Competitor cells were T-cell depleted using MACS beads. For all competitive transplantation experiments, at least two independent transplants, each with at least 4 recipients per condition of genotype were performed, and result of all recipients pooled for statistical analysis. Power calculation was based on results of the first experiment. In limiting dilution assays, cohorts of recipients received 20 or 50 HSCs together with 2 × 105 competitor cells, allowing calculation of HSC frequency based on the number of non-repopulated mice (<1% donor contribution) using Poisson statistics 15 weeks after reconstitution. For Mfn2 KO single cell transplantation, LT-HSCs were sorted directly into complete media (StemPro34, 100 ng ml−1 SCF, 100 ng ml−1 TPO, 50 ng ml−1 IL-6) and single cells were visually confirmed. Positive single cell wells were combined with 2 × 105 CD45.1 competitor bone marrow cells and transplanted into lethally irradiated CD45.1 recipient mice. Recipients showing ≥ 0.1% CD45.2 donor contribution were considered positive and GM/(B+T) ratios were calculated as previously described for characterizing heterogeneous HSC phenotypes37. In transplantations using WT or Prdm16−/− HSCs (Lin−cKit+Sca1+CD48−Flt3−CD150+) B6.CD45.2 cells were mixed with 2 × 105 freshly isolated B6.CD45.1 bone marrow cells and injected via tail vein into lethally irradiated (two doses of 478 cGy over 3 h using a Rad Source RS-2000 X-ray irradiator (Brentwood, TN)) B6.CD45.1+CD45.2+ F1 hybrid recipients. After 8 to 15 weeks, peripheral blood (PB) and bone marrow were analysed. Lentiviral particles were produced by seeding 293 cells at 7 × 105 per cm2, or PlatE cells (Cell Biolabs, San Diego, CA), in Ultra Culture serum-free media (Lonza, Basel, Switzerland) overnight followed by transfection of each packaging and expression construct (1:1:1) using Trans-It 293 (Mirus, Madison, WI) for 2 h. Media were pooled after 36–48 h, clarified and concentrated by ultracentrifugation (100,000g), resuspended in StemPro-34 media and stored at −80 °C. Virus titre was calculated from transduction of NIH-3T3 fibroblasts serial dilutions of the viral preparation. Sorted LT-HSCs were transduced with ≥ 150 MOI lentivirus particles in the presence of 6 μg ml−1 polybrene (Sigma) and spun at 900g for 20 min at 20 °C. Supernatant was aspirated and replaced with complete media and cultured overnight. Transduction efficiency of cells was confirmed after 24 h. To assess proviral copy number 15 weeks post-transplantation in vivo, splenocytes were harvested and sorted into donor (CD45.2) or competitor (CD45.1) populations and gDNA was isolated as previously described38. Amplification of the proviral WPRE region was achieved using SYBR Green qPCR assay using the primer pair WPREFor: 5′- -3′ and WPRERev: 5′- -3′. Quantification of proviral copies was derived from the linear regression of serial dilutions of viral vector and normalized to input cell number. Sorted or cultured cell populations (2–5 × 103 cells) were lysed in TRIzol LS reagent (Invitrogen, Carlsbad, CA) and RNA was isolated according to manufacturer’s instructions. cDNA was synthesized using Superscript III Reverse Transcriptase (Invitrogen) and target CT values were determined using inventoried TaqMan probes (Applied Biosystems, Carlsbad, CA, see Supplementary Table 2) spanning exon/exon boundaries and detected using a Viia7 Real Time PCR System (Applied Biosystems). Relative quantification was calculated using the ΔΔC method. To estimate relative copy number of Mfn1 and Mfn2 transcripts (Fig. 4a), copy numbers were derived from the linear regression of serial dilutions of respective cDNA plasmids and normalized to GAPDH-VIC values. To estimate relative copy number of flPrdm16 transcripts (Fig. 4d), a probe was designed to span the SET methyltransferase domain of Prdm16 (exon2/3 junction) and copy number was derived from the linear regression of serial dilutions of respective cDNA plasmids. Another probe (exon 14/15 junction) was used to quantify total Prdm16 copy numbers derived from the linear regression of serial dilutions of respective cDNA