Schwenk J.,Albert Ludwigs University of Freiburg |
Schwenk J.,Center for Biological Signaling Studies |
Harmel N.,Sensory Medical |
Brechet A.,Albert Ludwigs University of Freiburg |
And 14 more authors.
Neuron | Year: 2012
AMPA-type glutamate receptors (AMPARs) are responsible for a variety of processes in the mammalian brain including fast excitatory neurotransmission, postsynaptic plasticity, or synapse development. Here, with comprehensive and quantitative proteomic analyses, we demonstrate that native AMPARs are macromolecular complexes with a large molecular diversity. This diversity results from coassembly of the known AMPAR subunits, pore-forming GluA and three types of auxiliary proteins, with 21 additional constituents, mostly secreted proteins or transmembrane proteins of different classes. Their integration at distinct abundance and stability establishes the heteromultimeric architecture of native AMPAR complexes: a defined core with a variable periphery resulting in an apparent molecular mass between 0.6 and 1 MDa. The additional constituents change the gating properties of AMPARs and provide links to the protein dynamics fundamental for the complex role of AMPARs in formation and operation of glutamatergic synapses.
Duncker S.V.,University of Tübingen |
Franz C.,University of Tübingen |
Kuhn S.,University of Sheffield |
Kuhn S.,University of Tübingen |
And 14 more authors.
Journal of Neuroscience | Year: 2013
The encoding of auditory information with indefatigable precision requires efficient resupply of vesicles at inner hair cell (IHC) ribbon synapses. Otoferlin, a transmembrane protein responsible for deafness in DFNB9 families, has been postulated to act as a calcium sensor for exocytosis as well as to be involved in rapid vesicle replenishment of IHCs. However, the molecular basis of vesicle recycling in IHCs is largely unknown. In the present study, we used high-resolution liquid chromatography coupled with mass spectrometry to copurify otoferlin interaction partners in the mammalian cochlea. We identified multiple subunits of the adaptor protein complex AP-2 (CLAP), an essential component of clathrin-mediated endocytosis, as binding partners of otoferlin in rats and mice. The interaction between otoferlin and AP-2 was confirmed by coimmunoprecipitation. We also found that AP-2 interacts with myosin VI, another otoferlin binding partner important for clathrin-mediated endocytosis (CME). The expression of AP-2 in IHCs was verified by reverse transcription PCR. Confocal microscopy experiments revealed that the expression of AP-2 and its colocalization with otoferlin is confined to mature IHCs. When CME was inhibited by blocking dynamin action, real-time changes in membrane capacitance showed impaired synaptic vesicle replenishment in mature but not immature IHCs. We suggest that an otoferlin-AP-2 interaction drives Ca2+- and stimulus-dependent compensating CME in mature IHCs. © 2013 the authors.
Jeworutzki E.,CNR Institute of Biophysics |
Lopez-Hernandez T.,University of Barcelona |
Capdevila-Nortes X.,University of Barcelona |
Sirisi S.,University of Barcelona |
And 17 more authors.
Neuron | Year: 2012
Ion fluxes mediated by glial cells are required for several physiological processes such as fluid homeostasis or the maintenance of low extracellular potassium during high neuronal activity. In mice, the disruption of the Cl - channel ClC-2 causes fluid accumulation leading to myelin vacuolation. A similar vacuolation phenotype is detected in humans affected with megalencephalic leukoencephalopathy with subcortical cysts (MLC), a leukodystrophy which is caused by mutations in MLC1 or GLIALCAM. We here identify GlialCAM as a ClC-2 binding partner. GlialCAM and ClC-2 colocalize in Bergmann glia, in astrocyte-astrocyte junctions at astrocytic endfeet around blood vessels, and in myelinated fiber tracts. GlialCAM targets ClC-2 to cell junctions, increases ClC-2 mediated currents, and changes its functional properties. Disease-causing GLIALCAM mutations abolish the targeting of the channel to cell junctions. This work describes the first auxiliary subunit of ClC-2 and suggests that ClC-2 may play a role in the pathology of MLC disease. Video Abstract: Leukodystrophies are a group of genetic diseases affecting white matter. Jeworutzki et al. find that GlialCAM, a cell-adhesion molecule which is mutated in a leukodystrophy, serves as an auxiliary subunit of the chloride channel ClC-2. © 2012 Elsevier Inc.
