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In 3 article(s): Set(16756390.xml, 17280616.xml, 17608567.xml)
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To determine the functional interactions of RCK/p54 with miRISC, we employed affinity purification of RISC and target mRNA cleavage capabilities of miRISC when the target has perfectly complementary sequences to the miRNAs.

To dissect and understand the relationship between RNAi function and P-bodies, we affinity-purified RISC using Myc-Ago2 and expression vectors of the YFP-tagged P-body proteins, Lsm1, RCK/p54, Dcp2, and eIF4E. Ago2 interacted with these various P-body components in ways that were RNA-dependent or RNA-independent (Figure 1A).

(A) Affinity-purified miRISCs associated with PCK/p54 retain cleavage activity. To purify miRISC associated with RCK/p54, magnetic protein A beads coupled with rabbit IgG, rabbit anti-Ago2, or rabbit anti-RCK/p54 antibodies were incubated with HeLa cytoplasmic extracts. After immunoprecipitation, RISC activities were analyzed by incubating the supernatant (S) or bead (B) phases with 182-nt 32P-cap-labeled let-7 substrate mRNAs having a perfectly complementary or mismatched sequence to the let-7 miRNA. Cleavage products were resolved on 6% denaturing polyacrylamide gels. CE, cytoplasmic extract; PM, perfect match; MM, mismatch.

The glutathione S-transferase (GST)-Bcl-GL recombinant protein was expressed in Escherichia coli strain BL21 codon-plus RIL competent cells (Stratagene). Purification of the recombinant proteins was performed using Glutathione Sepharose 4B beads (GE Healthcare) under nondenaturing conditions according to the supplier's instructions. For confirmation of direct binding of BCL-GL and MELK, we removed GST from GST-fused BCL-GL protein using PreScission protease (GE Healthcare) according to the supplier's instructions.

Soluble fractions were filtered with 0.8 μm syringe filters and applied into a Ni-NTA affinity column pre-equilibrated with 30 mM Tris-HCl (pH 8.0), 50 mM NaCl, 5 mM β-mercaptoethanol. Target protein complexes (the Aα subunit with GST-tag and SV40 ST with His-tag) were eluted with elution buffer (30 mM Tris-HCl [pH 8.0], 50 mM NaCl, 300 mM imidazole, 5 mM β-mercaptoethanol) and dialyzed overnight at 4 °C in 30 mM Tris-HCl (pH 8.0), 50 mM NaCl, 5 mM DTT. Dialyzed protein was applied to a GST affinity column to remove free SV40 ST, and on-column cleavage with TEV protease was performed at 4 °C overnight. The flow-through fraction of the GST column was reapplied into the Ni-NTA column to remove cleaved His-tag and TEV protease.


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In 8 article(s): Set(17177603.xml, 17511879.xml, 16968134.xml, 17341134.xml, 16513846.xml, 18781224.xml, 16754960.xml, 17620405.xml)
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The MACS epitope tagged protein isolation kit (Miltenyi Biotec) was used to precipitate candidate partner proteins via the HA epitope. Eluted protein samples were split (1/3 and 2/3), size-fractionated in parallel on two polyacrylamide SDS-gels (concentration adjusted to protein size), electroblotted onto Immobilon-P membrane (Millipore) and subjected to antibody detection. HA-tagged (1/3 aliquot) partner proteins were identified using an anti-HA horseradish peroxidase (HRP)-conjugated antibody (clone 3F10; Roche 2012819). The co-precipitated 6× his-tagged STM protein was visualized by a primary penta-His mouse antibody [α-(H)5; Qiagen 34660] and secondary HRP-coupled goat anti-mouse IGG (Dianova 115-035-062). Peroxidase activity was detected non-radioactively via chemiluminescence (ECL PLUS kit; Amersham Biosciences) and documented on Kodak-X-omat AR films. Epidermal proteins after bombardement of leek epidermal cells were isolated in 50 mM Tris–HCl, pH 8.0, 150 mM NaCl, 1% Triton and size-fractionated by SDS–PAGE. GFP fusion proteins were detected after transfer to Immobilon-P membrane (Millipore) by a mouse monoclonal anti-GFP HRP-conjugated antibody (IgG1; Milteny Biotec 130-091-833).

(A) Nontransfected MDA-MB-231 cells were surface labeled with cleavable biotin and lysed immediately or allowed to internalize cell surface proteins for 15 min. Immunoprecipitations (IPs) were performed as indicated, and the Rab coprecipitating proteins were detected first with anti-biotin antibody followed by stripping and reprobing with anti–β1-integrin antibody. The quantification shows the amount of coprecipitated biotinylated protein relative to total precipitated integrin (means ± range; n = 2).

In a converse experiment, ephrin-B1 was co-immunoprecipitated with an anti-Cx43 antibody (Figure 6Cb).

Cells were lysed in 50 mM HEPES (pH 7.2), 150 mM NaCl, 1mM EDTA, 0.2% NP-40, and complete protease inhibitors. Cell lysates were resolved by standard Laemmli's SDS-PAGE (pH 8.8) unless otherwise stated. For immunoprecipitations: rat Bim antibody (Oncogene, San Diego, California, United States) was coated to goat-anti-rat immunoglobulin-agarose; rabbit Puma antibody was coated to protein A-sepharose; mouse NHE-1 antibody was coated to protein G-sepharose; rabbit Bcl-xL antibody was coated to goat-anti-rabbit immunoglobulin-agarose. Lysates were precleared with the appropriate agarose.

To explore the potential interplay between Trk receptors and Cdk5, we first examined if Trk receptors associated with Cdk5 or p35. TrkA, TrkB, or TrkC was overexpressed together with Cdk5 or p35 in COS7 cells, and immunoprecipitation was performed with Cdk5, p35, or pan-Trk antibody. Interestingly, all three Trk receptors were observed to associate with Cdk5 (Figure 1B) and p35 (Figure 1C), while no association was observed when immunoprecipitation was performed with IgG control. Since both TrkB and its ligand BDNF are abundantly expressed in the brain throughout development, we next proceeded to verify the interaction between TrkB and Cdk5/p35 in postnatal brains. We found that TrkB associated with both p35 and Cdk5 in postnatal day 7 (P7) rat brain lysates (Figure 1D).

TrkA, TrkB, and TrkC were overexpressed in COS7 cells and immunoprecipitated by pan-Trk antibody. Incubation with Cdk5/p25 revealed that TrkB and TrkC, but not TrkA, were phosphorylated by Cdk5/p25 in vitro (Figure 2A). This is in agreement with the lack of Cdk5 consensus sites in TrkA, and points to the possibility that Cdk5 may phosphorylate TrkB and TrkC at the Cdk5 consensus sites at the juxtamembrane region (Figure 1A). To examine this possibility, a GST fusion protein containing only the juxtamembrane region of TrkB was prepared.

For immunoprecipitation, 1–2 mg of protein lysates was incubated with 1 μg of the corresponding antibody at 4 °C overnight with rotation. Forty microliters of protein G Sepharose (Amersham Biosciences) pre-washed with 1× PBS was added and rotated at 4 °C for 1 h. After intense washing with the lysis buffer, the immunoprecipitated protein and its associated proteins were analyzed by SDS-PAGE and Western blotting.

(B) Cell lysates from HEK293T cells overexpressing Cdk5 and TrkA, TrkB, or TrkC were immunoprecipitated (IP) with Cdk5 antibody and immunoblotted with pan-Trk antibody. TrkA, TrkB, and TrkC were all observed to associate with Cdk5.

(C) Cell lysates from HEK293T cells overexpressing p35 and TrkA, TrkB, or TrkC were immunoprecipitated with p35 antibody and immunoblotted with pan-Trk antibody. TrkA, TrkB, and TrkC were all observed to associate with p35.

(D) Brain lysate from P7 rat brain was immunoprecipitated with pan-Trk, p35, or Cdk5 antibody and immunoblotted with p35, Cdk5, and TrkB antibodies. Rabbit normal IgG was used as a control. TrkB was observed to associate with both p35 and Cdk5 in P7 rat brain.

(F) Brain lysates from P7 p35+/+ or p35−/− mouse brains were immunoprecipitated with p35 and Cdk5 antibodies and immunoblotted with p35, Cdk5, and TrkB antibodies. Rabbit normal IgG served as a control. Association between Cdk5 and TrkB was abolished in p35−/− brain, indicating that p35 was required for the association between Cdk5 and TrkB.

(A) Lysates from COS7 cells overexpressing TrkA, TrkB, and TrkC were immunoprecipitated with pan-Trk antibody and incubated with Cdk5/p25 in an in vitro kinase assay. TrkB and TrkC, but not TrkA, were phosphorylated by Cdk5/p25.

(E) Full-length TrkB WT, M1, M2, and DM were overexpressed with or without Cdk5/p35 in HEK293T cells. In the absence of Cdk5/p35, Ser478-phosphorylated TrkB (p-Ser TrkB) was not detected. Overexpression of Cdk5/p35 resulted in phosphorylation of TrkB WT at Ser478, but phosphorylation at Ser478 was essentially abolished when TrkB M1 and DM were overexpressed. IP, immunoprecipitation.

(A) Cortical neurons were stimulated with BDNF for different time intervals. Lysates were immunoprecipitated (IP) with p35 antibody and subjected to in vitro kinase assay using histone H1 as substrate. BDNF stimulation for 15 min resulted in a marked increase in Cdk5 activity in cortical neurons. Quantification of the changes in phospho-Histone H1 level following BDNF stimulation was normalized to the value obtained from untreated cultures (time 0) and is shown in the histogram. *, p < 0.05.

(B) Addition of Trk inhibitor K252a abolished BDNF-induced increase in Cdk5 activity. Cortical neurons were pretreated with vehicle control (DMSO) or K252a for 30 min before stimulation with BDNF for 15 min. Lysates were immunoprecipitated with p35 antibody and subjected to in vitro kinase assay using histone H1 as substrate. We found that K252a pretreatment markedly reduced the increase in Cdk5 activity triggered by BDNF stimulation, indicating that the induction of Cdk5 activity was dependent on TrkB activation. Quantification of the changes in phospho-Histone H1 level following BDNF stimulation in the presence or absence of K252a treatment was normalized to the value obtained from untreated cultures (time 0) and is shown in the histogram. *, p < 0.05.

(C) Cortical neurons were treated with BDNF for 20 min. Lysates were immunoprecipitated with p35 antibody and immunoblotted with TrkB, p35, or Cdk5 antibody. While association between Cdk5 and p35 was not affected by BDNF stimulation, association between p35 and TrkB increased following 20 min of BDNF stimulation.

GST-pull down and co-immunoprecipitation assays were performed to further characterize this interaction.

Coimmunoprecipitation of BRCA1 and PP1. HEK293T kidney cells were transfected with vectors encoding untagged BRCA1 under the control of a CMV promoter, and vectors encoding Flag-PP1α, β, γ or Flag-Laf4. (A) A western blot probed with BRCA1 shows that immunoprecipitation of protein with an antibody against the Flag-PP1α, β or γ proteins, but not Laf4, co-immunoprecipitates BRCA1 (Lanes 1A-1D). It should be noted that the band observed slightly lower than BRCA1 in lane 1D is a non-specific background band. Lanes 1E-1H show immunoprecipitation of BRCA1 using antibodies against the amino and carboxy termini of BRCA1. (B) A western blot probed with an antibody against the Flag epitope. Lanes 2A-2D indicate immunoprecipitation of the Flag-epitope tagged PP1α, β or γ or Flag-Laf4. Lanes 2E-2G show co-immunoprecipitation of Flag-PP1α, β or γ with antibodies against BRCA1, and lane 2H shows a lack of coimmunoprecipitation of the negative control Flag-Laf4 by BRCA1.

UXT was previously reported to be expressed almost exclusively inside the nucleus of most cells (Markus et al., 2002). This was confirmed in our investigation for either endogenous or overexpressed UXT (Fig. 1 B). To further substantiate its interaction with p65, an in vitro coimmunoprecipitation assay was applied in which full-length HA-p65 and FLAG-UXT proteins were generated and labeled, respectively, with [35S]methionine by in vitro translation. The products were mixed and immunoprecipitated with either control IgG or anti-HA antibody. As shown in Fig. 1 C, UXT could be coprecipitated by antibody against the HA epitope but not by control IgG, which suggests that UXT indeed interacts directly with full-length p65.

To address the physiological relevance of this interaction in mammalian cells, we expressed HA-UXT in 293T cells and then stimulated cells with or without TNF-α for the indicated times. The fractionated cytoplasmic or nuclear extracts were immunoprecipitated with either anti-p65 antibody or IgG as a control, respectively. There was no detectable UXT that interacted with cytoplasmic p65 in the presence or absence of TNF-α (Fig. 1 D), which was consistent with the unique subcellular location of UXT. In addition, there was only a marginal amount of endogenous p65 in the nucleus devoid of TNF-α treatment. Consequently, no UXT was coimmunoprecipitated from this nuclear extract even though there existed a large amount of UXT. In contrast, there exhibited a strong interaction between nuclear p65 and UXT upon TNF-α stimulation. Furthermore, we tested whether endogenous UXT and p65 could interact in response to TNF-α. As shown in Fig. 1 E, endogenous UXT was coimmunoprecipitated by p65 antibody from cells treated with TNF-α. In contrast, UXT was barely detected in the immunoprecipitates without TNF-α treatment. One possible explanation for this phenomenon is that only after p65 translocation into the nucleus could UXT have access to p65. However, we could not formally rule out the possibility that posttranslational modifications of either protein were prerequisites for this interaction in vivo. Collectively, these results indicate that UXT interacts in vivo with p65 upon TNF-α stimulation.

We sought to establish whether the mitochondrial localized SIRT3 may also form a physical interaction with the FOXO family protein, FOXO3a, using co-immunoprecipitation (Co-IP) techniques. Carboxy-terminally myc tagged wild-type (p-myc-hSIRT3-wt), and mutant (p-myc-hSIRT3-mt) SIRT3 expression vectors were transfected into Cos-7 cells followed by Co-IP with an anti-myc antibody.

(A) FOXO3a binds to SIRT3 in vitro. Cos-7 cells were transfected with either SIRT3 wild-type (p-myc-hSIRT3-wt) or deacetylation mutant (p-myc-hSIRT3-mt) vectors and cell lysates were immunoprecipitated (IPd) with an anti-Myc antibody followed by Western analysis with an anti-FOXO3a antibody. (B) HCT116 cell lysates were IPd with either an anti-FOXO3a or anti-SIRT3 antibody, resolved by SDS-PAGE, and immunoblotted with anti-FOXO3a antibody. (C) Mitochondrial factions from HCT116 cells were IPd with either an anti-FOXO3a or anti-SIRT3 antibody and immunoblotted with anti-FOXO3a antibody.

Cells were fixed with 1% formaldehyde to crosslink protein-DNA interactions, sonicated, and fixed cells were immunoprecipitated with either an anti-FOXO3a antibody.


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In 7 article(s): Set(17511879.xml, 16756390.xml, 18836139.xml, 17280616.xml, 16513846.xml, 18188154.xml, 19131970.xml)
In 12 passage(s)
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In addition to the interactions in yeast, we performed co-immunoprecipitation experiments with epitope-tagged full-length proteins. The HA-tagged BLH proteins shown in Figure 2B were used to co-precipitate epitope-tagged STM-His (Figure 2C). The detection of the STM-His protein was strictly dependent on the presence of BLH partner proteins. The co-immunoprecipitation experiments therefore confirm that the full-length STM protein interacts with full-length ATH1, BLH3 and BLH9 proteins in vitro and substantiate the affinity of BLH/STM interactions.

To examine the RNA dependence of protein–protein interactions, TCEs (250 μg) were treated before immunoprecipitation with 0.2 μg/ul of RNase A for 20 min at room temperature. Myc-tagged proteins were precipitated by incubating overnight with polyclonal rabbit anti-Myc antibodies directly conjugated to agarose beads (Santa Cruz Biotech, California, United States).

HeLa cells were transiently co-transfected for 48 hours with 8 μg of plasmid constructs encoding Flag-tagged full-length Bcl-GL or a series of Flag-tagged partial Bcl-GL proteins (FL, N-1, N-2, N-3, N-4, C-1, C-2 and C-3) as well as the same amount of plasmid encoding HA-tagged WT-MELK, using the FuGENE6 transfection reagent (Roche). Cells were lysed with lysis buffer as described above. The lysates were pre-cleaned with normal mouse IgG (1.2 μg) and rec-Protein G Sepharose 4B (Zymed, San Francisco, CA, USA) at 4°C for 30 minutes. Subsequently, the lysate was incubated with anti-Flag agarose M2 gel (Sigma-Aldrich) at 4°C for 12 hours. After washing three times with lysis buffer, proteins on beads were eluted with SDS sample buffer.

To validate an interaction between WT-MELK and Bcl-GL, we constructed plasmids designed to express HA-tagged WT-MELK (HA-WT-MELK) and Flag-tagged Bcl-GL (Flag-Bcl-GL). These plasmids were co-transfected into HeLa cells and the proteins immunoprecipitated with anti-Flag antibody. Immunoblotting of the precipitates using anti-HA antibodies indicated that Flag-Bcl-GL was co-precipitated with HA-WT-MELK (Figure 3c).

To further determine which segment of Bcl-GL can interact with WT-MELK, we performed co-immunoprecipitation analyses using HA-WT-MELK and partial Bcl-GL proteins tagged with Flag (Figure 3e). After co-transfection of plasmid clones into HeLa cells, we performed immunoprecipitation with an anti-Flag antibody, and then immunoblotting with an anti-HA antibody.

(c) Interaction of MELK with Bcl-GL. Extracts from HeLa-cells transfected with HA (hemagglutinin)-tagged WT-MELK (HA-WT-MELK) or Flag-tagged Bcl-GL (Flag-Bcl-GL), or a combination of these, were harvested 36 hours after transfection. The cell lysates were immunoprecipitated with anti-Flag M2 antibody. Precipitated proteins were separated by SDS-PAGE and western blotting analysis was performed with an anti-HA antibody.

(f) Determination of the WT-MELK binding regions of Bcl-GL by immunoprecipitation. The HA-tagged WT-MELK and various peptide sequences of Flag-tagged Bcl-GL (Figure 3e) were pulled down by immunoprecipitation with Flag-M2 antibody and then immunoblotted with rabbit anti-Flag antibody. The expression of HA-tagged WT-MELK in total cell lysates was confirmed by western blotting analysis. As a control, immunoprecipitation was performed from cells co-transfected with pCAGGSn3FC (Mock) and HA-tagged WT-MELK (HA-WT-MELK) through all steps.

PP1 has 3 isoforms encoded by different genes that are 97% conserved across their catalytic domains and distinct roles for each isoform have yet to be determined. When we coimmunoprecipitated Flag-epitope tagged PP1α, β or γ with BRCA1, we observed that all 3 isoforms interacted with BRCA1. Additionally, we have identified the functional PP1 interacting domain within BRCA1. This domain is found in other PP1 regulatory proteins, suggesting that BRCA1 may regulate the activity of PP1 and could act as a scaffold protein to promote the dephosphorylation of BRCA1 associated proteins by PP1.

To probe for a possible signaling complex including the β1AR and a PDE, mouse neonatal cardiomyocytes were infected with an adenovirus encoding a Flag-tagged β1AR, and the receptor was subsequently immunoprecipitated using an antibody against the tag. A significant amount of endogenous PDE activity was recovered in the β1AR immunoprecipitation (IP) pellet (Figure 1A).

(B) Co-IP of β1AR and PDE activity from cardiomyocytes deficient in PDE4A, PDE4B, or PDE4D, and wild-type controls.

(A, B) Co-IP of exogenous β1AR and Myc-tagged PDE4D splice variants expressed in HEK293 cells. The efficiency with which β1AR pulls down the different PDE4D splice variants is quantified in (B). (C) Shown is the co-IP of exogenous β1AR and PDE4D8-Myc from extracts of MEFs derived from mice deficient in β-arrestin 1 and 2 (βarr1/2KO) or from wild-type controls (WT-MEF). (D, E) PDE4D3, and Flag-tagged receptors, β1AR and β2AR, were affinity purified after baculovirus expression (see Supplementary Figure 1). Purified PDE and (βARs) were then combined and the βARs immunoprecipitated.