plasmids. The values derived from total Prdm16 probe was subtracted from flPrdm16-specific probe to determine sPrdm16 transcript quantity. All values were normalized to relative multiplexed GAPDH-VIC values. Culture of sorted LT-HSCs was carried out using StemPro34 media (Invitrogen) supplemented with 10 mM HEPES and 50 ng ml−1 of recombinant murine SCF, TPO, IL-6 (Peptrotech, Rocky Hill, NJ) and cultured in 5% O at 37 °C. In some experiments, LT-HSCs were cultured in the presence of 500 ng ml−1 VIVIT (Millipore, Billerica, MA) or 30 μM mDivi1 (MolPort, Riga, Latvia). To demonstrate a mitochondrial fusion activity, cell fusion experiments were performed using MEFs as previously described37. Briefly, BacMam baculovirus constructs (Invitrogen) expressing the signalling peptide from cytochrome c fused to either GFP or RFP were transduced separately into MEF cells. Sorted GFP+ and RFP+ MEFs were co-cultured for 24 h and plasma membranes were fused using PEG-1500 (Roche, Basel, Switzerland. Fused cells were cultured in DMEM containing cyclohexamide (Sigma, St. Louis, MO) for 4 h and analysed for colocalization of mitochondrial labels. Early passage Prdm16−/− MEFs were transduced with 10 MOI retrovirus for 72 h and fixed with 4% paraformaldehyde for 10 min. Protein lysates were isolated and chromatin immunoprecipitation was carried out using the ChIP-IT Express Enzyme kit (Active Motif, Carlsbad, CA). Antibodies used for ChIP include anti-Flag and anti-TF2D. Primer probes were designed to span regions of the Mfn2 promoter previously shown to regulate Mfn2 transcriptional activity (see Supplementary Table 3)39. Quantification of precipitated Mfn2 promoter regions were derived from the linear regression of serial dilutions of bone marrow genomic DNA, normalized to input DNA concentration and quantifiable IgG detection was subtracted from sample values. Bone marrow was freshly isolated and lineage depleted with the MACS Lineage Depletion Kit (Miltenyi Biotech, San Diego, CA). Cells were cultured for 30 min in complete medium supplemented with 1 μM Indo-1 prepared as stock supplemented with Pluronic-F127 and incubated at 37 °C for 30 min. Cells were washed and stained for surface markers for 15 min, washed and allowed to rest in for 15 min PBS in PBS with Ca2+. FACS tubes were run at 37 °C in the sample port of the LSRII flow cytometer equipped with a 355 nm excitation laser. Events were collected for 40 s before incubation with 25 μM ATP or 1 μM SDF1 to induce calcium transients. The average ratio, R, of bound/free Indo-1 (405 nm/485 nm emission) before simulation was used to determine baseline values. Identical samples were equilibrated in 10 mM EGTA PBS without Ca2+ to determine R or stimulated with 1 μM ionomycin to determine R . The Indo-1 dissociation constant (K ) was assumed to be 237 nM at 37 °C based on previous studies40. The following equation was then used to relate Indo-1 intensity ratios to [Ca2+] levels; Sorted or cultured haematopoietic populations (2–5 × 103 cells) were collected in complete media and plated on onto MicroWell 96-well glass-bottom plates (Thermo, Waltham, MA) coated with 1ug ml−1 poly-d-lysine. Cells were allowed to adhere for 10 min and fixed with 4% PFA for 15 min. Cells were then permeabilized with 0.1% TritonX-100/PBS for 5 min and blocked with 2% BSA/PBS for 1 h at 4 °C. Cells were incubated with anti-Nfat1 (1:100), anti-Mfn2 (1:200), anti-tubulin (1:200), anti CD150-APC (1:100) or anti-Flag (1:250) (see Supplementary Table 1) overnight, washed and incubated with AlexaFluor secondary antibodies (Invitrogen) for 1 h. Cell nuclei were counterstained with DAPI and mounted with fluorescent mounting media (Vector Labs, Burlingame, CA). Confocal images were acquired with a Zeiss LSM 700 confocal microscope or a Leica DMI 6000B and images were deconvoluted and processed with Leica AF6000 software package. NIH-3T3, WT or Mfn2−/− MEF cells were plated at 2 × 104 cells per cm2 in triplicate overnight and transfected with 500 ng of pGF-Nfat, pGF-CMV or −950/+22 Mfn2-pGL4 reporter construct, 500 ng of cDNA plasmids as indicated and 500 ng of