Constantin C.E.,Albert Ludwigs University of Freiburg |
Muller C.S.,Albert Ludwigs University of Freiburg |
Leitner M.G.,University of Marburg |
Bildl W.,Albert Ludwigs University of Freiburg |
And 5 more authors.
Proceedings of the National Academy of Sciences of the United States of America | Year: 2017
Voltage-activated calcium (Cav) channels couple intracellular signaling pathways to membrane potential by providing Ca2+ ions as second messengers at sufficiently high concentrations to modulate effector proteins located in the intimate vicinity of those channels. Here we show that protein kinase Cβ (PKCβ) and brain nitric oxide synthase (NOS1), both identified by proteomic analysis as constituents of the protein nano-environment of Cav2 channels in the brain, directly coassemble with Cav2.2 channels upon heterologous coexpression. Within Cav2.2-PKCβ and Cav2.2-NOS1 complexes voltage-triggered Ca2+ influx through the Cav channels reliably initiates enzymatic activity within milliseconds. Using BKCa channels as target sensors for nitric oxide and protein phosphorylation together with high concentrations of Ca2+ buffers showed that the complex-mediated Ca2+ signaling occurs in local signaling domains at the plasma membrane. Our results establish Cav2-enzyme complexes as molecular entities for fast electrochemical coupling that reliably convert brief membrane depolarization into precisely timed intracellular signaling events in the mammalian brain.
News Article | November 30, 2016
No statistical methods were used to predetermine sample size. The experiments were not randomized and the investigators were not blinded to allocation during experiments and outcome assessment. Yeast strains are all based on the BY4741 laboratory strain28. Manipulations were performed using a standard PEG/LiAC transformation protocol29. All deletions were verified using primers from within the endogenous open reading frame. Primers for all genetic manipulations were planned either manually or using the Primers-4-Yeast web tool30. All strains, primers and plasmids used in this study28, 31, 32, 33, 34 are listed in Supplementary Table 4. SGA and microscopic screening were performed using an automated microscopy set-up as previously described11, 15, using the RoToR bench-top colony arrayer (Singer Instruments) and automated inverted fluorescent microscopic ScanR system (Olympus). Images were acquired using a 60× air lens with excitation at 490/20 nm and emission at 535/50 nm (GFP) or excitation at 575/35 nm and emission at 632/60 nm (RFP). After acquisition, images were manually reviewed using the ScanR analysis program. Manual microscopy was performed using by one of two apparatuses. (I) Olympus IX71 microscope controlled by the Delta Vision SoftWoRx 3.5.1 software. Images were acquired using a 60× oil lens and captured by PhoetometricsCoolsnap HQ camera with excitation at 490/20 nm and emission at 528/38 nm (GFP/YFP) or excitation at 555/28 nm and emission at 617/73 nm (mCherry/RFP). (II) VisiScope Confocal Cell Explorer system, composed of a Zeiss Yokogawa spinning disk scanning unit (CSU-W1) coupled with an inverted Olympus IX83 microscope. Images were acquired using a 60× oil lens and captured by a connected PCO-Edge sCMOS camera, controlled by VisView software, with wavelength of 488 nm (GFP) or 561 nm (mCherry/RFP). Images were transferred to Adobe Photoshop CS6 for slight adjustments to contrast and brightness. Lysates for immunoprecipitation were prepared from indicated strains in mid-logarithmic growth grown in YPD reach medium. Cells were harvested, washed in distilled water, and resuspended in lysis buffer (50 mM Tris HCl pH 7, 150 mM NaCl) supplemented with protease inhibitors (complete EDTA-free cocktail; Roche) and frozen in a drop-by-drop fashion in liquid nitrogen. Frozen cells were then pulverized in a ball mill (1 min at 30 Hz; Retsch) and thawed with nutation. Samples were thawed in 1 ml lysis buffer supplemented with protease inhibitors and 1% CHAPS (Sigma Aldrich) at 4 °C for 1 h. All samples were then clarified by centrifugation at 14,000g at 4 °C for 15 min. The remaining supernatant was added to GFP-trap (Chromotek) for 1 h followed by centrifugation