(A, B) Neonatal cardiac myocytes expressing a Flag-tagged β1AR were treated for 3 min with 100 nM Norepinephrine before cell lysis and IP with M1 (α-Flag) resin. The effect of a 5 min pre-treatment with 10 μM of the PDE4-specific inhibitor, Rolipram, or the PDE3-selective inhibitor, Cilostamide, on PKA-phosphorylation of the β1AR is detected in IBs using a PKA-site-specific antibody. (C, D) MEFs derived from mice deficient in PDE4D or wild-type controls were infected with adenovirus to express a Flag-tagged β1AR construct. At 40 h post-infection, cells were treated for 3 min with 100 nM Norepinephrine (NorEpi) before cell lysis and IP with M1 (α-Flag) resin. PKA-phosphorylation of the β1AR is detected in IB using a PKA-site-specific antibody. (E, F) Neonatal cardiac myocytes coexpressing a Flag-tagged β1AR and either GFP, a catalytically inactive PDE4D8 construct (PDE4D-DN; see also Supplementary Figure 5), or a catalytically inactive PDE3A1 (PDE3A1-DN) were subjected to α-Flag(M1)-IP, and the phosphorylation of the β1AR was subsequently detected in IB using a PKA-substrate-specific antibody.

Co-immunoprecipitation assays were used to confirm the protein–protein interactions. Subsequently, localization of ACBP4 and its interacting protein, AtEBP, was confirmed using transient expression of GFP- and DsRed-tagged fusion proteins in Nicotiana tabacum.

The abnormal cross-linking of basal transcription factors in ctk1Δ cells is confirmed by independent immunoprecipitation of TAP-tagged TFIIF subunits.


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Importantly, upon stimulation with the adenylyl cyclase activator, Forskolin, all PDE4D isoforms show the same increase in activity in both cell types (Figure 5C), suggesting that loss of one βAR subtype or the other has not perturbed overall cAMP signaling. It also demonstrates that the spatial dimension of cAMP signaling is lost when generalized adenylyl cyclase activation is induced with Forskolin.

DLK1 EGF repeats bind Notch1 EGF repeats in bacterial two-hybrid assays and inhibit activity of a Notch-dependent reporter gene.


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The greater binding affinity of AIRE–PHD1 for H3K4me0 peptides was confirmed by both tryptophan fluorescence spectroscopy and isothermal titration calorimetry (ITC), yielding dissociation constants of ∼4 μM, ∼20 μM and >0.5 mM for H3K4me0, H3K4me1 and H3K4me2, respectively (supplementary Fig S2C online; Table 1).

In agreement with the GST fusion pull-down experiments, fluorescence spectroscopy showed no binding of H3K4me0 to AIRE–PHD1 containing the APECED-causing C311Y mutation (Bjorses et al, 2000). Nevertheless, a second pathological mutant, V301M (Soderbergh et al, 2000), was still able to bind to H3K4me0, indicating that this mutation is not located in the H3 interaction site (Table 1).

Indeed, fluorescence spectroscopy and ITC assays showed that the alanine mutations R2A in the H3 peptide and D312A in AIRE–PHD1 markedly reduced the binding affinity (Table 1; Fig 4C) without affecting the protein fold (supplementary Fig S3 online).

Similarly, pull-down experiments with whole histones and the H3K4me0 peptide, together with fluorescence spectroscopy and ITC measurements performed on AIRE–PHD1-D297A showed reduced binding (Table 1; Fig 4).


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In 10 article(s): Set(18388858.xml, 18724936.xml, 17511879.xml, 17112379.xml, 18836139.xml, 16513846.xml, 16754960.xml, 18700817.xml, 18775702.xml, 17620405.xml)
In 52 passage(s)
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In a yeast two-hybrid screen performed with a meristem-enriched cDNA library, three interacting BLH (Bel1-like homeodomain) transcription factors were identified.

We initiated a search for potential STM interaction partner genes starting from a yeast two-hybrid screen performed with the MEINOXSTM domain as bait for a meristem-enriched cDNA library.

The Matchmaker GAL4 system (Clontech) was used to perform the yeast two-hybrid screen.

As a prerequisite to substantiate interactions between potential partner proteins identified in a yeast two-hybrid screen, C- and N-terminal fusions between the ORFs of STM and the GFP were constructed and expressed in leek or onion epidermal cells after particle bombardment.

Potential STM protein interaction partners were identified in a yeast two-hybrid screen performed with a meristem-enriched cDNA library (Materials and Methods) established in the vector pACT2 (prey) and the MEINOXSTM domain as a bait expressed from the pGDKT7 vector.

In contrast to bombardments with the ATH1 and BLH3 constructs, the number of BiFC-positive cells obtained with the BLH9/STM combination (e.g. BLH9-NYFP/STM-CYFP in Figure 4C) was generally low. This may indicate a weak interaction between STM and BLH9 as suggested by the yeast two-hybrid results, that chimeric BLH9-NYFP protein levels remain low in leak epidermal cells, or that steric constraints interfere with a reconstitution of YFP fluorescence.

To identify novel integrin-interacting proteins, we screened a mouse embryonic cDNA library using α2-integrin cytoplasmic domain as bait in a yeast two-hybrid screen. The bait comprised the conserved membrane-proximal sequence shared by most α-integrin subunits followed by the α2-specific segment. Several positive clones encoded the COOH-terminal part of Rab21. Rab21 is a ubiquitously expressed and poorly characterized member of the Rab family that has recently been shown to function on the endocytic pathway (Simpson et al., 2004).

To get evidence for the association with an independent method, we performed yeast two-hybrid studies with α-tail mutants. In remating tests, we found that the COOH-terminal part of Rab21 (amino acids 95–222) was able to associate with the cytoplasmic tails of α2- and α11-integrin (Fig. 1 D).

Mutagenesis of residue R1161 (in α2AA and α2AAKYA, removing another conserved charged residue) to alanine significantly reduced α2 tail association with Rab21 judged by yeast mating tests and immunoprecipitations (Fig. 1, D and E). In addition, F1159A mutation (α2AARA, possibly creating a conformational change) showed reduced association in the yeast assays, whereas in immunoprecipitations the reduction was not significant. These data on the mutant integrins suggest that the conformation of the cytoplasmic domain and residue R1161 of α2-integrin are important for the Rab21 association.

(D) α2- and α11-integrin cytoplasmic domains and their point mutants (indicated in bold) were cloned as Gal4BD fusions and used as baits with Rab21 (amino acids 95–222) Gal4AD prey in yeast two-hybrid assays.

Our data does not unambiguously show whether integrins and Rabs interact directly. The facts that the association was detected from a yeast two-hybrid screen and that it is abrogated by mutagenesis of the α-cytoplasmic domain suggest that it may be direct.

Here we show that Abp1 interacts with the DNA replication protein Cdc23 (MCM10) in a two-hybrid assay, and that the Δabp1 mutant displays a synthetic phenotype with a cdc23 temperature-sensitive mutant.

Two-hybrid analysis also shows that Cdc23 (Mcm10) can interact with Orc1, 2, 5, and 6 [28,36].

Using a two-hybrid system, we have identified Abp1 as a protein that interacts with Cdc23 (Mcm10).

To provide additional insights into how Cdc23 (Mcm10) might function during DNA replication initiation, we conducted a two-hybrid interaction screen to identify cDNAs encoding proteins that interact with Cdc23 (Mcm10). One of the proteins identified several times in our screen was ARS-binding protein 1 (Abp1), a protein previously shown to interact with both replication origins and centromere-associated DNA sequence elements. Cdc23 (Mcm10) fused to the DNA binding domain of Gal4 was able to activate lacZ expression from the GAL1 promoter when co-expressed with Abp1 fused to the Gal4 activation domain (Fig. 1A, row 1). As a negative control, when Abp1 was replaced with either Skb1 or Snf4 no LacZ expression was observed (Figure 1A, rows 2 and 3). The two-hybrid interaction between Snf1 and Snf4 is shown as a positive control (Figure 1A, row 4). These two transcription factors have been previously shown to interact using the two-hybrid assay (Durfee et al, 1993).

Cdc23 physically interacts with Abp1 in a yeast two-hybrid cDNA library screen. Row 1: Cdc23-Abp1 interaction; rows2, 3: negative controls; row 4: Snf1/Snf4 positive control. (A). β-galactosidase assay. (B). HIS3 expression in the presence of 50 mM 3-AT. All experiments shown in triplicate.

Consistent with the lacZ data, Cdc23 (Mcm10) fused to the DNA binding domain of Gal4 activated HIS3 under the control of the Gal4 promoter when co-expressed with Abp1 fused to the Gal4 activation domain (Figure 1B, row 1). Co-expression with either Skb1 or Snf4 failed to activate HIS3 (Figure 1B, row 2 and 3) suggesting that the interaction of Cdc23 and Abp1 in the two-hybrid system is specific. As expected, the positive control, Snf1/Snf4 was also able to confer HIS prototrophy (Figure 1B, row 4).

We found that the double mutant cdc23-M36 Δabp1 is less viable then either cdc23-M36 or Δabp1 when grown at 30°C (Figure 3A, lower panel, row 4), consistent with our two-hybrid data suggesting that Cdc23 (Mcm10) interacts directly with Abp1.

In an attempt to gain some insights into the function of Cdc23, we conducted a two-hybrid screen for Cdc23-interacting proteins. One of the proteins identified using this screen is Abp1, a protein that had previously been shown to bind to both ARS elements and centromeric DNA sequences. Although its primary function is thought be in chromosome segregation, we show that Abp1 may also have an additional role in DNA replication initiation.

We performed a yeast two-hybrid study to identify proteins that interact with exon11 of BRCA1 and identified Protein Phosphatase 1β (PP1β), an isoform of the serine threonine phosphatase, PP1.

We have used a yeast two-hybrid assay to detect proteins that interact with exon11 of BRCA1. This large exon encodes roughly 60% of the protein, and we wished to identify potentially important interacting proteins outside of the intensely studied RING and C-terminal regions of BRCA1.

We performed a yeast two-hybrid assay to identify proteins that interact with exon11 of BRCA1. In this study, 9 putative positives were identified including PP1β, an isoform of the serine/threonine phosphatase PP1 (not shown). These initial yeast two-hybrid studies led to further examination on the interaction of BRCA1 with PP1β.

We performed a yeast two-hybrid study to identify proteins that interact with exon11 of BRCA1. The region of BRCA1 encoded by exon 11 is known to interact with a number of proteins involved in DNA repair [23], as well as γ-tubulin [3] and several kinases including Aurora-A kinase [24] and ChkII [25]. Identification of additional interacting partners, particularly ones that could modify the activity of a BRCA1 through changes in phosphorylation, may further aid in clarifying its function and regulation. In this yeast two-hybrid study, we identified the serine/threonine phosphatase PP1β as a BRCA1 interacting protein, which could have important consequences on both the activity of BRCA1 and the regulation of PP1β activity.

By a two-hybrid approach, we identify a prefoldin-like protein, ubiquitously expressed transcript (UXT), that is expressed predominantly and interacts specifically with NF-κB inside the nucleus. RNA interference knockdown of UXT leads to impaired NF-κB activity and dramatically attenuates the expression of NF-κB–dependent genes. This interference also sensitizes cells to apoptosis by tumor necrosis factor-α.

To identify new components of the NF-κB enhanceosome, we performed a systematic yeast two-hybrid screening in which the cDNA fragment harboring the RHD of p65 (amino acids 1–312) was used as bait. Several positive clones were identified to encode full-length UXT (Fig. 1 A). In addition, previously confirmed p65-interacting proteins (e.g., IκBα and PIAS3) were screened out.

(A) Interaction between p65 and UXT in a yeast two-hybrid assay.

In the yeast two-hybrid system, we detected no interaction between CO and COP1, although an interaction between COP1 and the CO-related protein CO-LIKE3 (COL3) was previously detected by this method (Datta et al, 2006), and we were able to confirm this interaction.

DLK1 EGF repeats containing the DOS motif bind to specific Notch1 receptor EGF repeats in two-hybrid studies and in tissue culture [50], but the role of DLK1 in Notch signaling remains controversial because DLK1 lacks a DSL domain and does not activate mammalian Notch receptors [42,45–47].

Therefore, we turned to the yeast two-hybrid assay to test whether OSM-11 can interact with LIN-12 extracellular EGF repeats. Conventional wisdom suggests that the yeast two-hybrid system is not suitable for testing extracellular protein–protein interactions, especially for domains rich in disulfide bridges (e.g., EGF repeats). However, two-hybrid interactions have been demonstrated between Notch receptors and ligand pairs in other species for which biochemical interactions have been previously validated [38,39,50], as well as for numerous other extracellular proteins [81–86].

To validate our yeast two-hybrid approach, we first confirmed that the extracellular domain of LAG-2 [23] and the soluble DSL-domain LIN-12 ligand DSL-1 [35] interact with LIN-12 extracellular EGF repeats 1 through 6 in the two-hybrid assay (Figure 8). To the best of our knowledge, this is the first in vitro evidence that C. elegans DSL ligands may bind directly to LIN-12 Notch. As a negative control and to confirm specificity of the two-hybrid assay, we showed that the unrelated C. elegans ligands LIN-3 (an EGF homolog) and egg laying defective-17 (EGL-17) (an FGF homolog) do not interact with LIN-12 extracellular EGF repeats (Figure 8). The LAG-2 and DSL-1 interactions with LIN-12 in the two-hybrid assay are consistent with previous genetic studies in C. elegans and with biochemical analyses of Notch ligand/receptor interactions in other systems. Ligand-receptor interactions were only assayed using LIN-12 fused to the GAL4 activation domain (AD) as LIN-12 EGF fusion to the DNA-binding domain resulted in strong self-activation in the presence of AD empty vector (unpublished data).

DSL-1, OSM-11, LAG-2 extracellular domain (LAG-2Ex), EGL-17, or LIN-3 was fused to the GAL4 DNA binding domain (DB); the first six LIN-12 EGF repeats were fused to the GAL4 activation domain (AD). Pairwise interactions were tested with the yeast two-hybrid assay; positive interactions are indicated by blue staining. Both Notch DSL ligands and OSM-11 interacted with LIN-12 EGF repeats, whereas no interaction of LIN-3 EGF or EGL-17 FGF with LIN-12 Notch receptor EGF repeats was detected. LIN-12::DB fusion proteins exhibited strong self-activation (unpublished data); therefore, reciprocal fusions were not tested. Interaction controls are: (1) empty vectors; (2) DB-pRb and AD-E2F; (3) DB-Fos and AD-Jun; (4) Gal4p and pPC86; and (5) DB-DP1 and AD-E2F1.

We also confirmed previous studies [50] in which murine DLK1 EGF repeats 1 and 2 containing the DOS motif interacted specifically with murine Notch1 EGF repeats 12 and 13 in the same two-hybrid assay format (unpublished data). The two-hybrid interaction does not necessarily demonstrate that OSM-11 and LIN-12 interact in vivo; however, combined with the genetic interactions, the apical expression pattern of OSM-11 in VPCs, and previous studies of DLK1/Notch interactions, we favor a simple model in which OSM-11 binds directly to LIN-12 Notch EGF repeats.

Two-hybrid data and expression on the VPC apical surfaces suggest that OSM-11 may directly bind to the LIN-12 extracellular domain, although additional biochemical studies will be required to further confirm this.

To help identify factors that might be shuttled from the cytosol to the ER by the GET system, we performed a yeast two-hybrid (Y2H) screen for polypeptides that can interact with Get3. Y2H analysis, which reports on weak interactions occurring within the nucleus of assayed strains, is well suited for identifying Get3 binding proteins, as it can detect transient interactions that are independent of the presence of Get1 and Get2. We used yeast expressing Get3 as bait to screen a genomic library encoding prey proteins (James et al., 1996). Physical interactions caused activation of the Gal4-driven HIS3 reporter gene, allowing growth on plates lacking histidine. The strongest hit from the screen was a fragment of Sed5 (amino acid 197 to the C terminus) (Figure 2A), a TA protein that acts as a SNARE in vesicular traffic within the Golgi and between the Golgi and the ER (Hardwick and Pelham, 1992). The Get3-Sed5 interaction was dependent on the presence of the C-terminal TMD (Figure 2A).

Consistent with this idea, by a directed Y2H approach, we detected physical interactions between Get3 and several additional secretory pathway TA proteins, including the SNAREs Tlg2 and Sec22 and the peroxisomal TA protein Pex15. These interactions, as observed for Sed5, were dependent on the presence of the C-terminal TMD (Figure 3A).

First, our Y2H analysis indicates that Get3 can bind multiple secretory pathway TA proteins in a TMD-dependent manner.

(A) Yeast two-hybrid assay with Get3 as bait and Sed5197–340 (the strongest hit from the Y2H screen) as prey (in the presence or absence of its TMD). The growth on medium lacking histidine (−HIS) is indicative of a physical interaction.

(A) Y2H assay showing Get3 as bait and various TA proteins (in the presence or absence of their TMDs) as prey. The growth on medium lacking histidine (−HIS) is indicative of a physical interaction.

To explore the potential targets of p30 during infection, we have used the yeast two-hybrid system to screen a porcine macrophage (the natural viral host cell) cDNA library for cellular proteins that may interact with p30. We have identified heterogeneous nuclear ribonucleoprotein K (hnRNP-K) as the first cellular ligand of p30.

For the yeast two-hybrid assay, plasmids pGBT9 and pACT2 (BD Sciences) were used as sources of the GAL4 DNA-binding domain (BD) and transcriptional activation domain (AD), respectively.

pGBT9-p30 and unrelated control protein pGBT9-p54 were independently used as baits to screen a pACT2 cDNA library from pig macrophages in Saccharomyces cerevisiae reporter strain Y190 as previously published [18,20,21]. Yeast were sequentially transformed with bait plasmid and pACT2 library by the lithium acetate method. After auxotrophic and colony size selection, resulting clones were analyzed for expression of GAL4-dependent β-galactosidase. Plasmid DNA from those clones exhibiting β-galactosidase activity was isolated and retransformed into yeast strain Y190 with pGBT9-p30 to eliminate false positives. The sequence of inserts was determined by sequencing using specific primers and compared with the data base of the NCBI using the BLAST program. pGBT9-p30, pGBT9-p54 and pACT2-K were individually transformed in yeast and tested for β-galactosidase activity to exclude activation of gene reporter by itselves.

To identify cellular proteins interacting with ASFV early protein p30, yeast two-hybrid system was used to screen a porcine macrophage cDNA library. After selection from a total of 5 × 106 transformants screened, two potential positive clones were obtained in the reporter gene assay. DNA sequence analysis showed that cDNA contained in these clones, identical in size and composition, matched the cDNA sequence encoding hnRNP-K. cDNA sequences from positive clones represent nucleotides from 169 to 870 of the hnRNP-K cDNA sequence (GeneBank™ accession number 241477), with 98% nucleotide identity, corresponding to amino acid residues 13–246 of hnRNP-K protein.

In addition, diverse truncations of hnRNP-K were performed attending to previously well characterized functional domains [10] and tested for interaction using the yeast two-hybrid system. The results showed that none of the three different p30 truncations interacted with hnRNP-K (Fig. 2A). On the other hand, we could determine that the hnRNP-K fragment from amino acid residue 35–197 contained the interacting region with p30 (Fig. 2B).

By using the yeast two-hybrid system, we identified cellular hnRNP-K as an interacting protein with ASFV early protein p30.

(A) Schematic representation of the diverse p30 truncations tested for interaction with full length hnRNP-K in the yeast two-hybrid assay. (B) Schematic representation of the diverse hnRNP-K truncations tested for interaction with complete p30 in the yeast two-hybrid assay.

A bait-containing sequence encoding ACBP4 was constructed for yeast two-hybrid screens using a cDNA library derived from A. thaliana to identify proteins that interact directly with ACBP4.