either pSV-βGal or pLVX-IRES-mCherry plasmids with Lipofectamine 3000 according to manufacturer’s instructions for 24 or 48 h. Cells were lysed in reporter lysis buffer (Promega, Madison, WI) and analysed for luciferase activity using BrightGlo luciferase (Promega) and detected on a Synergy H2 plate reader (BioTek, Winooski, VT). To visualize βGal activity, cell lysate was incubated in Buffer Z (1mg ml−1 ONPG, 0.1 M phosphate, pH 7.5, 10 mM KCl, 1 mM βME, 1 mM MgSO ) at 37 °C for 1 h. Absorbance values were measured at 405 nm and used to normalize for transfection efficiency. In WT and Prdm16−/− MEFs, gene reporter luciferase values were normalized to mCherry excitation values. For total cell lysate experiments, MEF cultures were lysed in RIPA buffer, 50 mM Tris pH 7.5, 137 mM NaCl, 0.1% SDS, 0.5% deoxycholate and protease inhibitors (Roche). For subcellular fractionation studies, cells were scraped, washed in PBS. Cell pellets were lysed in 5× packed cell volume (pcv) Buffer A for 10 min on ice and vortexed for 15 s in the presence of 1/10 volume 3% NP-40. Plasma membrane lysis was verified by trypan blue staining. Lysate was spun at 15,000g for 10 min at 4 °C and the cytoplasmic fraction was saved. The remaining nuclear pellet was resuspended in 2.5× pc Buffer C and incubated at 4 °C for 1 h with rotation and spun at 15,000g for 10 min. The nuclear fraction was diluted with 2.5× volume of Nuclear Diluent Buffer and stored at −80 °C. To achieve even fractionation loading, equivalent percentages of nuclear and cytoplasmic fractions were loaded on each gel. All protein samples were denatured in 4× sample buffer at 95 °C and loaded onto 4–12% Bis-Tris SDS–PAGE gradient gels (Invitrogen). Gels were transferred onto 0.22 μm nitrocellulose membrane and stained with Ruby Red (Molecular Probes, Carlsbad, CA) to confirm transfer. Membranes were blocked with 3% non-fat milk or BSA in 0.1%Tween-20/TBS and incubated with anti-Mfn2 (1:200), anti-βGal (1:1,000), anti-Nfat1 (1:250), anti-tubulin (1:1,000), anti-lamin A/C (1:500) and anti-β-actin (1:5,000) overnight (see Supplementary Table 1). Membranes were washed, incubated with HRPO-conjugated secondary antibodies and exposed to X-ray film (Denville) after incubation with Super Signal West Fempto ECL reagent (Pierce). For mitochondrial length measurements, confocal or deconvoluted z-stacks were collected and projected as a z-project in ImageJ (NIH, Bethesda, MD). Individual mitochondria were manually traced, binned into length categories and expressed as percent of cellular mitochondria. The mean ± s.e.m. number of mitochondria falling into each length category collected from ≥ 15 fields (30–50 cells) are expressed. For Nfat nuclear localization quantification, confocal or deconvoluted z-stacks were collected and a 1-μm section in the centre of the cell was projected as a z-project in ImageJ. Nuclear boundaries were constructed using DAPI staining. The ratio of staining within the nuclear boundary to total staining was expressed as percent of Nfat signal. The mean ± s.e.m. for ≥ 10 fields (20–40 cells) are expressed. For immunofluorescence intensity measurements, confocal or deconvoluted z-stacks were collected and projected as a z-project in ImageJ. Thresholds were set based on IgG-stained negative control cells and the integrated density value of each signal per cell was recorded. The mean ± s.e.m. for ≥ 15 fields (30–50 cells) are expressed. For statistical analysis between two groups, the unpaired Student’s t-test was used. When more than two groups were compared, one-way ANOVA was used. Results are expressed as mean ± s.e.m. The Bonferroni and Dunnett multiple comparison tests were used for post-hoc analysis to determine statistical significance between multiple groups. All statistics were calculated using Prism5 (GraphPad, La Jolla, CA) software. Differences among group means were considered significant when the probability value, P, was less than 0.05. Sample size (‘n’) always represents biological replicates. Cochran test was used for exclusion of outliers. 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.