at 1,000g at 4 °C for 3 min, and the supernatant was set aside as the flow through. Beads were washed three times with lysis buffer supplemented with protease inhibitors, and bound proteins were released from the beads by a 5-min incubation at 95 °C in sample buffer. The total protein lysate, the flow through and the immunoprecipitation (IP) fraction were analysed by western blotting. Yeast proteins were extracted by either NaOH or TCA protocol as previously described9, 35 and resolved on polyacrylamide gels, transferred to nitrocellulose membrane blots, and probed with primary rabbit/mouse antibodies against HA (BioLegend, 901502), GFP (Abcam ab290), RFP (Abcam ab62341), histone H3 (Abcam ab1791), actin (Abcam ab8224), Sec65 (kindly provided by P. Walter) or Sec61 (kindly provided by M. Seedorf). The membranes were then probed with a secondary goat-anti-rabbit/mouse antibody conjugated to IRDye800 or to IRDye680 (LI-COR Biosciences). Membranes were scanned for infrared signal using the Odyssey Imaging System. Images were transferred to Adobe Photoshop CS6 for slight adjustments to contrast and brightness. Newly synthesized yeast proteins were radioactively labelled in vivo by a 7–10 min pulse with [35S]methionine at either 30 °C or 37 °C. Labelling was stopped by adding to the cells ice-cold TCA to a final concentration of 10%. Cells were then lysed and proteins were immunoprecipitated as previously described36 with antibodies against RFP (Abcam, ab62341), HA (BioLegend, 901502), Kar2 (kindly provided by P. Walter) or CPY (Abcam, ab113685). Protease inhibitors (complete EDTA-free cocktail; Roche) were used throughout the extraction and immunoprecipitation process. Immunoprecipitated samples were resolved on polyacrylamide gels, which were then exposed to Phosphor Screen (GE Life Sciences) and scanned by phosphorimager. Translocation efficiency was calculated as . The statistical significance of differences was measured using two-tailed student t-test with unequal variance, as indicated in the figure legends. For the Tetp-repression experiments, doxycycline (Sigma-Aldrich) was added to the overnight culture and to the back-dilution medium at a final concentration of 15 μg/ml. The ribosomal subunits RPL16a/b were conjugated to AVI-tag (biotin acceptor peptide), and Sec63 was conjugated to BirA (biotin ligase), allowing the specific biotinylation and streptavidin pull-down of ribosomes in close physical proximity to the ER membrane. By comparing the ribosomal footprints obtained from the total ribosome fraction and the streptavidin-pulled fraction, we measured ER-localized translation enrichment. Biotin induction was carried out at mid-logarithmic growth phase in the presence of cycloheximide, which was added to the medium 2 min before the addition of biotin, at a final concentration of 100 μg/ml. To induce biotinylation, d-biotin was added to the medium to a final concentration of 10 nM and biotinylation was allowed to proceed for 2 min at the same temperature as growth. Cells were harvested by filtration onto 0.45 μm pore size nitrocellulose filters (Whatman), scraped from the membrane, and immediately submerged in liquid nitrogen. The following steps of monosome isolation, streptavidin pulldown of biotinylated ribosomes, and library generation were done as previously described25. Sequencing reads were demultiplexed and stripped of 3′ cloning adapters using in-house scripts. Reads were mapped sequentially to Bowtie indices composed of rRNAs, tRNAs, and finally all chromosomes using Bowtie 1.1.0. Only uniquely-mapped, zero-mismatch reads from the final genomic alignment were used for subsequent analyses. These alignments were assigned a specific P-site nucleotide using a 15-nt offset from the 3′ end of reads. Gene-level enrichments were computed by taking the log ratio of biotinylated footprint density (reads per million) within a gene coding sequence (CDS) over the corresponding density of matched input ribosome-profiling experiment. Yeast genes were excluded from all analysis if they met any of the following criteria: had fewer than 100 CDS-mapping footprints in the input sample of a particular experiment; were annotated as ‘dubious’ in