The two-hybrid library screens were performed in the Saccharomyces cerevisiae strain YPB2 [MATa ara3 his3 ade2 lys2 trp1 leu2, 112 canr gal4 gal80 LYS2::GAL1-HIS3, URA3::(GAL1UAS17mers)-lacZ] (Kohalmi et al., 1998). Cotransformants were plated on synthetic dextrose agar plates lacking leucine, tryptophan, and histidine [SD-leu-trp-his] supplemented with 10 mM 3-AT (Kohalmi et al., 1998).

S. cerevisiae strain YPB2 was transformed with bait plasmid pAT188 and transformants were plated on synthetic dextrose agar plates lacking leucine [SD-leu]. An aliquot of transformants was also tested on [SD-leu-his] medium supplemented with 10 mM 3-amino-1, 2, 4-triazole (3-AT) because an absence of growth on this medium would confirm that the DB-‘bait’ fusion protein is unable to initiate transcription of HIS3. Subsequently, the bait-carrying strain was tested negative for β-galactosidase activity using the X-Gal (5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside) colony filter assay. This further showed that the bait was not able to activate transcription of the lacZ reporter gene. The prey vector pBI-771, a variant of pPC86 (Chevray and Nathans, 1992; Kohalmi et al., 1998), was introduced into this strain and its inability to grow on [SD-leu-trp-his] medium supplemented with 10 mM 3-AT and its lack of β-galactosidase activity were confirmed before the bait was further used in cDNA library screening.

To ensure sufficient coverage in the identification of potential proteins interacting with ACBP4, yeast two-hybrid screenings were also performed at the Molecular Interaction Facility, University of Wisconsin–Madison using yeast strains and vectors as previously described by James et al. (1996). For bait preparation, ACBP4 (amino acids 1–669) was cloned in-frame with the GAL4 DNA-binding domain of bait vector pBUTE (a kanamycin-resistant version of GAL4 bait vector pGBDUC1). The resulting vector was subject to DNA sequence analysis to confirm the presence of an in-frame fusion, before use in transformation of S. cerevisiae mating type strain PJ69-4A, followed by testing for autoactivation of the β-galactosidase reporter gene.

The yeast YPB2 transformed with the bait GAL4(DB)-ACBP4 could not grow on [SD-leu-his] and was tested negative on X-Gal colony filter assays (data not shown), suggesting that the pAT188 bait alone could not activate the transcription of reporter genes HIS3 and lacZ and was deemed appropriate for two-hybrid screens. A GAL4(TA) tagged A. thaliana cDNA library was introduced into the yeast YPB2 harbouring plasmid pAT188. The number of independent transformants was determined to be 2×106 following transformation and plating of an aliquot of the yeast transformation mixture on [SD-leu-trp]. A total of 100 putative positives were selected on [SD-leu-trp-his] supplemented with 10 mM 3-AT medium. When these putative positives were further screened for β-galactosidase activity using the X-Gal colony filter assay, nine yeast clones that appeared blue, at varying intensities, were identified as putative clones encoding interactors. Putative library plasmids were retrieved and their nucleotide sequences were searched against the BLAST server http://www.ncbi.nlm.nih.gov/cgi-bin/BLAST. Only one clone was in-frame to GAL4(TA), encoding a full-length ethylene-responsive element binding factor (ERF) protein AtEBP (Arabidopsis genome locus: AT3G16770). An AP2/EREBP (ethylene-responsive element binding protein) domain is present in AtEBP at amino acids 76–143 (Okamuro et al., 1997).

In another independent yeast two-hybrid screen using the Molecular Interaction Facility (University of Wisconsin–Madison), six putative positives were identified following selection on histidine drop-out and β-galactosidase assays. Subsequently, they were used to retransform yeast mating type strain PJ69-4A, and were validated in mating and selection assays using the ACBP4 bait, the empty bait vector, and unrelated baits. Five clones were tested positive and further identified by nucleotide sequence analysis. Results from analysis using the BLAST revealed that only one clone was in-frame and it encoded a full-length actin-depolymerizing factor 3 (ADF3, At5g59880) protein.

Results of X-Gal filter assays are shown in Fig. 1A. Positive protein–protein interaction results in activation of the reporter gene β-galactosidase in yeast cells, which turns yeast colonies blue in filter assays using X-Gal. Without interaction, the yeast colonies remain ‘colourless’. As shown in Fig. 1Aa, the GAL4(DB)-ACBP4 fusion interacted with GAL4(TA)-AtEBP, as indicated by the blue colour arising from the production of significant levels of β-galactosidase. No interactions were observed in control yeast cells harbouring GAL4(DB)-ACBP4+GAL4(TA) (Fig. 1Ab) and GAL4(DB)+GAL(TA)-AtEBP (Fig. 1Ac). Therefore, from yeast two-hybrid analysis, AtEBP was identified as a putative protein that interacts with ACBP4.

(A) Colony filter β-galactosidase assays of candidate proteins AtEBP from yeast two-hybrid screens. (a) YPB2/GAL4(DB)-ACBP4+GAL4(TA)-AtEBP; (b) YPB2/GAL4(DB)-ACBP4+GAL4(TA); (c) YPB2/GAL4(DB)+GAL(TA)-AtEBP.

Kelch-motif containing ACBP4 was used as bait in yeast two-hybrid screens from which an interactor (AtEBP) was retrieved.


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This assumption is supported by the fact that the MEINOXSTM domain is precipitated via the BELLBLH3 domain in co-immunoprecipitation experiments (data not shown).

Co-immunoprecipitation and BiFC experiments performed in the course of this study show that the MEINOXSTM and the BELLBLH3 domains are sufficient for interaction in vitro and in planta.

Mutagenesis of residue R1161 (in α2AA and α2AAKYA, removing another conserved charged residue) to alanine significantly reduced α2 tail association with Rab21 judged by yeast mating tests and immunoprecipitations (Fig. 1, D and E). In addition, F1159A mutation (α2AARA, possibly creating a conformational change) showed reduced association in the yeast assays, whereas in immunoprecipitations the reduction was not significant. These data on the mutant integrins suggest that the conformation of the cytoplasmic domain and residue R1161 of α2-integrin are important for the Rab21 association.

(A) GFP-Rab21–transfected HeLa cells on plastic (P) or on collagen (CI; 1 h) were subjected to immunoprecipitations (IPs) with the indicated antibodies followed by blotting with anti-α2 or anti-GFP. (B and C) Full-length Rab21WT tagged with Rluc (Rluc-Rab21), mutants (Rluc-Rab21GTP, -Rab21GDP, or -Rab21C-del), or Rluc alone were expressed in Saos-2 cells stably expressing chimeric α-integrin subunits (α2/α1) or (α2/α5) (B) or HT1080 cells (C), and immunoprecipitation was performed with the indicated antibodies. The coprecipitated luciferase activity is presented relative to the basal nonspecific activity detected in the relevant control immunoprecipitations (anti-EGFR). Efficiency of the integrin immunoprecipitation was analyzed by Western blotting with anti-β1 from the same beads (means ± SD; n = 5; ***, P < 7 × 10−5; **, P < 0.001).

(E) CHO cells were cotransfected with Rluc-Rab21 and GFP-α2 variants. Immunoprecipitation was performed with anti-GFP antibody, and the coprecipitated luciferase activity is presented relative to the activity detected in immunoprecipitations from α2WT-expressing cells. Efficiency of the integrin immunoprecipitation was analyzed by Western blotting with anti-α2 from the same beads (means ± SD; n = 5; *, P < 0.05; †, not significant). (F) Endogenous proteins were immunoprecipitated from MDA-MB-231 cells using the indicated antibodies and blotted with anti-β1 or anti-Rab antibodies.

(M) Immunoprecipitation (IP) was performed with anti–β1-integrin mAb or control mouse IgG from GFP-Rab21WT– or -Rab21CCSS–transfected MDA-MB-231 cells followed by Western blotting with anti-β1 or anti-GFP. Equal amounts of lysates were loaded on the gel to control for transfection efficiency.

(A) Cells transiently cotransfected with GFP-α2 and Rluc-Rab21 were subjected to the following immunoprecipitations (IPs): anti-GFP, anti-β1, and mouse IgG for control. The coprecipitated luciferase activity (cps) is shown, and the values above the bars indicate luminescence detected relative to the control immunoprecipitation.

To determine the effect of P-body disruption on Ago2 and RCK/p54 interactions, we immunopurified Myc-Ago2 and RCK/p54 after Lsm1 knockdown. HeLa cells were transfected for 48 h with Myc-Ago2 and control siRNA or siRNA against Lsm1, TCEs were prepared, and Myc-Ago2 was immunoprecipitated from an aliquot of TCE. TCEs and anti-Myc immunoprecipitation products were analyzed by immunoblot using anti-Myc, anti-RCK/p54, and anti-Lsm1 antibodies. Lsm1 siRNA treatment efficiently depleted Lsm1 protein levels without affecting Myc-Ago2 and RCK/p54 levels (Figure 5C). Immunoprecipitation using Myc antibodies showed that RCK/p54 interacted with Ago2 and this interaction was not significantly changed by depleting Lsm1 (Figure 5C). Since P-bodies were disrupted by Lsm1 depletion, these results demonstrate that Ago2 and RCK/p54 interaction does not require P-bodies.

Ago1 and RCK/p54 immunoprecipitated with Ago2 after RNase A treatment of HeLa cell extracts, suggesting that these proteins directly interact with Ago2. Interestingly, RCK/p54, Ago1, and Ago2 were also identified as a component of active RISC programmed with siRNA or miRNA and purified by biotin affinity to streptavidin-conjugated magnetic beads (Figures 2 and 3). We examined the P-body localization of Ago2 with Lsm1 and RCK/p54 by co-expressing YFP-tagged Lsm1 and RCK/p54 with CFP-Ago2. Interestingly, overexpressing YFP-RCK/p54 in HeLa cells increased the number of P-bodies (from ∼ 8 to ∼ 20 foci/cell). The number of P-bodies containing CFP-Ago2 also increased (Figure 1B). These results suggested a functional relationship between RCK/p54-Ago interactions and their localization to P-bodies.

TCEs from HeLa cells co-expressing Myc-Ago2 and YFP-Ago1, YFP-Dcp2, YFP-RCK/p54, YFP-eIF4E, YFP-Lsm1, or YFP were treated with +/− RNase A followed by Myc-Ago2 immunoprecipitation.

(C) RCK/p54 interacts with Myc-Ago2 in Lsm1-depleted cells. HeLa cells were transfected for 48 h with Myc-Ago2 and control siRNA or siRNA against Lsm1, TCEs were prepared, and Myc-Ago2 was immunoprecipitated from an aliquot of TCE.

Figure 2A shows that Bcl-xL measured in whole cell lysates from pretumourigenic CD45−/−lckF505 murine thymocytes is resistant to deamidation following γ irradiation, consistent with our previous findings [15]. Immunoprecipitation of the pro-apoptotic protein Bim, followed by immunoblotting for Bcl-xL, revealed that Bim sequestered only the N52/N66 Bcl-xL and failed to bind the slower migrating deamidated protein (Figure 2A, upper panel), although the amount of Bim in each immunoprecipitate was comparable (Figure 2A, lower panel). Because the BH3-only protein Puma, not Bim, plays a major role in DNA-damage triggered apoptosis [19,20], we also showed that both Puma and Bim are found in Bcl-xL immunoprecipitates from etoposide treated CD45−/−LckF505 thymocytes, whereas sequestration is ablated in wild-type cells, correlating with Bcl-xL deamidation (Figure 2B). A comparable result was obtained when Puma immunoprecipitates were blotted for Bcl-xL (Figure S2A).

(A) Bim binds to the native (Asn-Asn) but not deamidated forms of Bcl-xL. Wild-type (C57BL/6) thymocytes (1.5 × 107) were exposed to 5 Gy irradiation (IR) and then maintained in culture for the times shown, after which cells were lysed and either separated as whole cell lysates (WCL) or as Bim immunoprecipitates, followed by immunoblotting for either Bcl-xL or for Bim. Bim migrates as “extra-long” (EL) or “long” (L) forms.

(B) Bcl-xL was immunoprecipitated from lysates derived from purified DN thymocytes treated with/without etoposide (ut/E), followed by immunoblotting for Bim or Puma. The asterisk indicates the light chain of the Bcl-xL antibody used for immunoprecipitation.

(E) Primary thymocytes were retrovirally transduced with empty vector or Bcl-xL constructs (wild-type, N52A-N66A, or N52D-N66D). Bcl-xL was immunoprecipitated from lysates derived from 1.5 × 106 sorted GFP-positive cells per lane, followed by immunoblotting for Bim or Puma. Note that in the vector lane, at this exposure endogenous Bcl-xL is not visible because of the small number of cells used. The asterisk indicates the light chain of the Bcl-xL antibody used for immunoprecipitation.

Protein was immunoprecipitated using antibodies against BRCA1, or an antibody against the Flag epitope to immunoprecipitate Flag epitope tagged PP1α, β, or γ. BRCA1 coimmunoprecipitated all three PP1 isoforms, and conversely, PP1 α, β and γ coimmunoprecipitated BRCA1 (Figure 2), indicating that the interaction between BRCA1 and PP1 is specific.

(D) 293T cells were transfected with HA-UXT. 24 h after transfection, cells were treated with 10 ng/ml TNF-α for the indicated times and fractionated to cytoplasmic and nuclear fractions, which were immunoprecipitated and immunoblotted with the indicated antibodies, respectively. (E) 293T cells were treated with 10 ng/ml TNF-α for the indicated times. Whole cell lysates were immunoprecipitated and immunoblotted with the indicated antibodies. Bar, 10 μm.

To explore the UXT-binding region within p65, we constructed a series of p65 deletion mutants (Fig. 2 A). It was found that the loss of amino acids 1–285 at the N terminus of p65 resulted in its complete inability to interact with UXT (Fig. 2 B, top). In contrast, p65 fragments spanning amino acids 1–286, 1–312, or 1–372 fully retained their binding capability and interacted with UXT as well as the wild type (Fig. 2 B, middle). Because the RHD of p65 consisted of two Ig-like domains (Chen et al., 1998), we made two additional deletion mutants of p65 (amino acids 1–190 and 191–551), each of which contained only one Ig-like domain. Immunoprecipitation assays revealed that neither of them was able to interact with UXT (Fig. 2 B, bottom).

(B) Tagged full-length UXT was transfected into 293T cells along with p65 and its deletion mutants as indicated. Whole cell lysates were immunoprecipitated and immunoblotted with the indicated antibodies. (C) 293T cells were transfected with FLAG-UXT together with HA-p50, myc-cRel, and HA–lymphoid enhancer binding factor 1. Cell lysates were immunoprecipitated and immunoblotted with the indicated antibodies.

The co-IP of these PDE4D splice variants expressed exogenously in HEK293 cells identified PDE4D8 as the variant that most efficiently interacts with β1AR. Other long PDE4D splice variants were also recovered in β1AR IP pellets with the following rank order: PDE4D8>PDE4D9>PDE4D3>PDE4D5 (Figure 3A and B). Conversely, the short PDE4D form, PDE4D2, did not co-IP with the β1AR, indicating that the UCR domains unique to long PDE4 splice variants may contribute to the formation of the β1AR/PDE4D complex.

However, the co-IP of exogenous PDE4D and β1AR from extracts of mouse embryonic fibroblasts (MEFs) deficient in β-arrestin 1 and 2 (Kohout et al, 2001) was not decreased compared with wild-type controls, suggesting that formation of the β1AR/PDE4D complex is independent of β-arrestins (Figure 3C). To further characterize the interaction, we performed IPs using PDE4D and βARs that were purified from a baculovirus expression system to >90% purity (see Supplementary Figure 1 for the characterization of the purified proteins).

β1AR preferentially associates with PDE4D8 in cardiomyocytes as shown by co-IP of endogenous PDE with the β1AR (Figure 1C), as well as the selective activation of PDE4D8 in intact cells (Figure 5A). This preference of β1AR for PDE4D8 was confirmed by co-IP experiments with exogenous proteins (Figure 3A and B).

(B, C) Detergent extracts from neonatal cardiac myocytes were immunoprecipitated with PAN-selective antibodies for the PDE4 subtypes, PDE4A, PDE4B, and PDE4D (B), or with splice variant-selective anti-PDE4D antibodies (C).

It remains to be determined to what extent PDE4D9, which is activated after both β1AR and β2AR stimulation (Figure 5) and which also showed interaction with β1AR in co-IPs of exogenous proteins (Figure 3A and B), can substitute for interaction with the βARs in vivo.

To identify the mechanism of Plx1 recruitment to chromatin, we tested the interaction of Plx1 with the Mcm complex. We found that Plx1 co-precipitated with Mcm7 and that its binding was enhanced by activation of the checkpoint induced by pApT (Figure 5C).

(C) Co-immunoprecipitation of Plx1 and Mcm7. Extracts that were untreated (−) or treated with 50 ng/μl pA/pT (+) were immunoprecipitated with pre-immune (Pre-Imm) or anti-Plx1 (Anti-Plx1) antibodies.

Therefore, whether COP1 and CO interact in vitro was tested using a co-immunoprecipitation assay (Figure 3). COP1 attached to the GAL4 activation domain (GAD:COP1) and CO were made in an in vitro transcription/translation system and combined. GAD:COP1 was precipitated with anti-GAD antibody and CO was co-precipitated with GAD:COP1 (Figure 3).

In vitro precipitation experiments demonstrated that COΔB-box was co-immunoprecipitated with GAD:COP1, whereas COΔCCT was not. Therefore, the N-terminal region containing the B-boxes is not required for interaction with COP1, suggesting that the interaction with COP1 is mediated by the C-terminal region of CO that contains the CCT domain.

COP1 directly interacts with target proteins and directs them for degradation (Hoecker, 2005; Jiao et al, 2007). CO is composed of three domains, zinc-finger B-boxes, a central domain and the C-terminal CCT domain (Wenkel et al, 2006). CO and COP1 interact directly in vitro as demonstrated by immunoprecipitation experiments. This interaction was almost abolished when the C-terminal part of CO was removed, suggesting that COP1 interacts with the C-terminal region of CO, as was previously observed for interactions between COP1 and COL3 or between CO and SPA1 (Datta et al, 2006; Laubinger et al, 2006).

In vitro interaction between CO and COP1 detected by co-immunoprecipitation. 35S-methionine-labeled CO, COΔB-box or COΔCCT was incubated with 35S-methionine-labeled GAD:COP1 or GAD and co-immunoprecipitated with anti-GAD antibodies. Supernatant fractions and pellet fractions were resolved by SDS–PAGE and visualized by autoradiography using a phosphorimager. Quantification of the fractions of prey proteins that were co-immunoprecipitated by the indicated bait proteins GAD:COP1 or GAD. Error bars denote the standard error of the mean of two replicate experiments.

When both proteins were in the membranes, Bcl-XL bound to tBid, as assessed by co-immunoprecipitation. This interaction was not dependent on the detergent used to solubilize the liposomes (unpublished data), and at concentrations of 100 nM Bcl-XL and 20 nM tBid the interaction was not affected by the addition of Bax (Figure 2A, left panel, lanes 1 and 2). In the absence of membranes, interaction between Bcl-XL and tBid could not be detected by co-immunoprecipitation (unpublished data).

(A) Bax (100 nM) and/or tBid (20 nm) were incubated with 100 nM Bcl-XL (left panel) or 20 nM Bcl-XL (right panels) and liposomes. Samples were immunoprecipitated (IP) in either 2% CHAPS or 0.2% NP-40, as indicated, using an antibody with the indicated specificity and immunoblotted (IB) for the indicated protein.

(B–D) Mutations prevent the binding of Bcl-XL to tBid, Bax, or both. (B and D) Bcl-XL (20 nM), or the indicated Bcl-XL mutants, (C) Bcl-XL Y101K or (D) Bcl-XL ΔBH4, were incubated with (C and D) 20 nM tBid or (B) tBid-mt1 with or without Bax (100 nM) and with liposomes. Immunoprecipitations and immunoblotting were performed as in (A).