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Tissue samples from a Hungarian mummy have revealed that people in the early 17th and 18th centuries suffered from colon cancer, long before the modern plagues of obesity, physical inactivity and processed food were established as causes of the disease, according to new research. In a new study of 18th-century Hungarian mummies, scientists found that the genetic predisposition to colon cancer predates modern impacts on health. One of the mummies in the study carried a mutation in the adenomatous polyposis coli (APC) gene, which physicians now know raises the risk of colon cancer, said lead study author Michal Feldman, a research assistant formerly at Tel Aviv University in Israel. If the APC mutation is confirmed in other samples, it could mean that inherited changes in DNA play a bigger role in cancer evolution than do modern environmental impacts, Feldman told Live Science in an email. [10 Do's and Don'ts to Reduce Your Risk of Cancer] "Today, colorectal cancer is the third most common type of cancer, and it has a clear genetic background that is well-researched in modern populations," Feldman said. "In light of the many lifestyle and environmental changes human society has undergone during the last few centuries, we found it important to compare the spectrum of historical mutations to the modern spectrum." Because mummification preserves tissue, samples from such remains can give scientists invaluable information on anthropological, historical and medical details, Feldman said. In the past, studies of mummified remains have provided clues about the history of tuberculosis, clogged arteries and even air pollution. In the new study, Feldman's team collected tissue samples from 20 mummies that were excavated from sealed crypts in a Dominican church in Vác, Hungary. These crypts were used for the burial of several middle-class families and clerics from 1731 to 1838, and more than 265 mummies were found there in 1995, the researchers said. The mummies are now housed at the Hungarian National Museum in Budapest. The low temperature in the crypts, combined with constant ventilation and low humidity, were ideal conditions for natural mummification of the corpses, the researchers said. Some 70 percent of the bodies found in the location were completely or partially mummified, providing a rich source of preserved tissue and DNA samples for the scientists. [8 Grisly Archaeological Discoveries] By extracting DNA from the mummies, Feldman and her team were able to sequence and assess the presence of APC gene mutations. "The interesting thing about this study is that the APC mutation in cancer that was recently discovered in the past couple of decades is not new," said Dr. Sidney Winawer, a gastroenterologist at Memorial Sloan Kettering Cancer Center in New York, who was not involved in the study. "This opens up a whole new way of thinking. If this mutation was present so many years ago, why was it present there?" Additional historical samples need to be investigated, he said, in order to better understand the relationships between cancer and environmental factors, such as lifestyle, and between cancer and genetic changes. The findings were published online Feb. 10 in the journal PLOS ONE. Copyright 2016 LiveScience, a Purch company. All rights reserved. This material may not be published, broadcast, rewritten or redistributed.

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Panama's president Juan Carlos Varela (C) talks to Canal Administrator, Jorge Quijano during their visit to the new set of locks at the Panama Canal Expansion project on the pacific side in Panama City March 18, 2016. A view of the new set of locks of The Panama Canal Expansion project on the Pacific side in Panama City March 11, 2016. Panama is aiming to inaugurate its newly expanded canal at the end of June according to local media. PANAMA CITY The Panama Canal will next month impose new draft restrictions on ships due to falling water levels at nearby lakes that form part of the waterway, the authority that administers the canal said in a statement on Monday. Ships seeking to cross the waterway must comply with a maximum depth limit of 39 feet (11.89 meters) beginning on April 18, but the Panama Canal Authority (APC) said the impact on operations would be minimal. The "temporary and preventive measures" are connected to local climate impacts of El Niño, the seasonal weather phenomenon that has caused a drought in the canal's watershed, and will be implemented in 6-inch (15-cm) decrements that will be announced at least four weeks in advance. The APC added that ships loaded after March 21 will need to comply with the new restrictions.

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