the SGD database; mapped to the mitochondrial chromosome. Additionally, regions in which a CDS overlapped another same-strand CDS were excluded from enrichment calculations. TMD positions were predicted using the Phobius algorithm. TMD classification was divided based on the start site of the first predicted TMD: N-terminal TMDs start in the first 95 amino acids of the protein; downstream TMDs start after the first 95 amino acids of the protein. Genes that were dependent on SND components were identified by comparing the Sec63-BirA ER enrichment in a wild-type strain (yJW1784) with that in a Δsnd strain (yJW1811, yJW1812, or yJW1813) as previously described25. Briefly, log enrichments were separately normalized by subtracting the mean enrichment and dividing by the standard deviation of enrichments for the corresponding experiment. Genes were then binned by the minimum number of sequencing counts in either wild-type or Δsnd input sample, and the difference between normalized enrichments was compared within each bin. Enriched genes were defined as those genes whose Δsnd log enrichments were greater than 0.3 and whose enrichments increased in the Δsnd sample by at least two standard deviations compared to other genes in that bin. Depleted genes were defined as those genes whose wild type log enrichments were greater than 0.3 and whose enrichments decreased in the Δsnd sample by at least two standard deviations compared to other genes in that bin. Significant depletion of 10–23%, 9–42% and 14–45% was observed in Δsnd1, Δsnd2 and Δsnd3, respectively. Including or excluding SS-bearing proteins had no effect on this trend. Mitochondrial proteins were excluded from the analysis. Lysates for immunoprecipitations were prepared from yeast that expressed GFP-tagged SND genes or a constitutively expressed GFP-negative control, in mid-logarithmic growth grown in YPD reach medium. Cells were harvested, washed in distilled water, and resuspended in lysis buffer (50 mM Tris HCl pH 7, 150 mM NaCl) supplemented with protease inhibitors (complete EDTA-free cocktail; Roche) and frozen in a drop-by-drop fashion in liquid nitrogen. Frozen cells were then pulverized in a ball mill (1 min at 30 Hz; Retsch) and thawed with nutation. Samples were thawed in 1 ml lysis buffer supplemented with protease inhibitors and 1% digitonin (Sigma Aldrich) at 4 °C for 1 h. All samples were then clarified by centrifugation at 14,000g at 4 °C for 15 min. The remaining supernatant was added to GFP-trap (Chromotek) for 1 h followed by three washes with lysis buffer supplemented with protease inhibitors and 1% digitonin. Bound proteins were released from the beads by a 5-min acidic treatment (0.2 M glycine pH 2.5), which was neutralized with 1 M Tris pH 9.4. The eluted proteins were digested with 0.4 μg sequencing grade trypsin for 2 h in the presence of 100 μl of 2 M urea, 50 mM Tris HCl pH 7.5 and 1 mM DTT. The resulting peptides were acidified with trifluoroacetic acid (TFA) and purified on C18 StageTips. LC–MS/MS analysis was performed on an EASY-nLC1000 UHPLC (Thermo Scientific) coupled to a Q-Exactive mass spectrometer (Thermo Scientific). Peptides were loaded onto the column with buffer A (0.5% acetic acid) and separated on a 50-cm PepMap column (75 μm i.d., 2 μm beads; Dionex) using a 4-h gradient of 5–30% buffer B (80% acetonitrile, 0.5% acetic acid). Interactors were extracted by comparing the protein intensities to a GFP control. Yeast microsomes were extracted from the ADHp-SND2–GFP/SND3–HA strain as described37. In brief, spheroplasts of yeast were lysed by dounce homogenization (25 strokes) in lysis buffer (0.1 M sorbitol, 20 mM HEPES pH 7.4, 50 mM potassium acetate, 2 mM EDTA, 1 mM DTT, 1 mM PMSF) at 4 °C. The lysates were centrifuged at 1,000g and the resulting supernatant at 27,000g for 10 min at 4 °C. The crude membrane pellet was re-suspended in lysis buffer and layered onto a discontinuous sucrose density gradient consisting of 1.2 and 1.5 M sucrose. Following centrifugation at 100,000g for 60 min at 4 °C, the membranes at the 1.2–1.5 M sucrose interface were collected and washed twice in lysis buffer. The membrane pellets were re-suspended in membrane storage buffer (50 mM NaCl, 0.32 M sucrose, 20 mM HEPES pH 7.4, 2 mM EDTA containing protease inhibitors) and the protein concentration determined by a standard Bradford assay. Microsomes were solubilized in ComplexioLyte 48 buffer (1 mg/ml, Logopharm) for 30 min at 4 °C38. Solubilized extracts were centrifuged at 100,000g for 30 min at 4 °C, supplemented with glycerol (5%) and coomassie G-250 (0.3%) and loaded on a 3.5–15% linear native polyacrylamide gel. The BN-PAGE gel was prepared as described39. The gel buffer contained 25 mM imidazole and 500 mM 6-aminohexanoic acid. The cathode chamber was first filled with cathode buffer B (50 mM Tricine, 7.5 mM imidazole and 0.02% coomassie) and subsequently replaced by cathode buffer B/10 (containing 0.002% coomassie) after the gel running front had covered a third of the desired distance of electrophoresis. The anode chamber was filled with 25 mM imidazole pH 7.0. A high-molecular-weight calibration kit for native electrophoresis from GE Healthcare was used as a standard. For 2D BN-PAGE, the excised lanes were equilibrated in 2D-dissociation buffer (60 mM Tris/HCl pH 6.8, 10% glycerol, 2% SDS, 5% v/v β-mercaptoethanol, 6 M urea) before separation on the second dimension by SDS–PAGE. After electro-blotting, the nitrocellulose membrane was detected with the indicated antibodies. The HEK293 cell line used was obtained from DSMZ (no. ACC 305). DSMZ supplied verification of authentication of the cells, tested by DSMZ via short tandem repeat loci (STR profile). The cell line is routinely tested for mycoplasma contamination. This cell line was chosen as it is routinely used for fractionation experiments. Rough microsomes from human cells were prepared as described40. Briefly, 30 × 106 HEK293 cells were harvested and washed once with PBS and twice with buffer 1 (50 mM HEPES/KOH pH 7.5; 0.25 M sucrose; 50 mM KOAc; 6 mM MgOAc; 4 mM PMSF; 1 mM EDTA; 1 mM DTT; 0.1 mg/ml cycloheximide; 0.3 U/ml RNAsin (Promega); protease inhibitor cocktail). After homogenization in buffer 1 using a glass/Teflon homogenizer, the suspension was centrifuged at 1,000g for 10 min. The supernatant was centrifuged at 10,000g for 10 min. The new supernatant was layered onto 0.6 M sucrose in buffer 2 (50 mM HEPES/KOH pH 7.5, 0.6 M sucrose, 100 mM KOAc, 5 mM MgOAc, 4 mM DTT, 0.1 mg/ml cycloheximide, 40 U/ml RNAsin) and centrifuged at 230,000g for 90 min. The resulting membrane pellet was previously shown to comprise rough ER. Here, it was resuspended in buffer 2 and adjusted to 2.3 M sucrose, which was overlaid with 1.9 and 0 M sucrose, respectively, in buffer 2. After flotation at 100,000g for 18 h, the interphase between 0 and 1.9 M sucrose, two fractions of the remaining supernatant, and the pellet were collected. After centrifugation of the interphase at 100,000g for 1 h, the membrane pellet corresponded to purified rough ER. All steps after the first washing step were carried out on ice. Western blot analyses employed antibodies against β-actin (Sigma), CAML (Synaptic Systems SA7679), or rabbit antibodies that were raised against the depicted proteins: the C-terminal peptide of hSnd2 (KRQRRQERRQMKRL) plus an N-terminal cysteine; or an internal peptide of SRα (KKFEDSEKAKKPVR) plus a C-terminal cysteine, cross-linked to KLH. The SRα and β-actin antibodies were visualized using ECL Plex goat-anti-rabbit IgG-Cy5-conjugate or ECL Plex goat-anti-mouse IgG-Cy3-conjugate (GE Healthcare) and the Typhoon-Trio imaging system (GE Healthcare) in combination with Image Quant TL software 7.0 (GE Healthcare). The hSnd2 and CAML antibodies were visualized using secondary peroxidase (POD)-coupled anti-rabbit antibody (Sigma) plus ECL (GE Healthcare) and the Fusion SL luminescence-imaging system (Peqlab) in combination with Image Quant TL software 7.0. Ribosome-profiling data have been deposited in Gene Expression Omnibus (GEO) under accession number GSE85686. Gel source images can be found in Supplementary Fig. 1. Other data that support the findings of this study are available from the authors on reasonable request.
Zotz J.S.,RWTH Aachen |
Wolbing F.,TU Munich |
Lassnig C.,University of Veterinary Medicine Vienna |
Kauffmann M.,RWTH Aachen |
And 8 more authors.