Although sequestration of membrane-bound tBid by Bcl-XL appears to account for its antiapoptotic function in both liposomes and MLM, we sought to determine whether Bcl-XL also could interact stably with Bax. When tested in the absence of tBid, a stable interaction between Bax and Bcl-XL was not detected (Figure 2A, left panel, lane 3), suggesting that Bcl-XL does not sequester Bax in solution. However as expected, co-immunoprecipitation of Bcl-XL and Bax was observed in control experiments where membranes were solubilized with the nonionic detergent NP-40, known to induce a conformational change in Bax required for heterodimerization with Bcl-XL that is also seen in cells when apoptosis is induced [20,26].

(A) Bax (100 nM) was incubated in the presence of liposomes and in the absence or presence of tBid (20 nM). Conformation-altered Bax was immunoprecipitated using the 6A7 antibody with or without the addition of 2% CHAPS to solubilize the liposomes prior to immunoprecipitation and analyzed by immunoblotting using an α-Bax antibody. The asterisk denotes the light chain of the 6A7 antibody.

(B) Bax was incubated in the presence of liposomes, tBid, and increasing concentrations of Bcl-XL. Immunoprecipitations and immunoblotting were performed as in (A) without the addition of 2% CHAPS.

(C) Bax was incubated for 2 h with liposomes (without tBid) at increasing concentrations of Bcl-XL or Bcl-XL Y101K. Immunoprecipitations and immunoblotting were performed as in (B).

(F) Membrane-bound Bcl-XL inhibits the liposome-induced Bax conformational change with 50 μM m1Bid but not with 50 μM Bak BH3 peptide. Immunoprecipitations and immunoblotting were performed as in (B).

Pioneering experiments to identify relevant binding partners for Bcl-2 and Bcl-XL using immunoprecipitation in transfected cells suggested a lack of correlation between Bax binding and inhibition of apoptosis, as only certain Bcl-XL point mutants that could no longer bind to Bax lost function [40].

Immunoprecipitation of Bcl-XL was performed using the polyclonal Bcl-XL antibody in assay buffer containing either 2% CHAPS or 0.2% NP-40. Immunoprecipitates were collected as previously described [7] and washed three times in assay buffer containing the appropriate detergent. Immunoprecipitation using the conformation-specific 6A7 Bax antibody was performed on whole membranes and washed three times with assay buffer containing 2% CHAPS.

Immunoprecipitation experiments using tagged TBK1 suggested that its interaction with DDX3X and the transcription factor IRF3 are significantly weaker than the interaction between TBK1 and TANK and therefore not detected by coimmunoprecipitation under stringent conditions (Supplementary Figure 1A).

The wild-type MDC1 derivative efficiently co-immunoprecipitated MRN at physiological salt concentrations, whereas only low levels of MRN were recovered in immunoprecipitates of the MDC1SDTDΔ mutant (Fig 4B), consistent with the SDTD region being the principal MRN interaction interface.

(B) Indicated expression constructs were transfected into human embryonic kidney 293 cells. After 48 h, extracts were prepared, immunoprecipitated (IP) with GFP antibodies and immunoblotted. Immunoprecipitations and washes were performed at 150 mM salt; INP, input (5%). (C,D) Indicated osteosarcoma (U2OS) cell lines were treated with two rounds of control (CNTL) or MDC1-targeting siRNA for 72 h. Cells were then treated with 5 Gy of X-rays and processed for immunofluorescence 4 h later with MDC1, (C) NBS1 or (D) 53BP1 antibodies (non-irradiated cells are shown in supplementary Fig S4B,C online).

Co-IP experiments were repeated using the HCT116 SIRT3 expressing stable cell lines. FOXO3a was found to interact with SIRT3 in both whole cell (Fig. 1B) and mitochondrial extracts (Fig. 1C). All fractions and samples in the Co-IP experiments were checked for the presence of SIRT3, using a SIRT3 specific antibody (Biomol, Plymouth Meeting , PA), as well as tubulin (Santa Cruz Biotechnology, Santa Cruz, CA), and Cytochrome C (Mitosciences, Eugene, OR) to ensure fraction purity (data not shown). These experiments demonstrate that both wild type and mutant SIRT3 form a physical interaction with FOXO3a in mitochondrial extracts.

To corroborate the interaction from yeast two-hybrid analysis, co-immunoprecipitation studies were performed according to Mongiat et al. (2003). All constructs used in these interaction assays were derivatives of vector pBluescriptII KS(–) (pKS). The HindIII-SacI fragment from pBI-771 carrying GAL4(TA) (amino acids 768–881) was cloned into corresponding restriction sites on pKS. The GAL4(TA)-ACBP4 fusion construct was prepared by inserting ACBP4 cDNA from pAT181, on a 2 kb EcoRI-BamHI fragment, into the EcoRI-BglII sites of pKS-TA with the 5′ of TA-ACBP4 adjacent to the T3 promoter.

Co-immunoprecipitation with monoclonal anti-GAL4(TA) antibody (Clontech, USA) was performed following Mongiat et al. (2003).

(B) Co-immunoprecipitation of ACBP4 and AtEBP using the anti-GAL4(TA) monoclonal antibody. Autoradiograph of a 12% SDS-PAGE (left panel) showing the in vitro transcribed and translated ADF3, AtEBP, and GAL4(TA)-ACBP4, respectively, as indicated. The right panel shows the co-immunoprecipitation of equimolar amounts of GAL4(TA)-ACBP4 and ADF3 or AtEBP using the anti-GAL4(TA) antibody. Arrows indicate the positions of these proteins.

Co-immunoprecipitation of in vitro transcription/translation products to the GAL4(TA)-ACBP4 fusion protein, immobilized to protein A/agarose beads, using monoclonal antibody against GAL4(TA), showed that the GAL4(TA)-ACBP4 fusion protein significantly binds AtEBP (Fig. 1B). However, no binding of GAL4(TA)-ACBP4 to ADF3 was observed (Fig. 1B), perhaps due to the lack of cofactors which must be present for their in vitro interaction.

The interaction of AtEBP and ACBP4 was further substantiated by co-immunoprecipitation and by using autofluorescent protein fusions in the transient expression of tobacco leaf epidermal cells. ACBP4 and AtEBP showed overlapping expression patterns in leaves and stems and both were inducible by ACC, MeJA treatment, and infection with the fungal pathogen, Botrytis cinerea.


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In density gradient fractionations, expression of GFP-Rab21 shifted the integrins toward the denser Rab-positive fractions (Hughes et al., 2002), and GFP-Rab21 cofractionated with α2-integrin in fractions 3–9 (Fig. 3 D). A further shift in the endogenous integrin pool (to fractions 5–11) was observed upon expression of GFP-Rab21GTP, and GFP-Rab21GDP was also observed in the denser fractions (Fig. 3 D). In the lighter fractions (3–5), GFP-Rab21 and integrin were found to cosediment with the Golgi-marker GM130, whereas in the denser fractions, cosedimentation was observed with the ER marker P115 (fractions 6–8) and EEA1 (fractions 7–9; Fig. S3 A, available at http://www.jcb.org/cgi/content/full/jcb.200509019/DC1). This data, together with the abundance of integrin vesicles observed in Rab21-expressing cells (Fig. 2), suggests that Rab21 targets integrins to the endocytic fraction in human cells.


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To determine whether the Bcl-XL/Bax heterodimer also prevented the subsequent oligomerization of Bax, we examined oligomerization by cross-linking. In these experiments, the cross-linker was added to reactions containing an equal amount of membrane-bound Bax in the absence or presence of Bcl-XL (Figure S3). In these reactions, membrane-bound Bcl-XL inhibited Bax oligomerization, as detected by cross-linking concomitant with inhibition of dye release from liposomes. Taken together, these results suggest that, when bound to Bcl-XL, Bax function is neutralized, both in recruitment of other Bax molecules through autoactivation and in oligomerization to permeabilize membranes (Figure 6C, step 6).

(D and E) Bcl-XL inhibits liposome-induced cross-linking of Bax. (D) Bax (100 nM) was incubated with liposomes for 2 h either alone (left panel), with 20 nM tBid (middle panel), or with 20 nM tBid and 100 nM Bcl-XL (right panel). Cross-linking with DSS was performed for 30 min at room temperature with or without 2% CHAPS to solubilize the liposomes prior to cross-linking, as indicated. Results were analyzed by immunoblotting. (E) Bax (100 nM) was incubated with or without liposomes for 2 h. Cross-linking and immunoblotting were performed as in (D).

To investigate further the effects of the membrane surface on Bax and the inhibition of these effects by Bcl-XL, cross-linking experiments using disuccinimidyl suberate (DSS) were performed. Cross-linking of Bax into higher-order structures after Bax binds to membranes has been observed previously [18].

Nevertheless, incubation with liposomes did result in the cross-linking of Bax into higher-order complexes (Figure 5D, left panel). As expected from previous results [7,9], the interactions between Bax monomers induced by incubation with membranes were not resistant to detergent solubilization prior to cross-linking. The Bax–Bax cross-links were reduced in the absence of liposomes (Figure 5E), suggesting that, similar to binding by the 6A7 antibody, they result from a liposome-induced conformational change in Bax. Addition of tBid to Bax and liposomes resulted in a similar cross-linking pattern, but these Bax oligomers were resistant to solubilization of the membrane with detergent (Figure 5D, middle panel). Membrane-bound Bcl-XL not only prevented the formation of detergent-resistant Bax cross-links but also prevented the cross-linking of Bax that resulted when Bax contacted the membrane surface (Figure 5D, right panel).

Kelch motifs, structural repeats first observed in the Drosophila actin cross-linking protein kelch, allow protein folding into a cylindrical ‘β-propeller structure’ (Adams et al., 2000) forming a potential protein–protein interaction domain (Andrade et al., 2001).


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Arabidopsis leaves were fixed in a solution of 4% (v/v) paraformaldehyde and 0.5% (v/v) glutaradehyde in 0.1 M phosphate buffer (pH 7.2) for 20 min under vacuum and then a further 3 h at room temperature. The specimens were then dehydrated in a graded ethanol series, infiltrated in stepwise increments of LR white resin (London Resin, Theale, Berkshire, UK) and polymerized at 45 °C for 24 h. Materials for immuno-gold labelling were prepared according to the procedure of Varagona and Raikhel (1994) with the modification as described. Specimens (90 nm) were sectioned using a Leica Reichert Ultracut S microtome and mounted on formvar-coated slotted grids. Grids were incubated in a blocking solution of TTBS containing 1% (w/v) fish skin gelatin and 1% (w/v) BSA for 30 min. Anti-ACBP4 antibodies diluted 1:50 in blocking solution were added and incubated at room temperature for 2 h. The grids were then rinsed three times, each for 5 min, in TTBS and then incubated with 10 nm gold-conjugated goat anti-rabbit IgG secondary antibody (Sigma), diluted 1:20 with blocking solution. Grids were rinsed three times, each for 5 min in TTBS, following by three 5-min rinses in distilled water. After being stained in 2% (w/v) uranyl acetate for 6 min followed by 2% (w/v) lead citrate for 6 min, the sections were visualized and photographed using Philips EM208s electron microscope operating at 80 kV. Controls were performed excluding the primary antibody.

(B, C, D) Immuno-gold labelling of ACBP4 in an Arabidopsis leaf cell using transmission electron microscopy. Transverse sections were stained with affinity-purified ACBP4-specific antibodies. (B) Transverse sections of leaves stained with ACBP4-specific antibodies. (C) Magnification of the boxed area in (B). (D) Control labelling of a leaf cell using secondary antibodies alone. Arrowheads, gold particles. V, vacuole; C, cytosol; Ch, chloroplast; N, nucleus; Cw, cell wall; Bars in (B) represent 2 μm, and in (C, D), 0.2 μm.

Immuno-electron microscopy was carried out using transverse sections of leaves of 2-week-old Arabidopsis germinated and grown in MS medium under a 16/8 h light/dark regime. Although immuno-gold labelling with the anti-ACBP4 antibodies was mostly evident in the cytosol, some signals were detected at the periphery of the nucleus, (Fig. 3B, C). In the control, when the primary antibody was replaced by blocking solution, no significant immuno-gold labelling was observed (Fig. 3D). The immunolocalization of signals at the periphery of the nucleus may have culminated from the interaction of ACBP4 with AtEBP.

In this study, GFP:AtEBP was not confined to the nucleus but was also detected in the cytosol where it could interact with ACBP4. ACBP4:DsRed, transiently-expressed in tobacco leaves, was predominantly targeted to the cytosol but immuno-electron microscopy indicated localization of ACBP4 in the cytosol with signals detected at the periphery of the nucleus, perhaps as a consequence of its interaction with AtEBP.


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To establish whether the N-terminal region of α−parvin, which is excluded from our structural analysis, contributes to LD recognition, we compared binding of a fluorophore-labeled LD1 peptide to either α-parvin-CHC or full-length α-parvin by fluorescence anisotropy.


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To investigate this hypothesis, we transiently co-transfected two plasmid clones designed to express HA-tagged MELK (WT or D150A) and Flag-tagged Bcl-GL into COS7 cells, and then performed a TUNEL assay and FACS analysis to measure the proportions of apoptotic cells (see Material and methods).

As reported previously [14], we demonstrated by TUNEL assay and FACS analysis that introduction of full-length Bcl-GL into COS7 cells induced apoptosis. However, under the same conditions, addition of exogenous WT-MELK suppressed induction of apoptosis by Bcl-GL, but addition of D150A-MELK did not (Figure 5b–d).

Notably, we did not detect any cell-cycle-dependent alteration of MDC1 S329/T331 phosphorylation relative to total MDC1 protein content by using flow cytometry, which suggests that SDTD phosphorylation occurs throughout interphase (supplementary Fig S1B online).

(A) Altered steady-state levels of superoxide as shown by increased oxidation of DHE in HCT116 SIRT3 overexpressing cells. Control (pcDNA vector), wild-type SIRT3 (wt-SIRT3), or a deacetylation SIRT3 mutant (mut-SIRT3) cell lines were analyzed by flow cytometry for the amount of hydrolyzed DHE per 10,000 cells represented as Mean Florescent Intensity (MFI).


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To visualize protein–protein interactions in vivo, we used fluorescence resonance energy transfer (FRET) as a probe. In FRET, a fluorescent donor molecule transfers energy via a nonradiative dipole–dipole interaction to an acceptor molecule [46]. We used a well-known donor: acceptor fluorescent-protein pair, CFP:YFP, with a Förster distance (R 0) of 4.9 nm [47]. To determine whether Ago1 and Ago2 interacted in vivo with each other and with RCK/p54, we measured the FRET efficiency between the donor, CFP-Ago2, and acceptor, YFP-RCK/p54. To do so, we used a method in which the donor signal lost during FRET is restored by deliberately photobleaching the acceptor fluorophore to abolish its capacity as an energy acceptor [48– 50]. In cells expressing YFP-RCK/p54 and CFP-Ago2, the FRET efficiency was 21.07% ± 2.52% (Figure 1C and (Figure1D). In cells expressing CFP-Ago2 and YFP-Lsm1, FRET efficiency was not significant (1.62% ± 1.11%), corroborating our immunoprecipitation results (Figure 1A). Furthermore, cells co-expressing YFP-RCK/p54 and CFP showed no significant FRET efficiency (1.64% ± 1.28%). Similar to the YFP-RCK/p54 and CFP-Ago2 pair, YFP-Ago1 and CFP-Ago2 showed an efficient FRET (19.61% ± 4.51%), indicating a direct interaction between Ago1 and Ago2 in vivo [28].

Interestingly, the FRET efficiency between Ago1 and Ago2 decreased to 12.13% ± 1.6% when we used CFP-Ago1 and YFP-Ago2, indicating that the energy transfer efficiencies were sensitive to the orientation of donor: acceptor pair in the ribonucleoprotein (RNP) complex. Moreover, only moderate energy transfer efficiency (6.41% ± 1.96%) was seen when YFP-RCK/p54 and CFP-Ago1 were used in FRET experiments, suggesting that this donor: acceptor pair was not as ideally oriented for an efficient energy transfer as the pair CFP-Ago1 and YFP-Ago2. Alternatively, RCK/p54-Ago1-Ago2 is assembled in an RNP complex where the donor: acceptor pair is affected by the location of the probe. Nonetheless, the efficiency of energy transfer was well above the background control (0.99%). As a control experiment, cells co-expressing YFP-Ago1 and CFP showed no significant FRET efficiency (0.99% ± 0.67%). Taken together, these results indicate that Ago1 and Ago2 directly interact in vivo with each other and with RCK/p54.

To visualize protein–protein interactions in vivo, we used FRET as a probe. In cells expressing YFP-RCK/p54 and CFP-Ago2, the FRET efficiency was 21.07% ± 2.52% (Figure 1C and 1D). We also observed an efficient FRET between YFP-Ago1 and CFP-Ago2; however, FRET between RCK/p54 and Ago1 was moderate (6.41% ± 1.96%). Since FRET is quite sensitive to the orientation of the donor: acceptor pair, it is possible that CFP and YFP in Ago1 and RCK/p54 are not suitably positioned for efficient energy transfer. Nonetheless, the FRET efficiency between Ago1 and RCK/p54 was significantly above the 0.9% background. Taken together, these results demonstrate that Ago1, Ago2, and RCK/p54 directly interact in vivo.

(C) Visualization of interactions between RCK/p54 and Ago2 in P-bodies by FRET. HeLa cells expressing YFP-RCK/p54 and CFP-Ago2 were fixed at 24 h post-transfection. FRET was measured by an acceptor photobleaching method. Fluorescence images of donor (CFP-Ago2) and acceptor (YFP-RCK/p54) molecules were taken before and after photobleaching YFP. FRET efficiencies were calculated as described [48,49,68], and data were analyzed by Leica confocal software. Arrows point to P-bodies, which are enlarged in insets.

Whether the interaction between CO and COP1 also occurred in vivo in plant cells was tested using fluorescent resonance energy transfer (FRET). Microprojectile bombardment was used to co-express cyan fluorescent protein (CFP):COP1 and yellow fluorescent protein (YFP):CO in leaf epidermal cells of Arabidopsis. CFP:COP1 and YFP:CO colocalized to the nucleus and also colocalized in speckles within the nucleus (Figure 4A and B). Physical interaction of CFP:COP1 and YFP:CO was tested by measuring FRET using photoacceptor bleaching, as previously described (Wenkel et al, 2006) (Figure 4C and D). Quantification of FRET signals demonstrated that FRET occurred between YFP:CO and CFP:COP1 both in the nucleus and specifically in nuclear speckles (Figure 4C and D). In control experiments using YFP and CFP, YFP:CO and CFP or YFP and CFP:COP1 FRET was detected at significantly lower levels (Figure 4C). These experiments demonstrate that YFP:CO and CFP:COP1 colocalize and physically interact in the nuclei of plant cells.

The COP1 full-length cDNA was isolated by RT–PCR and produced as entry clone through BP reaction of Gateway system from Invitrogen. Then, the entry clone was utilized for the construction of destination vectors for plant transformation, FRET experiments and in vitro-binding assay. All plasmids for plant transformation were introduced into Agrobacterium strain GV3101 (pMP90RK) and transformed into WT Columbia, cop1–4 or SUC2:CO (An et al, 2004) plants by the floral dip method (Clough and Bent, 1998).

(C) Quantification of FRET in vivo between CFP:CO and YFP:COP1. YFP:CO detected as an increase in CFP fluorescence after photobleaching of YFP. Quantification of FRET efficiencies after acceptor photobleaching measured in nuclei and nuclear speckles. Data are mean±s.d. of 10–20 cells from three separate experiments. (D) Visualization of increase in CFP fluorescence after YFP photobleaching. Left-hand panel, cells expressing CFP:COP1 and YFP, which exerts an effect as a negative control. Right-hand panel, cells expressing CFP:COP1 and YFP:CO. Scale bar: 6 μm in (A) and 8 μm in (D).

Fluorescence resonance energy transfer (FRET) pairs GFP/DsRed were analysed using a confocal laser-scanning microscope (Zeiss LSM510 META). FRET measurements of DsRed emission with zero contribution from GFP, was accomplished as described by Erickson et al. (2003) using the following settings: excitation at 488 nm and emission filters, BP 505–530 nm for GFP and BP 600–637 nm for DsRed.

(C–F) FRET detection in tobacco leaf epidermal cells co-expressing GFP:AtEBP and ACBP4:DsRed; (C) differential interference contrast image of D-F; (D) green channel shows GFP:AtEBP; (E) red channel shows FRET signal of ACBP4:DsRed; (F) co-localization of two signals is indicated by a yellow colour in merged images of (D) and (E).

In FRET analysis, in cells co-expressing GFP:AtEBP and ACBP4:DsRed, not only GFP:AtEBP green fluorescence (Fig. 2D) but also ACBP4:DsRed red fluorescence (Fig. 2E), which overlapped with the GFP signals (Fig. 2F), were detected, indicating that FRET occurred between GFP:AtEBP and ACBP4:DsRed.


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Isothermal titration calorimetry (ITC) confirmed the ability of Munc13–13–150(K32E) to bind to RIM2α82–142 (Figure 4B), yielding an apparent Kd of 0.10 μM and a 1:1 stoichiometry.

Moreover, the gel filtration and ITC data (Figure 4A and4B), as well as the high quality of the1H-15N HSQC spectrum of Munc13–13–150(K32E)/RIM2α82–142 complex and the observation of only one set of cross-peaks (Figure 4C), demonstrate a 1:1 stoichiometry. All these results strongly suggest that the monomer A/RIM2α82–142 complex observed in the crystals faithfully reflects the true Munc13–1/RIM2α binding mode, whereas the presence of a second Munc13–13–150(K32E) molecule in the crystals must be considered a consequence of crystal packing.

The greater binding affinity of AIRE–PHD1 for H3K4me0 peptides was confirmed by both tryptophan fluorescence spectroscopy and isothermal titration calorimetry (ITC), yielding dissociation constants of ∼4 μM, ∼20 μM and >0.5 mM for H3K4me0, H3K4me1 and H3K4me2, respectively (supplementary Fig S2C online; Table 1).

Indeed, fluorescence spectroscopy and ITC assays showed that the alanine mutations R2A in the H3 peptide and D312A in AIRE–PHD1 markedly reduced the binding affinity (Table 1; Fig 4C) without affecting the protein fold (supplementary Fig S3 online).

Similarly, pull-down experiments with whole histones and the H3K4me0 peptide, together with fluorescence spectroscopy and ITC measurements performed on AIRE–PHD1-D297A showed reduced binding (Table 1; Fig 4).

(C) ITC data for binding of H3K4me0 peptide to AIRE–PHD1 (PHD1), AIRE–PHD1-D297A (D297A) and AIRE–PHD1-D312A (D312A). The upper panels show the sequential heat pulses for peptide–protein binding, and the lower panels show the integrated data, corrected for heat of dilution and fit to a single-site-binding model using a nonlinear least-squares method (line). N, Kd, ΔH and ΔS represent measured stoichiometric ratio, dissociation binding constant, differential enthalpy and differential entropy, respectively. AIRE, autoimmune regulator; GST, glutathione-S-transferase; H3K4me0, histone H3 non-methylated at lysine 4; ITC, isothermal titration calorimetry; PHD, plant homeodomain.

The Kd values for the binding of p53 TAD2 to p62 and Tfb1 PH-Ds determined by isothermal titration calorimetry (ITC) are 3175±570 and 391±74 nM, respectively (Di Lello et al, 2006). In the binding of VP16 TAD to Tfb1 PH-D, the Kd value estimated by NMR titration experiment was ∼4000–7000 nM (Di Lello et al, 2005). Compared with these Kd values, the binding of hTFIIEα AC-D to p62 PH-D is rather strong.

The binding affinity of p53 TAD2 to p62 PH-D is regulated by S46 and T55 phosphorylation (Di Lello et al, 2006). The Kd values of p53 TAD2 to p62 PH-D are reported for the unphosphorylated form as 3175±570 nM, for the S46-phosphorylated form as 518±92 nM, for the T55-phosphorylated form as 457±75 nM and for both S46- and T55-phosphorylated form as 97±33 nM. Very recent ITC studies demonstrated the Kd value of hTFIIEα336−439 to p62 PH-D to be 45±25 nM (Di Lello et al, 2008).


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Various forms of gE and the gE-gI heterodimer were subcloned, expressed, and purified from baculovirus-infected insect cell supernatants by nickel affinity and/or IgG affinity and gel-filtration chromatography as described previously [12]. Two recombinant forms of the Fc fragment of IgG1, wtFc and heterodimeric Fc, which contain two and one gE-gI binding sites, respectively, were also produced in CHO cells and purified as described previously [12].

The predicted C2A domain of Munc13–1 encompasses approximately residues 3–130. Using Munc13–1 fragments containing residues 3–132 (Munc13–13–132), 3–150 (Munc13–13–150), 3–209 (Munc13–13–209), and 105–228, we previously showed that the C2A domain is essential for binding to the RIM2α ZF domain (RIM2α82–142), but additional sequences at its C-terminus are necessary for tight binding [20]. In agreement with these conclusions, Munc13–13–150 and Munc13–13–209 (but not Munc13–13–132) largely co-elute with RIM2α82–142 in gel filtration experiments with an apparent molecular weight characteristic of a 1:1 heterodimer (Figure 1B and [20]). Interestingly, the apparent molecular weights observed for isolated Munc13–13–132, Munc13–13–150, and Munc13–13–209 in gel filtration were significantly higher than their monomeric molecular weights (Figure 1B and unpublished data), suggesting that they form stable dimers.

Gel filtration showed that both point mutations disrupt dimerization of Munc13–13–128, Munc13–13–150, and Munc13–13–209, but preserve heterodimerization with RIM2α82–142 (Figure 4A and unpublished data).

Moreover, the gel filtration and ITC data (Figure 4A and4B), as well as the high quality of the1H-15N HSQC spectrum of Munc13–13–150(K32E)/RIM2α82–142 complex and the observation of only one set of cross-peaks (Figure 4C), demonstrate a 1:1 stoichiometry. All these results strongly suggest that the monomer A/RIM2α82–142 complex observed in the crystals faithfully reflects the true Munc13–1/RIM2α binding mode, whereas the presence of a second Munc13–13–150(K32E) molecule in the crystals must be considered a consequence of crystal packing.

The flow-through fraction of the Ni-NTA column was concentrated and applied into a Superdex 200 size-exclusion column (Amersham Biotech) pre-equilibrated with 30 mM Tris-HCl (pH 8.0), 50 mM NaCl, 5 mM DTT.

(A) Quantification of Bax binding to membranes. Bax (100 nM) and tBid (20 nM) or tBid-mt1 were incubated with liposomes in the presence of increasing concentrations of Bcl-XL, Bcl-XL Y101K, or Bcl-XL ΔBH4. Membrane-bound protein was separated from soluble protein by Sepharose CL-2B gel filtration chromatography and quantified by immunoblotting.

Although our results indicate that Bcl-XL inhibits tBid-mediated activation of Bax by sequestering tBid in a stable complex, it is unclear how Bcl-XL inhibits Bax binding to membranes. For example, Bcl-XL might inhibit Bax binding to membranes by preventing it from interacting with tBid. Alternatively or in addition, Bcl-XL might directly inhibit Bax binding to membranes. To determine the mechanism(s) involved, we measured Bax liposome binding by gel filtration chromatography for reactions containing the different mutant proteins (Figure 4A).

The incubation of Bax with liposomes alone does not cause sufficiently tight membrane binding by Bax to survive gel filtration chromatography (Figure 1B).

All sample components (buffers, liposomes, etc.) were added prior to the addition of recombinant proteins, which were added in the order Bcl-XL, Bax, and tBid. Membrane-bound protein was separated from soluble (“free”) protein using gel filtration chromatography on Sepharose CL-2B resin. Membrane binding was measured by comparing the intensities of membrane-bound proteins (fractions 3 and 4) with total proteins (fractions 3 and 4 plus fractions 8–11). Separation of membrane-bound protein from soluble (“free”) protein by liposome floatation on a sucrose density gradient was performed as previously described [57].


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X-ray diffraction studies guided by nuclear magnetic resonance (NMR) experiments reveal the crystal structures of the Munc13–1 C2A-domain homodimer and the Munc13–1 C2A-domain/RIM ZF heterodimer at 1.44 Å and 1.78 Å resolution, respectively. The C2A domain adopts a β-sandwich structure with a four-stranded concave side that mediates homodimerization, leading to the formation of an eight-stranded β-barrel. In contrast, heterodimerization involves the bottom tip of the C2A-domain β-sandwich and a C-terminal α-helical extension, which wrap around the RIM ZF domain. Our results describe the structural basis for a Munc13–1 homodimer–Munc13–1/RIM heterodimer switch that may be crucial for vesicle priming and presynaptic plasticity, uncovering at the same time an unexpected versatility of C2 domains as protein–protein interaction modules, and illustrating the power of combining NMR spectroscopy and X-ray crystallography to study protein complexes.

Guided by solution nuclear magnetic resonance (NMR) experiments, we have solved the X-ray crystal structure of the Munc13–1 C2A-domain homodimer at 1.44 Å resolution, designed a mutation that disrupts homodimerization, and solved the X-ray crystal structure at 1.78 Å resolution of a Munc13–1 fragment bearing this mutation bound to the RIM2α ZF domain.

Moreover, our data uncover an unexpected versatility of C2 domains as protein–protein interaction modules that underlies this switch, and emphasize that combining NMR spectroscopy with X-ray crystallography provides a powerful approach to investigate protein complexes at atomic resolution.

Note also that the mutated Munc13–1 side chain (E32) is close to the interface with the ZF domain, but is facing away, consistent with the conclusion drawn from the NMR analysis that the mutation does not substantially alter the binding mode.

Thus, NMR studies showed that a long α-helix (helix a1) and an SGAWFY motif at the end of the short α-helix at the C-terminus of the RIM2α ZF domain (helix a2) bind to Rab3A, and that the interaction involving the SGAWFY motif is released upon binding of Munc13–1 to the ZF domain. Figure 7B shows that the α-helical extension at the C-terminus of the Munc13–1 C2A domain would have steric clashes with Rab3A upon binding to RIM2α. All these observations lead to the model of Figure 7C, which proposes that formation of the Munc13–1/RIM/Rab3A tripartite complex requires disruption of the Munc13–1 homodimer and a change in the relative orientation of the structural elements of RIM.

Our results illustrate the power of combining X-ray crystallography with NMR spectroscopy to study structural aspects of protein complexes, and suggest that a complex cascade of protein–protein interactions, including a Munc13–1 homodimer–Munc13–1/α-RIM heterodimer switch, may regulate synaptic vesicle priming and some forms of presynaptic plasticity.

(E) Superposition of the structure of the RIM2α ZF domain observed in the heterodimer (blue) and its solution structure determined in isolation by NMR spectroscopy (red) [20].

To confirm the specificity of AIRE–PHD1 for H3K4me0, we compared the binding of histone H3 N-terminal peptides—H3K4me0, H3K4me1, H3K4me2 and H3K4me3—to AIRE–PHD1 by using two dimensional 1H-15N nuclear magnetic resonance (NMR). A discrete set of chemical shift changes was observed on addition of all four histone H3 peptides to AIRE–PHD1 (supplementary Fig S2A,B online). However, the intensity of the changes was inversely related to the methylation level of the H3 peptide: the H3K4me0 peptide induced the largest changes (maximum average chemical shift change Δδmaxav=0.9 p.p.m.; Fig 2). The addition of H3K4me0 and H3K4me1 peptides resulted in chemical shift changes in the slow- to intermediate-exchange regime (supplementary Fig S2A online), indicating low micromolar binding affinities. By contrast, the NMR data on addition of H3K4me2 and H3K4me3 peptides were in the fast-exchange regime, indicating millimolar binding affinities (supplementary Fig S2B online).

The mapping of the H3/AIRE interaction site uniquely to AIRE–PHD1 was further confirmed by NMR titrations of histone H3 peptides into AIRE–PHD2, which bound neither methylated nor H3K4me0 peptides (data not shown).

Accordingly, the amides of C310, L308 and G306 showed high protection factors in NMR deuterium exchange experiments, confirming their involvement in H-bonds (Fig 3B).

The formation of salt bridges between the side chains of R2 and D312, and between K4 and D297 seemed to be crucial for binding specificity, as indicated experimentally by the large NMR chemical shift changes for G313 (near to D312) and D297 (Fig 2).

The model of AIRE–PHD1 complexed with H3K4me0 was in perfect agreement with the experimental chemical shift perturbation data, as the peptide-binding region coincided with the binding surface identified by NMR spectroscopy (Fig 3A). In fact, the H3K4me0 peptide induced chemical shift changes in AIRE–PHD1 residues that map only on one side of the protein surface, involving residues in the N terminus of the PHD finger, the first β-strand, and the loop connecting the first and the second β-strands (D297, G305, G306, L308, C310, D312 and G313; Fig 2; supplementary Fig S5 online). A similar pattern of chemical shift changes indicated the same binding site for H3K4me1.

Furthermore, two tertiary structures of a zinc-finger domain of hTFIIEα (Okuda et al, 2004) and a winged helix/forkhead domain of hTFIIEβ (Okuda et al, 2000) have been solved by NMR spectroscopy.

We have determined structures of both free hTFIIEα AC-D and its form bound to the PH-D of hTFIIH p62 by using NMR spectroscopy. The structures reveal that hTFIIEα AC-D recognizes p62 PH-D tightly through a combination of hydrophobic and electrostatic interactions.

To characterize the precise interaction at the molecular level, we first solved a solution structure of the AC-D of hTFIIEα using NMR spectroscopy (Figure 1B and Table I).

These interacting amino acids were also observed in the NMR titration experiments. In hTFIIEα AC-D, the NMR signals of E386, F387, E388, E389, V390, A391 and D392 were changed significantly upon addition of p62 PH-D (Supplementary Figure 1B) and also in p62 PH-D the NMR signals of K19, Q53, K54, I55, S56, E58, K60, A61, I63, Q64, L65, Q66, T74, T75 and F77 were changed by adding hTFIIEα AC-D (Supplementary Figure 2B).

Recently, the structure of a complex of the PH-D of Saccharomyces cerevisiae Tfb1, a homologue of human p62, with a TAD2 of activator p53 was determined by NMR spectroscopy (Di Lello et al, 2006).

To investigate whether the STDE is involved in the binding, we prepared a longer construct (residues 351–439) containing both acidic regions and performed the NMR titration experiment under the same conditions (Supplementary Figure 4). The result was that the NMR signals of STDE showed no significant changes and the Kd of 400±43 nM was almost the same as that estimated using hTFIIEα AC-D.

The binding site of hTFIIH p62 PH-D was localized to the second β-sheet (S5, S6 and S7), the loops between S1 and S2 and between S5 and S6 and the C-terminal H1 helix, where a substantial positive cluster is formed. Therefore, it is reasonable to speculate that the N-terminal highly acidic tail of hTFIIEα AC-D strongly binds to the positively charged surface of hTFIIH p62 PH-D. This is supported by the result that the binding is strengthened by removing NaCl from the buffer in the NMR titration experiments.

While this manuscript was in preparation, an independent study reported the NMR structure of the C-terminal CH domain of α-parvin in complex with a 10-residue peptide derived from the paxillin LD1 motif (Wang et al., 2008).

This analysis together with NMR studies of a spin-labeled LD1 peptide supports the surprising finding that LD motifs can associate with a single binding site bidirectionally.

However, we previously demonstrated that LD-binding to the FAT domain of FAK does not reside in a local peptide sequence such as the PBS (Hoellerer et al., 2003) and thus investigated the interaction of α-parvin-CHC with LD motifs using solution NMR. 1H-15N HSQC monitored titrations of 15N-enriched α-parvin-CHC were performed with peptides representing all five paxillin LD motifs. Each peptide was found to induce resonance-specific chemical shift perturbations (Figure 2 and data not shown), indicating an interaction with α-parvin-CHC.

As measured by NMR, none of these substitutions substantially altered the binding affinity for α-parvin-CHC, indicating that the predicted electrostatic contacts are energetically neutral under the assay conditions (50 mM sodium phosphate and 100 mM NaCl [pH 6.9]) used.

The resulting KD-value for α-parvin-CHC is similar to the corresponding value obtained by NMR, supporting the validity of our results (Table 2). Importantly, the KD-values for α-parvin-CHC and full-length α-parvin are the same within error (Table 2), suggesting that the N-terminal region of α-parvin (residues 1–242) makes little net contribution to LD binding. We thus conclude that the crystal structures of the α-parvin-CHC/LD complexes along with the solution NMR studies presented here provide a relevant description of LD recognition by α-parvin.

This binding site is consistent with the one described in a recent solution NMR structure of α-parvin-CHC in complex with an LD1 peptide (Wang et al., 2008). Using NMR titrations, we have shown that this binding site interacts with all five paxillin LD motifs, exhibiting a preference for LD1, LD2, and LD4 over the less conserved LD3 and LD5. This could be due to differences in specific contacts and/or helical propensity of the latter two LD motifs.

Recent NMR studies (Zhang et al., 2008) have also demonstrated that the single LD binding site of PKL/GIT1 binds to both LD4 (Turner et al., 1999) and LD2. Taken together, these observations suggest that paxillin LD motifs are promiscuous protein interaction modules.

Data were recorded on home-built or Bruker spectrometers with 11.7, 14.1, 17.6, and 22.3 T field strengths and processed with NMRPipe (Delaglio et al., 1995). Backbone chemical shift assignments were obtained using standard triple resonance experiments. Peptide titration experiments were performed by mixing two stock solutions (in 50 mM sodium phosphate, 100 mM NaCl, 2 mM DTT, 5% D2O, and 30 μM DSS [pH 6.9]) containing 235 μM 15N-enriched α-parvin-CHC and either no or a maximum concentration of LD peptide at the required protein/ligand ratios (Figure S2). Phase-sensitive gradient-enhanced 1H-15N HSQC spectra (Kay et al., 1992) were recorded at 25°C.


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To map the phosphorylation site systematically, we decided for a peptide array that displays 73 peptides containing all serine and threonine residues of DDX3X (Supplementary Figure 5).

Incubation of the peptide array with purified TBK1 produced a number of phosphorylation signals (Figure 7A). The TBK1 control peptide was phosphorylated by TBK1, suggesting that the activation loop is indeed an autophosphorylation site.

Analysing the remaining hits in the peptide array, we identified 11 potential phosphorylation sites in 9 DDX3X-derived peptides (Figure 7A). Four of these target sites (S181, S183, S240 and S269) were found in the DEAD domain that contains the ATPase activity of DDX3X. The seven remaining sites were scattered throughout the helicase domain (S429, T438, S442, S456, S520, T542 and S543). We used the information gathered in the peptide array to derive a TBK1 consensus phosphorylation site (Figure 7B). Strikingly, TBK1 showed a strong preference for serine over threonine. In fact, none of the 12 peptides displayed on the array that contain exclusively threonines was phosphorylated.

Mapping of the TBK1 phosphorylation site in DDX3X. (A) Peptides derived from DDX3X or control peptides derived from TBK1 or IRF3 along with alanine mutants were incubated with TBK1 in the presence of [γ-32P]ATP. Each array contained a total of 82 peptides spotted in triplicate. (B) Phosphorylation sites obtained from the peptide array (see also text) were used to build a TBK1 phosphorylation consensus sequence.


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To test whether ephrin-B1 and Cx43 physically interact, we performed a pull-down assay in NIH 3T3 cells expressing ephrin-B1. We used a recombinant protein consisting of the extracellular domain of Eph-B2 receptor fused to the Fc fragment of human IgG (EphB2-Fc) to pull down ephrin-B1. Cx43 was detected in the pull down, indicating that it interacts with ephrin-B1 (Figure 6Ca).

To identify the domain of ephrin-B1 required for the interaction with Cx43, we performed a pull down using a recombinant protein consisting of the extracellular domain of ephrin-B1 fused to the Fc fragment of human IgG (ephrinB1-Fc). Cx43 was not detected in the pull down indicating that the intracellular domain of ephrin-B1 is required for the interaction with Cx43 (Figure 6Cb).