FASEB Journal | Year: 2016
Antigen-induced mast cell (MC) activation via cross-linking of IgE-bound high-affinity receptors for IgE (FcϵRI) underlies type I allergy and anaphylactic shock. Comprehensive knowledge of FcϵRI regulation is thus required. We have identified a functional interaction between FcϵRI and CD13 in murine MCs. Antigen-triggered activation of IgE-loaded FcϵRI results in cocapping and cointernalization of CD13 and equivalent internalization rates of up to 40%. Cointernalization is not unspecific, because ligand-driven KIT internalization is not accompanied by CD13 internalization. Moreover, antibody-mediated cross-linking of CD13 causes IL-6 production in an FcϵRI-dependent manner. These data are indicative of a functional interaction between FcϵRI and CD13 on MCs. To determine the role of this interaction, CD13-deficient bone marrow-derived MCs (BMMCs) were analyzed. Intriguingly, antigen stimulation of CD13-deficient BMMCs results in significantly increased degranulation and proinflammatory cytokine production compared to wild-type cells. Furthermore, in a low-dose model of passive systemic anaphylaxis, antigen-dependent decrease in body temperature, reflecting the anaphylactic reaction, is substantially enhanced by the CD13 inhibitor bestatin (-5.9 ± 0.6°C) and by CD13 deficiency (-8.8 ± 0.6°C) in contrast to controls (-1.2 ± 1.97°C). Importantly, bestatin does not aggravate anaphylaxis in CD13-deficient mice. Thus, we have identified CD13 as a novel negative regulator of MC activation in vitro and in vivo. © FASEB.
Schwenk J.,Albert Ludwigs University of Freiburg |
Schwenk J.,Center for Biological Signaling Studies |
Perez-Garci E.,University of Basel |
Schneider A.,Albert Ludwigs University of Freiburg |
And 15 more authors.
Nature Neuroscience | Year: 2016
GABAB receptors, the most abundant inhibitory G protein-coupled receptors in the mammalian brain, display pronounced diversity in functional properties, cellular signaling and subcellular distribution. We used high-resolution functional proteomics to identify the building blocks of these receptors in the rodent brain. Our analyses revealed that native GABAB receptors are macromolecular complexes with defined architecture, but marked diversity in subunit composition: the receptor core is assembled from GABAB1a/b, GABAB2, four KCTD proteins and a distinct set of G-protein subunits, whereas the receptor's periphery is mostly formed by transmembrane proteins of different classes. In particular, the periphery-forming constituents include signaling effectors, such as Cav2 and HCN channels, and the proteins AJAP1 and amyloid-β A4, both of which tightly associate with the sushi domains of GABAB1a. Our results unravel the molecular diversity of GABAB receptors and their postnatal assembly dynamics and provide a roadmap for studying the cellular signaling of this inhibitory neurotransmitter receptor. © 2016 Nature America, Inc. All rights reserved.
Bildl W.,Albert Ludwigs University of Freiburg |
Haupt A.,Albert Ludwigs University of Freiburg |
Haupt A.,Logopharm GmbH |
Muller C.S.,Albert Ludwigs University of Freiburg |
And 8 more authors.
Molecular and Cellular Proteomics | Year: 2012
Affinity purification (AP) of protein complexes combined with LC-MS/MS analysis is the current method of choice for identification of protein-protein interactions. Their interpretation with respect to significance, specificity, and selectivity requires quantification methods coping with enrichment factors of more than 1000-fold, variable amounts of total protein, and low abundant, unlabeled samples. We used standardized samples (0.1-1000 fmol) measured on high resolution hybrid linear ion trap instruments (LTQ-FT/Orbitrap) to characterize and improve linearity and dynamic range of label-free approaches. Quantification based on spectral counts was limited by saturation and ion suppression effects with samples exceeding 100 ng of protein, depending on the instrument setup. In contrast, signal intensities of peptides (peak volumes) selected by a novel correlation-based method (TopCorr-PV) were linear over at least 4 orders of magnitude and allowed for accurate relative quantification of standard proteins spiked into a complex protein background. Application of this procedure to APs of the voltage- gated potassium channel Kv1.1 as a model membrane protein complex unambiguously identified the whole set of known interaction partners together with novel candidates. In addition to discriminating these proteins from background, we could determine efficiency, cross-reactivities, and selection biases of the used purification antibodies. The enhanced dynamic range of the developed quantification procedure appears well suited for sensitive identification of specific protein-protein interactions, detection of antibody-related artifacts, and optimization of AP conditions. © 2012 by The American Society for Biochemistry and Molecular Biology, Inc.