Cx43 could be detected in the pull downs from cells expressing either ephrin-B1 wild type or ephrin-B1ΔPDZ, however, the relative abundance of phosphorylated versus unphosphorylated band was changed. More phosphorylated Cx43 (slower mobility) was observed in the ephrin-B1 wild-type pull down, whereas more unphosphorylated Cx43 was detected in the ephrin-B1ΔPDZ pull down (Figure 7A). These results indicate that the PDZ binding domain of ephrin-B1 is not required for its interaction with Cx43; however, ephrin-B1ΔPDZ and wild-type ephrin-B1 interact preferentially with different forms of Cx43.

In our pull-down assay, wild-type ephrin-B1 interacted preferentially with phosphorylated Cx43 whereas ephrin-B1ΔPDZ interacted preferentially with unphosphorylated Cx43, suggesting that the interaction between ephrin-B1 and Cx43 might not be direct, and that these proteins might interact differently when at the cell surface or in the cytoplasm.

To assess whether MELK has a role in mammary carcinogenesis, we knocked down the expression of endogenous MELK in breast cancer cell lines using mammalian vector-based RNA interference. Furthermore, we identified a long isoform of Bcl-G (Bcl-GL), a pro-apoptotic member of the Bcl-2 family, as a possible substrate for MELK by pull-down assay with recombinant wild-type and kinase-dead MELK.

To investigate the biological functions of MELK in breast cancer cells, we searched for substrates of MELK in cancer cells by in vitro protein pull-down assays using wild-type MELK (WT-MELK) and kinase-dead MELK (D150A-MELK) recombinant proteins. Comparison of silver staining of SDS-PAGE gels containing the pulled-down proteins identified an approximately 30 kDa protein in the lane corresponding to proteins pulled-down with WT-MELK but not in that corresponding to proteins pulled-down with D150A-MELK (Figure 3a).

Furthermore, we demonstrated that His-tagged WT-MELK could pull-down with Bcl-GL but His-tagged D150A-MELK could not, indicating that, in vitro, Bcl-GL interacts directly with WT-MELK but not with D150A-MELK (Figure 3d).

Thus, to investigate the biological significance of MELK in breast cancer cells, we searched for a possible substrate(s) of MELK by means of in vitro pull-down assays with recombinant wild-type MELK (WT-MELK) and kinase-dead MELK (D150A-MELK).

(E) The membrane fraction of adult brain lysates was incubated with or without Flag-tagged Cdk5. Flag-tagged Cdk5 pulled down TrkB from the membrane fraction of adult brain lysates.

Quality of the purified GST and GST-fusion proteins used in the GST pull-down assay was verified by Coomassie blue staining.

GST-pull down and co-immunoprecipitation assays were performed to further characterize this interaction.

GST-pull down assay to identify the region of BRCA1 interacting with PP1. (A) Fragments used in GST pull down assays (BR 2 to 5) are diagrammed. (B) Gel depicting co-precipitation of GST-bound BR-4 with PP1. Following incubation of GST-BRCA1 proteins with equal amounts of cell lysate, a western blot was performed and probed with an antibody to the catalytic region of PP1. (C) Analysis of the effect of mutations of the KVTF PP1 interacting domain on the BRCA1- PP1 interaction. GST-bound-BR4 V-A and GST-bound-BR4 F-A binds PP1 with decreased intensity, compared to WT GST-bound-BR4.

To study APPL1−Rab5 interaction in solution, we performed glutathione S-transferase (GST)-mediated pull-down assays. The APPL1 BAR-PH domain (residues 5−385) and a longer fragment with a 40-residue extension downstream of the PH domain, APPL1 (5−419), were each effectively pulled down by GTP-bound GST−Rab5 fusion protein (Figure 4). The APPL1 protein was pulled down by either WT Rab5 preloaded with non-hydrolysable GTP analog (GppNHp) or Rab5-Q79L defective in GTP hydrolysis (with or without preloaded GTP analog), but could not be effectively pulled down by either the WT Rab5 preloaded with GDP or Rab5-S34N defective in GTP binding (Figure 4 and data not shown).

Therefore, we tested APPL1 binding specificity towards other members in the Rab5 subfamily, using GST−Rab21 (full length) and GST−Rab22 (2−192) to pull down APPL1 (5−419).

On the other hand, we were unable to detect any binding between APPL1 and Rab22 in the pull-down assay (Figure 4).

To further define the Rab5−APPL1 binding mode, we performed extensive pull-down analyses between variants of Rab5 and APPL1, looking for reversal mutants that could rescue the lost binding ability of others. We identified one such pair; APPL1-N308D abolished the binding to Rab5, while Rab5-L38R had no effect on APPL1 binding. However, Rab5-L38R was found to bind with APPL1-N308D, but not with the other tested APPL1 variants of similar hydrophobic-to-charged mutations, including V25D, A318D, and L321D (Supplementary Figure 6). This result suggests that Rab5-L38R restores binding for APPL1-N308D through complementary, electrostatic, yet specific interactions. It further implies that the position 308 in the β3 strand of APPL1 PH domain is in the vicinity of position 38 in the α1 helix of Rab5 in their complex.

Combined results from our mutagenesis pull-down experiments (Figures 4, 5 and 6), crystal structures of the BAR-PH domain of APPL1 (Figure 1), and structures of GTPase domain of human Rab5 in different nucleotide binding modes (Zhu et al, 2003, 2004) clearly explain the requirement of GTP-bound Rab5 for APPL1 binding. Based on available information, we have modeled the interaction between the two proteins. With the assumption that both proteins remain rigid bodies, our complex model satisfies constraints imposed by the mutagenesis pull-down results (Figure 7).

In the Rab5−APPL1 pull-down experiment, 30 μg GST−Rab fusion protein (52 kDa) was incubated with 60 μl of 30% slurry of GSH–Sepharose 4B (GE Healthcare) at 22°C for 30 min. Nucleotide loading reaction was performed on the GSH beads in an exchange buffer of (1 × PBS, 2 mM DTT, 1 mM MgCl2, 4 mM EDTA, and 400 μM GppNHp or GDP) at 22°C for 30 min. Increasing the magnesium ion concentration to 20 mM terminated the loading reaction.

Pull-down analysis of APPL1-Rab interaction. GST fusion proteins of Rab5, Rab21, and Rab22 were used to pull down His-tagged APPL1 (5−385) and (5−419) fragments in the presence of GDP or GTP analog (GppNHp).

(A) Pull-down assay on Rab5 variants. Point mutations in switch regions were introduced in the full-length Rab5-Q79L background of the GST fusion construct. Each mutant was expressed in E. coli, purified, and equal amounts of each Rab5 sample was used to pull down recombinant proteins of His-tagged WT APPL1 (5−419) (top panel) and His-tagged rabaptin5 (551−862) (bottom panel). The results were visualized by Coomassie blue stain.

Consistent with prior reports [23,24], we failed to observe stable direct interaction between purified ST and PP2A C subunit using a glutathione S-transferase (GST) pull-down assay (unpublished data).

To assess which of the PDE4 subtypes contribute to the activity recovered in the β1AR IP, cardiomyocytes deficient in PDE4A, PDE4B, and PDE4D were subjected to pull-down experiments.

In pull-down experiments using purified proteins (Figure 3D and E), β1AR efficiently interacts with PDE4D, whereas β2AR has negligible affinity for PDE4D, underscoring the direct mode of PDE4D–β1AR interaction versus the indirect, β-arrestin-dependent mode of PDE4D–β2AR interaction.

In agreement with the GST fusion pull-down experiments, fluorescence spectroscopy showed no binding of H3K4me0 to AIRE–PHD1 containing the APECED-causing C311Y mutation (Bjorses et al, 2000). Nevertheless, a second pathological mutant, V301M (Soderbergh et al, 2000), was still able to bind to H3K4me0, indicating that this mutation is not located in the H3 interaction site (Table 1).

Similarly, pull-down experiments with whole histones and the H3K4me0 peptide, together with fluorescence spectroscopy and ITC measurements performed on AIRE–PHD1-D297A showed reduced binding (Table 1; Fig 4).

(A) Pull-down assay of AIRE–PHD1 (PHD1) and AIRE–PHD1-D297A (D297A) mutant proteins with histones.

To identify the region of Plx1 that binds to the Mcm complex, we used truncated proteins fused to GST in pull-down experiments. One of the truncated proteins contained the protein interaction domain known as the Polo box, which has been shown to interact with phosphorylated proteins (Elia et al, 2003). The GST–Polo box-containing fragment derived from Plk1, which is highly similar to the Polo box of Plx1, was incubated with egg extracts. Proteins bound to the Polo box domain were then eluted using a phosphorylated peptide with a strong affinity for the Polo box, and subsequently subjected to matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) to identify the sequence of the interacting proteins. As shown in Figure 5D, the GST–Polo box was able to pull down four major proteins, Mcm2, Mcm4, Mcm6 and Mcm7, as revealed by MALDI-TOF analysis. As the Polo box interacts with phosphorylated residues and the Mcm proteins could be eluted using a phosphorylated peptide, it is likely that the interaction is mediated by phosphorylated serine/threonine residues present on endogenous Mcm proteins. The binding of the Polo box to endogenous Mcm proteins was consistent with the ability of full-length Plx1 to interact with chromatin and Mcm7. We then tested whether the Polo box binding to Mcm proteins was enhanced by the activation of checkpoint. In line with the results obtained with full-length Plx1, Polo box binding to the Mcm complex was enhanced by checkpoint activation and was inhibited by caffeine (Figure 5E).

(D) Pull-down with GST–Polo box and MALDI-TOF analysis. Interphase extracts were incubated with GST–agarose beads or agarose beads with GST fused to the Polo box derived from human Plk1. After pull-down, proteins were consecutively eluted using 3 mg/ml of phosphopeptide solution (first elution=E1, second elution=E2, see Supplementary data for the sequence of the peptide and the Polo box) and subjected to MALDI-TOF analysis. Band 1 corresponds to Mcm2, band 2 to Mcm6, band 3 to Mcm4 and band 4 to Mcm7. The GST fusion protein containing the Polo box from Plk1 is shown at the bottom. (E) GST–Polo box pull-down of nuclear proteins probed with anti-Mcm7 antibodies (GST–Polo box). The extract used for the pull-down was supplemented with nuclei and buffer (Buff), 2.5 μM aphidicolin (APH) or 2.5 μM aphidicolin and 5 mM caffeine (Caff), as indicated. The total amount of unbound Mcm7 extracted from nuclei was loaded as a control (Input).

To confirm this and to identify the AC-D-binding region in p62, we performed a GST pull-down assay using hTFIIEα AC-D and GST-fused p62 deletion mutants (Figure 2A). After purification by glutathione-Sepharose column chromatography, all samples containing the C-terminal region, namely full-length GST–p621−548, GST–p62109−548, GST–p62238−548 and GST–p62333−548, were considerably degraded or incompletely translated (data not shown).

(B) GST pull-down assay of hTFIIEα AC-D with wild type and p62 mutants. Lane 1, GST; lane 2, GST–p62 (1–548aa); lane 3, GST–p62 (1–108aa); lane 4, GST–p62 (1–238aa); lane 5, GST–p62 (1–333aa); lane 6, GST–p62 (109–548aa); lane 7, GST–p62 (238–548aa); lane 8, GST–p62 (333–548aa); lane 9, 20% input.

Finally, the interaction of Vav3 and ERα was assessed by GST pull-down analysis.

For pull down reaction, 5~10 ug of GST or GST-Vav3-DH+PH was incubated with 1 mg of cell extracts from MCF7 cells in a binding buffer [20 mM of Tris.CL, PH.

We performed GST pull down experiment to confirm the interaction between Vav3 and ERα. A GST fusion protein including the DH and PH domain of Vav3 (GST-Vav3-DH+PH) was generated (Figure 7A). Cell extract derived from MCF7 cells was incubated with immobilized GST-Vav3-DH+PH fusion protein or GST protein. Then, the pull down samples were fractionated by SDS-PAGE (Figure 7B) and subjected to western blot analysis for ERα. We found that GST-Vav3-DH+PH fusion protein, but GST protein, interacted with ERα (Figure 7C). In summary, we found that Vav3 contains NLS in the PH domain and three LXXLL motifs in the DH domain.

Vav3 complexes with ERα by GST pull down analysis. (A) GST-Vav3-DH+PH fusion protein. (B) GST-Vav3-DH+PH fusion protein (lane 3 and 4) and control GST protein (lane 1 and 2) were subjected to pull down reaction in the absence (lane 1 and 3) and presence (land 2 and 4) of cell extract derived from MCF7 cells.

Next, we used the recombinant, purified MRN complex in peptide pull-down experiments. These showed that MRN bound to the phosphorylated but not the unphosphorylated form of the SDTD peptide—confirming the direct nature of the interaction—yet did not bind to either version of the H2AX C-terminal peptide (Fig 3A). This indicates the specificity of MRN for the phosphorylated MDC1 SDTD motif and also shows that if, as previously reported, NBS1 binds to γH2AX directly (Kobayashi et al, 2002), this interaction must be less stable than MRN binding to the phosphorylated SDTD motif.

(B) Silver-stained SDS–polyacrylamide gel of an SDTD peptide pull-down. PP, S329 T331 doubly phosphorylated peptide and −, its non-phosphorylated equivalent; B, bead-interacting proteins removed from extracts in a pre-clearing step; M, molecular weight markers.

(B) A 200 ng portion of CK2-phosphorylated or mock-phosphorylated GST-SDTD6 was incubated in pull-down reactions with 25 μl of in vitro-translated (IVT) HA fusions corresponding to amino-acid residues 1–348 of NBS1 (HA-fNBS1), or analogous proteins bearing point mutations predicted to abolish either FHA (R28A/H45A) or BRCT2 (K160M) phosphorylation-dependent interactions.

For the glutathione S-transferase (GST) pull-down experiments, plasmids pGEX-RNP-K (kindly provided by Dr. Levens) and pGEX-4T (GE Healthcare) were used to express GST-hnRNP-K fusion protein or GST alone in Escherichia coli.

GST-hnRNP-K and GST proteins were produced in E. coli BL21 cells, previously transformed with vector pGEX-RNP-K or pGEX-4T. Cells were induced with 0.1 mM IPTG for 2 h at 37 °C. Bacteria were harvested and suspended in lysis buffer (PBS, 1% Triton X-100, 1 mM PMSF, 5 mM DTT, and anti-proteases), and sonicated on ice. GST-hnRNP-K and GST alone were purified from cleared lysates by mixing with glutathione-sepharose 4B beads (GE HealthCare), 5 ml of cleared lysate/400 μl of beads, for 1 h at 4 °C. After extensive washing, GST-hnRNP-K or GST beads were incubated in binding buffer (50 mM HEPES, ph 7.5, 50 mM NaCl, 0.1% Nonidet P-40 with protease inhibitor mixture (Roche Molecular Biochemical)) at 4 °C for 1 h with insect cell extracts containing either p30 or p54 ASFV proteins overexpressed in a baculovirus system [8]. Equal amounts of GST, GST-hnRNP-K and ASFV proteins p30 and p54 were used as judged by Coomasie Blue staining.

The interaction was further confirmed by in vitro binding assays using a GST-hnRNP-K fusion protein bound to glutathione-sepharose 4B beads. GST pull-down experiments were carried out followed by Western blot with specific antibodies. First, p30 and another unrelated ASFV protein (in this case, p54 as negative control) were used to bind GST fusion protein or GST alone. Protein p30, and not p54, was retained in the presence of GST-hnRNP K, showing a band of appropriate size (30 kDa) for p30 in Western blotting. This band did not appear in the presence of GST alone, indicating specific interaction of p30 with hnRNP K and not with GST (Fig. 1A). Identical results were obtained in subsequent experiments using BA71V infected or mock-infected cells extracts instead of baculovirus infected cell extracts in the pull-down assay (Fig. 1B).

This interaction was further confirmed by an in vitro GST-fusion pull-down assay, using either p30 obtained from baculovirus system or ASFV infected cell extracts.

In contrast, previous data based on less-sensitive pull-down experiments suggested that α-parvin binding is limited to LD1 and LD4 (Nikolopoulos and Turner, 2000).

These immobilized template assays also allowed us to determine whether those basal transcription factors travelling with RNApII in Ctk1-deficient cells remain stably associated with polymerase after it runs off the template. A single round of transcription on the HIS4 template was allowed, and RNApII was immunoprecipitated from the supernatant through the Rpb3 subunit (Figure 5E). Although we can efficiently pull down Rpb3 itself, we failed to coimmunoprecipitate any of the basal transcription factors that cross-linked throughout genes in ctk1Δ cells (Figure 5F).


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Consistent with this, our analytical ultracentrifugation (AUC) data showed that BAR-PH protein has a higher dimerization affinity in solution (kd=0.34 nM) than BAR domain alone (kd=0.13 μM; Supplementary data).


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To determine whether isolated CgE (gE residues 213–390, where residue 1 is the first residue of the hydrophobic leader peptide in the immature gE and residue 420 is the first residue of the predicted transmembrane region) binds Fc and to measure the affinity of the interaction, we performed a surface plasmon resonance assay using two forms of recombinant Fc derived from human IgG1: wild-type Fc (wtFc) and nbFc, which contains six point mutations in the CH2-CH3 linker that abrogate binding to gE-gI (Met252 to Gly, Ile253 to Gly, His310 to Glu, His433 to Glu, His435 to Glu, and Tyr436 to Ala) [12]. The Fc proteins were immobilized on the surface of a biosensor chip, and the binding of CgE was assayed at pH 8 and pH 6.

Surface plasmon resonance studies were performed using a BIAcore 2000 instrument (Biacore, Uppsala, Sweden). In this system, binding between a molecule coupled to a biosensor chip (the “ligand”) and a second molecule injected over the chip (the “analyte”) results in changes in the surface plasmon resonance signal that are read out in real time as resonance units [54]. wtFc and nbFc, both derived from human IgG1, were purified from CHO cell supernatants as described previously [12] and immobilized on a CM5 biosensor chip (Biacore) using primary-amine coupling as described by the manufacturer. Equilibrium binding data for CgE binding to wtFc (coupling density, 440 resonance units), nbFc (coupling density, 405 resonance units), and a mock-coupled flow cell were collected for a CgE concentration series (three-fold dilutions of CgE from 30 μM to 5 nM) at pH 8 (50 mM HEPES [pH 8.0], 150 mM NaCl, 3 mM EDTA, 0.005% [vol/vol] P-20 surfactant) and pH 6 (50 mM sodium phosphate [pH 6.0], 150 mM NaCl, 3 mM EDTA, 0.005% [vol/vol] P-20 surfactant). After each injection of CgE, a 30-s injection of 250 mM di-ammonium hydrogen citrate (pH 5.0) was used to disrupt the interaction and restore the surface.

Binding affinity between full-length Rab5-Q79L and APPL1 (5−419) was quantitatively determined in a surface plasmon resonance (SPR) experiment. Rab5 was coupled to the SPR biosensor chip in random orientations, and APPL1 (5−419) was applied as the analyte at concentrations of 0.15−12 μM (Supplementary Figure 4). The dissociation constant, kd, for the Rab5−APPL1 interaction measured from this experiment was 0.9 (±0.7) μM, with kon and koff of 1.3 (±0.6) × 103 M−1 s−1, and 1.2 (±0.4) × 10−3 s−1, respectively. This kd value is typical for an interaction between a Rab and its effectors (Eathiraj et al, 2005).


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The binding mode observed for Fc and the CgE portion of gE-gI in the gE-gI/Fc crystal structure was verified by an independent theoretical prediction of the CgE interaction with Fc. The gE-gI/Fc model resulting from prediction and crystallographic methods is consistent with biochemical data characterizing the interaction, provides insight into the molecular basis for the observed pH dependence of the gE-gI/Fc interaction, and allows mapping of CgE residues important for IgG binding and cell-to-cell spread. In addition, the orientation of the gE-gI/IgG complex on a membrane, as predicted from the complex crystal structure, demonstrates that antibody bipolar bridging can occur on the surface of an infected cell or virus.

Crystallization trials were conducted for various forms of CgE, gE, and gE-gI (including CgE [residues 213–390], gE [residues 21–419], gE2 [residues 21–390], gE-gI [gE plus gI residues 21–266], gE2-gI2A [gE2 plus gI residues 21–208], and gE2-gI2B [gE2 plus gI residues 21–201]) both alone and complexed with wtFc or heterodimeric Fc. The only isolated protein to crystallize was CgE (described above), and the only complex of the six possible gE-gI/Fc complexes that crystallized was one that contained gE residues 21–419 and gI residues 21–266 and wtFc (residues 223–447). The complex crystals grew from drops containing a 2:1 molar ratio of gE-gI and wtFc mixed with an equal volume of well solution (0.1 M MES [pH 6.0] or 0.1 M HEPES [pH 7.0] and 0.9–1.1 M sodium malonate), resulting in a final pH of approximately 7.5. Microseeding increased the reproducibility of crystal growth.

(C) Stereo superposition of the crystallographically determined CgE/Fc complex and the complex predicted with RosettaDock [34] using the structures of CgE and Fc.

X-ray diffraction studies guided by nuclear magnetic resonance (NMR) experiments reveal the crystal structures of the Munc13–1 C2A-domain homodimer and the Munc13–1 C2A-domain/RIM ZF heterodimer at 1.44 Å and 1.78 Å resolution, respectively. The C2A domain adopts a β-sandwich structure with a four-stranded concave side that mediates homodimerization, leading to the formation of an eight-stranded β-barrel. In contrast, heterodimerization involves the bottom tip of the C2A-domain β-sandwich and a C-terminal α-helical extension, which wrap around the RIM ZF domain. Our results describe the structural basis for a Munc13–1 homodimer–Munc13–1/RIM heterodimer switch that may be crucial for vesicle priming and presynaptic plasticity, uncovering at the same time an unexpected versatility of C2 domains as protein–protein interaction modules, and illustrating the power of combining NMR spectroscopy and X-ray crystallography to study protein complexes.

Guided by solution nuclear magnetic resonance (NMR) experiments, we have solved the X-ray crystal structure of the Munc13–1 C2A-domain homodimer at 1.44 Å resolution, designed a mutation that disrupts homodimerization, and solved the X-ray crystal structure at 1.78 Å resolution of a Munc13–1 fragment bearing this mutation bound to the RIM2α ZF domain.

Moreover, our data uncover an unexpected versatility of C2 domains as protein–protein interaction modules that underlies this switch, and emphasize that combining NMR spectroscopy with X-ray crystallography provides a powerful approach to investigate protein complexes at atomic resolution.

Our results illustrate the power of combining X-ray crystallography with NMR spectroscopy to study structural aspects of protein complexes, and suggest that a complex cascade of protein–protein interactions, including a Munc13–1 homodimer–Munc13–1/α-RIM heterodimer switch, may regulate synaptic vesicle priming and some forms of presynaptic plasticity.

The structural information available on protein–protein interactions involving ZF domains is also limited [25]. The X-ray structure of the Munc13–1/RIM2α heterodimer described here provides the first high-resolution view of a complex directly involving a member from the family of ZF domains that includes α-RIMs and other Rab effectors. The structure shows that two surfaces at opposite sides of the RIM2α ZF domain interact with two different structural motifs of Munc13–13–150. Thus, the N-terminal loop region of the RIM2α ZF domain binds to the tip of the C2A-domain β-sandwich, whereas the crevice formed by the C-terminal β-hairpin and helix a2 binds the C-terminal helix of Munc13–13–150. Interestingly, in the complex between the α subunit of Hif-1 (Hif-1α) and the TAZ1 domain of CREB-binding protein (a ZF domain unrelated to the RIM2α ZF domain), Hif-1α also wraps around the TAZ1 domain, interacting with surfaces at opposite sides of the domain [31]. An emerging theme from these observations is that, because of their small size, ZF domains may often use multiple surfaces to increase the affinity and specificity of interactions with target proteins, although further research will be necessary to assess the generality of this notion.

Based on the APPL1 BAR-PH crystal structure, we find that, in general, the PH domain is more conserved than the BAR domain, and most of the highly conserved positions are located closer to the BAR−PH interface rather than the central region of the symmetric dimer.

We have determined the crystal structure of full-length SV40 ST in complex with the full-length Aα subunit of PP2A. This structure reveals two novel zinc-binding motifs formed by the unique C-terminal domain, the structural linkage of the J and unique domain of ST, and the interaction site of ST with the structural A subunit. Together with our biochemical data, we provide a structural basis for understanding the tumorigenic activity of ST protein.

Crystal structure of the complex was determined by a combination of molecular replacement, using the PP2A A subunit structure as the searching model, and single-wavelength anomalous dispersion of intrinsic zinc atoms in ST, and was refined at 3.1 Å resolution (Table 1).

Secondary structures in the determined crystal structure are indicated above the aligned sequences. Solid and empty stars indicate residues interacting with PP2A A subunit using side-chain and main-chain, respectively. Solid and empty squares represent residues involved in interactions between the J domain and the unique domain with side-chain and main-chain atoms, respectively.

Structural comparison of the A-ST crystal structure with previously reported crystal structures of A, AC complex, and A-B56-C complexes [25,26,45,46] also support the conclusion that HEAT repeats 2–10 and HEAT repeats 13–15 form two relatively rigid blocks. However, there is substantial structural flexibility between these two structural blocks, due to the result of accumulative conformational changes in HEAT repeats 10–13 (Figures S1 and S2).

Here we report the crystal structure of SV40 ST in complex with the murine PP2A A subunit. It is striking that all four A-ST complexes in the asymmetric unit of our crystal lattice have essentially the same structure, except the flexible HEAT repeats 11–15 of the A subunit that are not involved in the A-ST interaction. This observation argues strongly that the ST structure as well as the A-ST interface observed in our crystal structure are independent of crystal packing and should be physiologically relevant.

In our ST crystal structure, instead of forming a binuclear cluster, these two zinc ions are located in two separate positions and form two novel zinc-binding motifs. The first and second cysteine clusters (residues 111–116, 138–143, respectively) form two zinc-binding motifs together with the conserved Cys103 and His122, respectively. Both zinc ions interact with helix α5 and stabilize the structure of the C-terminal unique domain.

However, the first zinc-binding motif does not interact with the A subunit of PP2A in the crystal structure. Instead, the crystal structure suggests that the first zinc-binding motif may directly interact with the C subunit near its active site, since the first zinc-binding motif is spatially close to the active site of the PP2A C subunit in the structural superposition of PP2A and A-ST complexes (Figure 5).

In addition to the inhibition of the phosphatase activity of the PP2A AC dimer, the J domain may also play a role in the oncogenic activity of ST by providing an additional binding site for Hsp70, even when in complex with the PP2A AC complex, as suggested by our crystal structure.

Here, we report the first crystal structure of the C-terminal calponin homology domain (CHC) of α-parvin at 1.05 Å resolution and show that it is able to bind all the LD motifs, with some selectivity for LD1, LD2, and LD4.

We present the first high-resolution crystal structure of α-parvin-CHC and show that it interacts with all five paxillin LD motifs in solution. Three cocrystal structures of α-parvin-CHC with 20-residue peptides representing LD1, LD2, and LD4 allow us to characterize the interaction at atomic resolution and to highlight binding-induced conformational changes in α-parvin-CHC.

To elucidate the molecular basis for LD recognition by α-parvin-CHC, we cocrystallized α-parvin-CHC with peptides representing the high-affinity ligands LD1, LD2, and LD4. The corresponding structures at 2.1 Å, 1.8 Å, and 2.2 Å resolution, respectively, were solved by molecular replacement with the α-parvin-CHC apo structure (Table 1).

We have applied a combination of high-resolution X-ray crystallography and solution techniques to characterize the recognition of paxillin LD motifs by α-parvin. Our results demonstrate that full-length α-parvin contains a single LD binding site, formed by the N-linker helix, helix A, and helix G of the C-terminal type-5 CH domain.


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The proteins in the supernatant were subjected for SDS-PAGE, which was visualized by Coomassie Blue staining.

The samples were fractionated in SDS-PAGE and stained with Coomassie Blue.


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The ChIP assays were performed in terms of A20, IκBα, or GAPDH promoters using antibodies and corresponding primers as described.

To substantiate this finding, we took advantage of the chromatin immunoprecipitation (ChIP) assay to examine whether binding of NF-κB to its endogenous promoter was influenced in vivo when the expression of UXT was diminished. Consistently, deficiency of the endogenous UXT level resulted in considerable decreases in the amount of p65 associated with its endogenous cognate promoters upon TNF-α stimulation.

Given that UXT interacted with p65 and was essential to maintain the presence of NF-κB inside the nucleus, we wondered whether UXT was an integral component of the NF-κB transcriptional enhanceosome in vivo. To address this possibility, we transfected HA-UXT into 293T cells and performed systematic ChIP assays on the promoters of A20 and IκBα as described in Materials and methods. It turned out that UXT was indeed present within the NF-κB transcriptional enhanceosome. Notably, its presence became much more prominent upon stimulation, which suggested that UXT was dynamically recruited onto the enhanceosome. In addition, UXT had nothing to do with the transcription complex on the GAPDH promoter, indicating the selectivity of UXT action (Fig. 7 A). We also stimulated 293T cells and performed similar ChIP assays to confirm again that endogenous UXT was recruited onto the NF-κB enhanceosome in response to stimulation (Fig. 7 B).

After 10 ng/ml TNF-α stimulation, ChIP assays were performed on A20, IκBα, or GAPDH promoters as described.

Next, we studied in vivo histone binding by protein chromatin immunoprecipitation (ChIP) assays and observed that AIRE is found in complexes with a small fraction of histone H3 but not with H3K4me3. By contrast, binding of ING2, used as a positive control, was detected for both H3 and H3K4me3 (supplementary Fig S7 online). By using DNA ChIP analysis, we found that AIRE interacts with the insulin, involucrin and S100A8 promoter regions, but much less with the S100A10 and GAPDH promoters (Fig 5B).

(B) DNA ChIP analysis with anti-AIRE and IgG was performed from the stably transfected cells, as indicated. The fold differences are normalized to input fractions and shown in comparison with the background level (=1) with each primer set in each condition. The data are the averages of two independent experiments. AIRE, autoimmune regulator; ChIP, chromatin immunoprecipitation; HEK, human embryonic kidney; PHD, plant homeodomain.

Chromatin immunoprecipitation indicated that DDX3X is recruited to the IFN promoter upon infection with Listeria monocytogenes, suggesting a transcriptional mechanism of action. DDX3X was found to be a TBK1 substrate in vitro and in vivo. Phosphorylation-deficient mutants of DDX3X failed to synergize with TBK1 in their ability to stimulate the IFN promoter. Overall, our data imply that DDX3X is a critical effector of TBK1 that is necessary for type I IFN induction.

To address this question, we used chromatin immunoprecipitation (ChIP) and amplified the enhanceosome-binding region of the IFN-β promoter by quantitative PCR (Figure 5A). IRF3, absent under non-stimulated conditions, was recruited to the IFN promoter upon infection with L. monocytogenes (Figure 5B). The specificity of this signal was ascertained using a control serum for ChIP.

ChIP suggests that DDX3X can be recruited to the IFN promoter, positioning DDX3X downstream of TBK1 at the level of IRF3.

Cells were cultured for 24h and prepared using a ChIP assay kit from Upstate Biotechnology, Inc. (Lake Placid, NY) according to the manufacturer's recommendations and preformed as previously describe 28. Primers for the FOXO3a and SCO2 promoter are shown (Supplemental Methods) and their location is shown in supplemental Fig. S3. IP and transient transfection IP westerns were done as previously described 28. Bands for all IPs were detected using an ECL protocol (Santa Cruz Biotechnology, Santa Cruz, CA) and visualized with Fuji Las-3000 intelligent darkbox (FujiFilm Medical Systems, Stamford, CT).

ChIP analysis showed that wt-SIRT3 cells have an increase in FOXO3a binding to upstream regulatory regions of gene promoters that each contain two canonical FOXO3a binding sites roughly 1 kb upstream of the transcription start site (supplemental Fig. S3). FOXO3a binding to both the MnSOD (Fig.3D, upper panel) and SCO2 (lower panel) promoters is increased in the wt-SIRT3, as compared to mt-SIRT3 cells. These experiments imply that SIRT3 may increase FOXO3a DNA-binding.

(D) ChIP analysis of FOXO3a finding to the MnSOD and SCO2 promoters in HCT116 cells that overexpress either a wild-type or deacetylation null SIRT3 gene.

Chromatin immunoprecipitation (ChIP) experiments show that Ctk1 associates with RNApII throughout elongation (Kim et al, 2004a).

To determine whether Ctk1 regulates the disposition of basal transcription factors, ChIP experiments on three constitutively expressed genes (PMA1, ADH1, and PYK1) were performed as described previously (Kim et al, 2004a). ChIP analysis using polyclonal antibodies against TBP (TATA-binding protein), TFIIB (Sua7), TFIIE (Tfa2) and TFIIH (Tfb1 and Kin28) generally confirmed the localization of these basal transcription factors at promoter regions in a wild-type (WT) background.

Consistent with this, when ChIP was carried out using strains deleted for the other subunits of the Ctk1 complex, including Ctk2 (cyclin subunit) or Ctk3 (accessory factor), TBP occupancy was again increased in the coding region of PMA1 (Supplementary Figure S1).

To exclude the possibility that the abnormal cross-linking of basal transcription factors in ctk1Δ cells was due to issues with the specific antibodies, the experiment was repeated with strains carrying TAP-tagged Tfg1, Tfg2, or Tfg3 (all subunits of TFIIF). ChIP was carried out with IgG-agarose as described previously (Kim et al, 2004a). Again, spreading of TFIIF from the promoter throughout the transcribed region was seen in ctk1Δ cells (Figure 2, Tfg1–TAP result shown is representative). Some promoter preference was still seen, but the qualitative difference upon Ctk1 deletion was clear.

A second model that could explain the cross-linking of basal factors throughout a gene is that they fail to release from RNApII as it moves into productive elongation. To test whether the extended pattern of basal factor cross-linking was completely coincident with RNApII, ChIP experiments were carried out at the termination site for PMA1 (Figure 3). RNApII levels decreased between primer sets 7 and 9 in WT and ctk1Δ strains, indicative of transcription termination in both backgrounds (Ahn et al, 2004; Kim et al, 2004b). TBP and TFIIH (Kin28) decrease in parallel with RNApII in ctk1Δ cells (Figure 3A), supporting the idea that these basal transcription factors travel with elongating RNApII.

ChIP analysis for TBP was performed in these strains to see whether aberrant cross-linking was observed. As serine 2 phosphorylation is known to be required for cotranscriptional 3′-end processing (Ahn et al, 2004), we also tested for the polyadenylation factor Rna14 as a positive control (Figure 4B). In accordance with our previous report, Rna14 cross-linking at the 3′-end of PMA1 was reduced to about one-third of WT levels.

As in vitro experiments with mammalian Cdk9 (Wada et al, 1998; Chao and Price, 2001) and ChIP experiments with Ctk1 (Cho et al, 2001) also suggest that these kinases function after initiation, we modified the protocol.

There are at least two possible variations of this model that could explain the ChIP results (Figure 5G). In the first, basal factors release from the promoter in ctk1Δ cells, but remain associated with the polymerase as it moves through the gene body. In the second, RNApII actually remains tethered to the basal factors at the promoter while transcribing. This would result in the transcribing polymerase ‘pulling' the transcribed region past basal factors, thereby allowing cross-linking with downstream sequences.

Chromatin immunoprecipitation was performed as described previously (Ahn et al, 2004). For precipitation of TAP-tagged proteins, 10 μl of rabbit IgG agarose (Sigma) was incubated with chromatin solution overnight at 4°C. For Rpb1 (BWG16 Covance), Sua7, TBP, Kin28, Tfb1, Tfa2, and Rap1 (y-300, Santa Cruz) immunoprecipitations, the antibodies were preincubated with protein A-sepharose CL-4B (Amersham) for 1 h at room temperature and then incubated with chromatin solution overnight at 4°C. For Rpb3 immunoprecipitation, antibody (1Y26, NeoClone) was preincubated with protein G-sepharose (Amersham). PCR amplification was performed with primer pairs as described (Ahn et al, 2004). PCR signals were quantitated by Fujix PhosphoImager. To control for amplification efficiency and label incorporation with different primers, the ratio of each gene-specific product to that of a non-transcribed region of chromosome V was calculated from the input sample signals. The signal for each specific gene primer in the immunoprecipitation was then divided by this ratio to convert the signal to normalized units. This value was divided by the immunoprecipitation signal of the non-transcribed control product to determine the fold enrichment of the ChIP over background signals. As this number is a ratio, a value of 1 represents ‘no signal' (i.e., a signal equal to background).

Ctk1 is necessary for the dissociation of basal transcription factors from elongating RNA polymerase II (RNApII). (A) Schematic of the PMA1, ADH1, and PYK1 genes. The UAS of each gene is indicated by an open box (see Figure 3C). The TATA/promoter region and open reading frames are represented by black and grey boxes, respectively. Arrows indicate the position of the major polyadenylation sites reported previously (Kim et al, 2004a) and bars below the genes show the relative positions of PCR products in ChIP analysis. (B) Occupancy of Rpb1 and basal transcription factors (Sua7, TBP, Kin28, Tfb1, Tfa2) at the indicated regions in WT (YSB726) or ctk1Δ (YSB854) cells. INPUT was used to normalize the PCR amplification and the asterisk marks a non-transcribed PCR fragment indicated in all reactions as a background control. (C) Quantitation of the ChIP experiments in (B), with PMA1 as representative. The x axis indicates the specific primer pair used in each PCR. The y axis shows the specific signal relative to the negative control (i.e., a ratio of one is equivalent to background).

(A) ChIP analysis was carried out using Rpb3–TAP (YSB956) or Tfg1–TAP tagged strains (YSB925) +/− Ctk1. Similar cross-linking patterns were seen with Tfg2–TAP (YSB926) or Tfg3–TAP (YSB927) strains (not shown).

Basal transcription factors coincide with elongating RNApII in ctk1Δ cells. (A) ChIP analyses were carried out with antibodies against Rpb3, TBP, or Kin28 in WT (YSB726) and ctk1Δ (YSB854) backgrounds. Numbers (6–9) correspond to PMA1 primer locations in Figure 1A. PCR products from (A) and (C) are shown in Figure 1B. (B) Quantitation of results in (A). (C) Occupancies of Rap1, Rpb1, and TBP at the indicated regions of genes were determined using the indicated polyclonal antibodies in both WT (YSB726) and ctk1Δ (YSB854) backgrounds. The upstream activating sequence (UAS) regions of each gene are depicted in Figure 1A.


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The amount of biotinylated (intracellular) β1-integrin was determined with ELISA.

HEK293 cells were seeded at 5 × 105 cells per well in six-well plates. Cells were transfected with 500 ng of pDM128 (Rev reporter plasmid), 30 ng of Rev expression plasmid, 300 ng of DDX3X expression plasmid and 500 ng of TBK1 expression plasmid. All samples were normalized for DNA content using an empty plasmid. Cells were harvested 24 h post-transfection and CAT levels were analysed by ELISA (Roche) according to the manufacturer's instructions.

(B) HEK293 cells were transfected with the reporter plasmid pDM128 and constant amounts of Rev DDX3X wt or K230A and TBK1 wt or K38M were added as indicated. CAT expression was measured 24 h post-transfection by ELISA (Roche).


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(C) 293T cells were induced by 10 ng/ml TNF-α for 30 min, and nuclear extracts were prepared and incubated with the indicated antibodies to perform EMSA supershift assays.

Alternatively, we performed supershift assay to further substantiate this observation. Stimulation of 293T cells with TNF-α led to a strong NF-κB DNA-binding band composed of κB probe, NF-κB, and its cofactors in EMSA. Interestingly, this EMSA band was markedly diminished when antibody against UXT was introduced into the reaction mixture (Fig.


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Because UXT interacted specifically inside the nucleus with p65 in a signal-dependent manner, we went on to address whether this interaction was important for regulating NF-κB activity and genes responsive to it. We transfected 293T cells with UXT or other control plasmids, prepared nuclear extracts, and performed electrophoretic mobility shift assay (EMSA) as indicated. Consistently, there was no NF-κB binding to its cognate probe without TNF-α treatment. Notably, the overexpression of UXT alone did not induce any detectable basal NF-κB binding activity.

Overexpression of UXT does not markedly affect NF-κB activation induced by TNF-α. (A) 293T cells were transfected with an equal amount of UXT, A20, or control vector. 24 h after transfection, cells were stimulated with 10 ng/ml TNF-α for the indicated times. Equal amounts (10 μg) of nuclear extracts were subjected to EMSA. For competition analysis, 100-fold excess of unlabeled wild-type or mutant κB probes were added to the reaction mixtures. For supershift assays, nuclear extracts were incubated with antibody as indicated.

EMSA was performed to test endogenous NF-κB or sp1 binding to their cognate probes.

Via EMSA, we then analyzed endogenous NF-κB DNA binding activity in cells with reduced UXT expression. Expectedly, TNF-α alone induced endogenous NF-κB to bind to its cognate probe strongly and specifically. Considerably, this interaction was markedly diminished in nuclear extracts from UXT-specific knockdown cells. In addition, this reduction was correlated with the effectiveness of the siRNA administered (Fig. 5 B).

EMSA was performed to test endogenous NF-κB or sp1 binding to their cognate probes.

Furthermore, EMSA confirmed that the loss of UXT also suppressed the constitutive DNA binding activity of NF-κB but not the control sp1 (Fig. 9 E). These results indicate that the elevated expression of UXT strongly correlates with constitutive NF-κB activity in prostate cancer cell lines and again substantiates the notion that UXT is essential for NF-κB function in the nucleus.


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The effect of depleting RCK/p54 on localization of Ago2 was next examined by immunofluorescence analysis of HeLa cells expressing Myc-Ago2 and siRNAs against RCK/p54. As shown in Figure 4B, 4C, 4F, and 4G, depleting RCK/p54 disrupted the cellular P-body structures. In these P-body-deficient cells, Ago2 proteins were diffused throughout the cytoplasm and no longer accumulated at specific foci (Figure 4A and 4E). This cytoplasmic redistribution of Ago2 suggests that its localization to P-bodies is driven in mammalian cells by factors such as RCK/p54.

We next visualized Lsm1 and Ago2 by immunofluorescence using antibodies against Lsm1 and Myc tag and found that Lsm1 and Ago2 co-localized in P-bodies (Figure 5B). To determine the status of P-body structures and RISC localization in cells after Lsm1 knockdown, Lsm1-depleted cells were analyzed by immunofluorescence. We found that P-body structures were drastically disrupted, Ago2 was diffused throughout the cytoplasm, and Lsm1 and Ago2 were minimally co-localized (Figure 5B). These results show that P-body structures were drastically disrupted and Ago2 was diffuse throughout the cytoplasm when Lsm1 was depleted by RNAi.

(B) Subcellular localization of endogenous and exogenous UXT. 293T cells were transfected with (top) or without (bottom) FLAG-UXT. Immunofluorescentmicroscopy was performed with the indicated primary antibodies.

Alternatively, we performed immunofluorescence analysis to support this speculation. Interestingly, when transfecting cells with siRNA against UXT and stimulating them with TNF-α, there were an apparently decreased percentage of cells that displayed focused nuclear p65. Approximately 90% of control cells displayed p65 inside the nucleus upon stimulation, whereas only 35–50% displayed p65 in the case of the UXT knockdown specimen (Fig. 5 E).

Consistent with these data, S329/T331-phosphorylated MDC1 was detected by immunofluorescence in both control and γ-irradiated cells (Fig 2B; note the absence of staining in cells treated with MDC1 small interfering RNA (siRNA)), confirming that SDTD-phosphorylated MDC1 is present in the absence of damage and also forms IRIF.

(B) Fluorescence microscopy demonstrating a shift in the subcellular localization of GFP-Sed5 from Golgi in control (WT) strains to a partially cytosolic localization in a Δget3 strain, and both cytosolic and few large puncta in Δget1/2 strains. The GFP-Sed5 puncta in get mutants do not colocalize with the Golgi marker Anp1-RFP.

(C) Fluorescence microscopy demonstrating colocalization of GFP-Sed5 and Get3-tdRFP in cytosolic aggregates that form in a Δget1/2 background.

(B) Fluorescence microscopy of control (WT) and Δget1/2 strains expressing a broad variety of TA proteins. GFP-Scs2, GFP-Sbh1, and GFP-Ysy6 under a galactose-inducible (GAL) promoter. Cherry-Sbh2 was expressed from a plasmid under the constitutive TEF2 promoter.

(C) Fluorescence microscopy of control (WT) and Δget1/2 strains expressing two mitochondrial TA proteins, Cherry-Fis1 and Cherry-Tom22, expressed from a plasmid under the constitutive TEF2 promoter.

(A) Fluorescence microscopy showing the localization of GFP-Ubc6 and mitochondrially targeted dsRED (MTS-RFP) in a control (WT) or Δget1/2 strain.

(B) Fluorescence microscopy of a time course monitoring the subcellular localizations of the peroxisomal TA protein GFP-Pex15 as well as dsRED targeted to the mitochondria (MTS-RFP) following induction of Pex15 from a galactose inducible promoter in a control (WT), get1/2, or get3 strain.

To ascertain the time in infection and the subcellular compartment at which the interaction occurs, ASFV infected cells were examined by confocal laser scanning microscopy at different times post infection (at least 30 infected cells were analyzed for each time point at 0, 6, 8 and 12 hpi).

Subcellular distribution of p30 and hnRNP-K were analyzed by immunofluorescence microscopy in ASFV or mock infected cells. ASFV infection resulted in an intensification of nuclear, but not cytoplasmic, staining of hnRNP-K when compared to uninfected cells. When nuclei of infected cells were examined in detail, nuclear hnRNP-K was detected in characteristic granular structures coincident with areas completely devoid of nucleic acid staining. Interestingly, these kind of hnRNP-K accumulation was also observed in cells transiently expressing p30 (pCMV-p30), but not in mock-infected cells, suggesting the involvement of p30 on these changes (Fig. 4) during ASFV infection.

Colocalization of p30 and hnRNP-K in the nucleus of infected cells. Cells were infected with BA71V strain and analyzed at different times post infection by confocal immunofluorescence microscopy acquiring 0.1 μm optical sections from the Z-axis. A representative image of an infected cell at 8 hpi is shown (A). p30 was detected with a monoclonal anti-p30 followed by Alexa 647-conjugated goat anti-mouse antibody (red) and hnRNP-K with an anti-hnRNP-K specific serum followed by Alexa 488-conjugated goat anti-rabbit antibody (green). Discrete spots of colocalization of both proteins (orange) can be discerned in the cell nucleus.

ASFV infection induces changes in subcellular distribution of hnRNP-K. (A) Vero cells were either infected with BA71V (1 pfu/cell) or transfected with pCMV-p30 and analyzed by immunofluorescence microscopy at 8 hpi or 24 h post transfection, respectively. ASFV protein p30 was detected with a mouse anti-p30 monoclonal antibody and rhodamine red conjugated corresponding secondary antibody (red). hnRNP-K distribution in the cell nucleus was detected with a rabbit antiserum anti-hnRNP-K and Alexa Fluor 488 conjugated corresponding secondary antibody (green). Nuclei of cells were stained with Hoechst 3332. Accumulation of hnRNP-K in infected cells as spots which are coincident with the absence of nucleic acids staining can be appreciated in BA71V infected and pCMV-p30 transfected cells.


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Rho GTPases, including RhoA, Rac1, and Cdc42, are key regulators of actin cytoskeleton dynamics. Since BDNF stimulation has been observed to activate Rac1 and Cdc42 in neurons [15], we were interested to delineate if Rho GTPases contribute to BDNF-stimulated dendritic growth. To investigate if Rho GTPases are involved, and to identify the Rho GTPase(s) implicated, hippocampal neurons were transfected with WT or DN Rac1, Cdc42, or RhoA. We found that while overexpression of WT and DN Rac1 increased the basal number of dendrites in the absence of BDNF treatment, overexpression of both forms of Rac1 abolished BDNF-stimulated dendritic growth. On the other hand, while overexpression of DN RhoA slightly enhanced primary dendrites irrespective of BDNF stimulation, overexpression of both WT and DN forms of RhoA inhibited BDNF-stimulated dendritic growth. Remarkably, in contrast to the inhibition of BDNF-stimulated dendritic growth in cells overexpressing WT Rac1 and RhoA, BDNF stimulation of hippocampal neurons overexpressing WT Cdc42 resulted in an increase in primary dendrites, which was nearly abolished by overexpression of DN Cdc42 (Figure 6A). Our observations therefore suggest that while Rac1 and RhoA may also modulate BDNF-stimulated dendritic growth, it is the activation of Cdc42 following BDNF stimulation that most likely mediates the increase in primary dendrites by BDNF.

In the current study, we demonstrated that Ser478 phosphorylation of TrkB by Cdk5 is essential for the Cdc42-dependent increase in primary dendrites triggered by BDNF, thus adding a new regulatory component to the mechanisms involved in Rho GTPase activation by neurotrophin. Although the precise downstream pathways by which this phosphorylation affects Cdc42 activation remains to be determined, our observations provide some interesting insights.

Interestingly, APPL1 would bind to Rab21 in a GTP-dependent manner (Figure 4), indicating that APPL1 is an effector for both Rab5 and Rab21.

GTPase binding has emerged as a major function of PH domains in addition to lipid binding (Lemmon, 2004). For example, PH domains in some guanine nucleotide-exchange factors (GEF) have been shown to bind directly to their cognate small GTPases (Rossman et al, 2002, 2003; Lu et al, 2004), and our data now show direct interaction between the APPL1 PH domain and Rab5. So far, only two crystal structures of small GTPase−PH domain complexes are available. One is Ran−RanBD1 (PDB file 1RRP). The interactions between the Ran GTPase domain and RanBD1 PH core domain is fairly minor, occurring between the switch I region of the GTPase (equivalent to the 40's in Rab5) and strand β2 of the PH domain. This interaction alone is unlikely to be sufficient to form a stable complex. Indeed, Ran has a long C-terminal peptide beside the GTPase domain, while the PH domain of RanBD1 has an extra N-terminal peptide. These two terminal peptides wrap around the partner proteins forming the major interaction between Ran and RanBD1. Such an interaction seems not to be required for Rab5 and APPL1, because the GTPase domain of Rab5 and BAR-PH domain of APPL1 are sufficient to mediate their interaction. The second published small GTPase−PH complex is that of Ral−Exo84 (PDB file 1ZC3). In this complex, the PH domain of Exo84 uses L1, β5, and L6 to interact with the interswitch and switch II regions of Ral forming an intermolecular β-sheet extension mediated by the PH β5 strand and GTPase β2 strand (Jin et al, 2005). Our mutagenesis analysis points to a different surface region (β3, L3, and β4) of the PH domain for Rab5 binding. Therefore, the Rab5−APPL1 interaction represents a new GTPase−PH binding mode.


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We also found that MELK physically interacted with Bcl-GL through its amino-terminal region. Immunocomplex kinase assay showed that Bcl-GL was specifically phosphorylated by MELK in vitro.


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More importantly, when siRNA of UXT, which blocked NF-κB activity, was used in combination with TNF and CHX, we detected drastically morphological changes under light microscopy and ∼50% of cells undergoing apoptosis. Alternatively, we used annexin V and TUNEL methods to quantitatively measure the percentages of cells undergoing apoptosis. Consistently, a considerable increase of apoptotic cells was observed in samples with reduced UXT (Fig. 8, B and C). These indicated that the knockdown of UXT sensitizes 293T cells to apoptosis induced by TNF-α.


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The cytoplasmic retention of the chimeric GFP-STM protein was further substantiated by confocal laser microscopy scanning and staining of nuclei with propidium iodide. The comparison of a median Z-stack projection through the nucleus either analysed for GFP-STM fluorescence (Figure 1F), the corresponding propidium iodine stains (Figure 1G) and their overlay in Figure 1H confirm exclusion of GFP fluorescence from the nuclear compartment.

We further extended these binding studies and confirmed the above Rab5-binding site in vivo in the cell, by monitoring Rab5-mediated APPL1 recruitment to early endosomes in the cell via confocal microscopy. In this case, the RFP (DsRed-monomer)−Rab5-Q79L fusion protein was expressed in PC12 cells, targeted to the early endosomes, and recruited effectively the coexpressed GFP (green fluorescence protein)−APPL1 to these early endosomes (Figure 5C). Importantly, APPL1 (5−385), that is, the BAR-PH domain, was sufficient to target to Rab5-Q79L containing early endosomes (Figure 5C). In contrast, one of the Rab5-binding defective mutants (A318D) failed to target the early endosomes and exhibited a diffused pattern throughout the cytoplasm in the cell (Figure 5C).

(C) Mutational effects on APPL1 targeting to Rab5-positive early endosomes in the cell. RFP−Rab5-Q79L was coexpressed with GFP−APPL1 (full length, FL; BAR-PH domain; or BAR-PH mutant) in PC12 cells as indicated, followed by confocal fluorescence microscopy. Shown are typical confocal microscopic images indicating the RFP−Rab5-Q79L labeled early endosomes (red) and the colocalization of GFP−APPL1 or mutants (green) in the same cells. Scale bar, 16 μm.

Tobacco leaf epidermal cells from agroinfiltration were examined under a Zeiss LSM 510 inverted confocal laser-scanning microscope (Zeiss, Jena, Germany) following the settings described by Goodin et al. (2002) with minor modifications. Single optical sections were scanned as resulting images for each transient expression. For each plasmid construct, 10–15 cells were imaged with similar results. GFP fluorescence was excited at 488 nm, filtered through a primary dichroic (UV/488/543), a secondary dichroic of 545 nm, and subsequently through BP505–530 nm emission filters to the photomultiplier tube (PMT) detector. DsRed fluorescence was excited at 543 nm, the emission was passed through similar primary and secondary dichroic mirrors and finally through a BP560–615 nm emission filter to the PMT detector. Fluorescence resonance energy transfer (FRET) pairs GFP/DsRed were analysed using a confocal laser-scanning microscope (Zeiss LSM510 META). FRET measurements of DsRed emission with zero contribution from GFP, was accomplished as described by Erickson et al. (2003) using the following settings: excitation at 488 nm and emission filters, BP 505–530 nm for GFP and BP 600–637 nm for DsRed.

Following agroinfiltration of tobacco leaves with both ACBP4:DsRed and GFP:AtEBP, observations were carried out by confocal microscopy using a green filter to investigate the fluorescence pattern of GFP and a red filter to visualize the fluorescence of DsRed (Fig. 2). GFP:AtEBP was located mainly in the nucleus, with some signals at the cytosol and the plasma membrane (Fig. 2A). ACBP4:DsRed was localized predominantly to the cytosol, inclusive of signals detected in the cytosol surrounding the nucleus (Fig. 2B). Signals of both fusion proteins were common to the cytosol.

Confocal images indicating co-localization of ACBP4:DsRed and GFP:AtEBP fusion proteins transiently-expressed in tobacco leaves. Representative tobacco leaf epidermal cells are shown by laser-scanning confocal microscopy following agroinfiltration of plasmid pAT282 or pAT225.


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To identify novel interactors of TBK1, we generated stable RAW264.7 cell lines that express a GS-TAP-tagged version of TBK1 and purified the TBK1 protein complex by tandem affinity purification (Figure 1A and B) (Burckstummer et al, 2006).

GS-TAP-tagged DDX3X, isolated by affinity purification using rabbit IgG agarose, was also recruited to the enhanceosome region upon L. monocytogenes infection (Figure 5C). This recruitment was specific because a control region (Figure 5A) could not be amplified by PCR under these conditions. This suggests that DDX3X exerts a direct effect on the IFN-β promoter.

Tandem affinity purification using the GS-TAP cassette was performed as described previously (Burckstummer et al, 2006).

The DEAD-box helicase DDX3X is a target of TBK1. (A) Schematic representation of the tandem affinity purification protocol: GS-TAP-tagged TBK1 expressed in RAW264.7 cells was purified using rabbit immunoglobulin G (IgG) agarose and eluted by tobacco etch virus (TEV) protease cleavage. Next, the remaining complex was purified using streptavidin agarose and eluted by boiling in SDS sample buffer. (B) The final TAP eluate was separated on an SDS–PAGE and stained by silver staining. Protein complex composition was analysed by LC-MSMS. The position of several complex components is depicted next to the region where it was identified.


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According to bimolecular fluorescence complementation, STM is targeted into the nuclear compartment through heterodimerization with BLH partner proteins, which are expressed in distinct SAM domains from the center to the periphery.

The method of choice to prove a physical interaction between candidate partner proteins in plant cells is BiFC (31,36).

As a potential transcription factor, STM was expected to direct GFP fluorescence into the plant cell nucleus. However, both fusion proteins, GFP-STM and STM-GFP remained in the cytoplasm (Figure 1B and C).

The principle of BiFC is the formation of a fluorescent complex by fragments of the enhanced YFP brought together by the association of two interaction partners (31,36). To verify STM interaction partners we constructed translational fusions between the N- or the C-terminal YFP sub-domains and the full-length STM protein or individual full-length BLH partner proteins. Optimal combinations of chimeric gene constructs are indicated in Figure 4 above each corresponding experiment. Chimeric genes were co-delivered into leek epidermal cells by bombardment of tungsten particles double-loaded with two appropriate chimeric gene constructs. Any detectable fluorescence was dependent on the combination of the C-terminal with the N-terminal YFP sub-domain fused to STM or BLH partner proteins. In contrast, the fusion of the C- or the N-terminal YFP sub-domain to the STM protein and their co-bombardment in plant cells gave no fluorescence (data not shown). BiFC therefore provided no evidence for the formation of STM homodimers but substantiated the formation of STM/BLH heterodimers.

In contrast to results obtained with STM-GFP fusions, however, the BiFC signal exhibited a high preference for the nucleus whereas fluorescence was low in the cytoplasm. The BiFC results therefore, confirm that full-length STM and BLH proteins interact in plant cells, but more notably, that BLH/STM heterodimers are efficiently incorporated into the nuclear compartment.

Reminiscent, however, of the MEIS/PBC situation in animals, the GFP and BiFC data show that the TALE-HD transcription factor STM is targeted into the nuclear compartment through interaction with three BLH partner proteins, which are members of the non-MEINOX class of TALE-HD proteins in plants.

The BELL domain is essential for the interaction of STM with BLH partner proteins. BiFC experiments with BLH3 deletion polypeptides (BLH3 ΔC or BELLBLH3 see schematic drawing in Figure 4G) demonstrated that amino acid sequences upstream and downstream of the BELL domain including the homeodomain are dispensable for heterodimerization with STM (Figure 4H and I). BiFC fluorescence still exerted a preference for the nucleus if sequences C-terminal to the BELL domain including the DNA-binding homeodomain were deleted (BLH3 ΔC, Figure 4H). In contrast, YFP fluorescence reconstituted with the isolated BELL domain (BELLBLH3 in Figure 4I) remained mainly cytoplasmic. An effective NLS therefore must be located in the BLH3 protein N-terminal to its BELL domain. In the STM protein, the MEINOXSTM domain is sufficient to interact with full-length BLH3 (Figure 4J). However, we have been unable to reconstitute YFP fluorescence in BiFC experiments with the isolated MEINOXSTM and BELLBLH3 domains although several compatible N- or C-terminal combinations with each YFP sub-domain were tested. One explanation for this may be that the relative arrangement of YFP sub-domains sterically interferes with their functional association.

Co-immunoprecipitation and BiFC experiments performed in the course of this study show that the MEINOXSTM and the BELLBLH3 domains are sufficient for interaction in vitro and in planta.


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(A) Immunodepletion of Tipin. Extracts were mock depleted (Mock) or Tipin depleted using anti-Tipin antibodies (ΔTipin).


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For example, p300/CBP played a major role in the acetylation of p65 in vivo (Chen et al., 2002). Conversely, acetylated p65 was subjected to deacetylation by HDAC3 (Chen et al., 2001). Recent work also revealed that SIRT1 physically interacted with p65 and promoted p65 deacetylation (Yeung et al., 2004).


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