Tyrosine Phosphorylation within the Intrinsically Disordered Cytosolic Domains of the B-Cell Receptor: An NMR-Based Structural Analysis

<div><p>Intrinsically disordered proteins are found extensively in cell signaling pathways where they often are targets of posttranslational modifications e.g. phosphorylation. Such modifications can sometimes induce or disrupt secondary structure elements present in the modified protein. CD79a and CD79b are membrane-spanning, signal-transducing components of the B-cell receptor. The cytosolic domains of these proteins are intrinsically disordered and each has an immunoreceptor tyrosine-based activation motif (ITAM). When an antigen binds to the receptor, conserved tyrosines located in the ITAMs are phosphorylated which initiate further downstream signaling. Here we use NMR spectroscopy to examine the secondary structure propensity of the cytosolic domains of CD79a and CD79b <i>in vitro</i> before and after phosphorylation. The phosphorylation patterns are identified through analysis of changes of backbone chemical shifts found for the affected tyrosines and neighboring residues. The number of the phosphorylated sites is confirmed by mass spectrometry. The secondary structure propensities are calculated using the method of intrinsic referencing, where the reference random coil chemical shifts are measured for the same protein under denaturing conditions. Our analysis revealed that CD79a and CD79b both have an overall propensity for α-helical structure that is greatest in the C-terminal region of the ITAM. Phosphorylation of CD79a caused a decrease in helical propensity in the C-terminal ITAM region. For CD79b, the opposite was observed and phosphorylation resulted in an increase of helical propensity in the C-terminal part.</p></div

Secondary structure propensity of CD79b and the effects of phosphorylation.

<p>The ITAM region in the sequence is underlined and all shifts are plotted against their corresponding residue number (<b>A</b>) positive values of secondary chemical shifts (Δδ^Cα–Δδ^Cβ) indicate an overall tendency for α-helical structure with an increased propensity in the region Thr206 to Gly216. (<b>B</b>) change of secondary chemical shifts upon phosphorylation (Δδ^Cα–Δδ^Cβ)_P–(Δδ^Cα–Δδ^Cβ). Positive values for residues Tyr207 to Gly216 indicates an increased helical content in this region following phosphorylation. (<b>C</b>) Δδ^CO secondary chemical shifts. The Δδ^CO shift pattern agrees well with the pattern of (Δδ^Cα–Δδ^Cβ) indicating an overall tendency for α-helical structure (<b>D</b>) change of secondary chemical shifts (Δδ_P^CO–Δδ^CO). Phosphorylation increases the tendency for α-helical structure in the C-terminal part of the ITAM region.</p

Chemical shift changes induced by tyrosine phosphorylation.

<p>(<b>A</b>) δ−δ_P (black bars) and (δ−δ_P)_UREA (gray bars) of CD79a and CD79b calculated from Cα chemical shifts. For CD79a, significant δ−δ_P values can be observed surrounding Tyr188, Tyr199 and Tyr210 indicating phosphorylation of these sites. For CD79b, such values are observed surrounding Tyr196 and Tyr207. A comparison between δ−δ_P and (δ−δ_P)_UREA reveals that a dominating part of the chemical shift changes induced by phosphorylation is still present in 6 M urea. (<b>B</b>) δ−δ_P (black bars) and (δ−δ_P)_UREA (gray bars) of CD79a and CD79b calculated from CO chemical shifts. Analysis using CO chemical shifts results in similar patterns as Cα. (<b>C</b>) Overlays of ¹H-¹⁵N-HSQC spectra of CD79a_P and CD79b_P (red) and the corresponding spectra of CD79a and CD79b (black). Phosphorylation induces changes in the amide chemical shifts of targeted tyrosines as well as surrounding residues. Phosphorylated tyrosines show a ¹H downfield shift while the direction of the ¹⁵N shifts varies. Upon phosphorylation, the amide peaks of the tyrosines tend to move into already crowded areas of the spectra.</p

Secondary structure propensity of CD79a and the effects of phosphorylation.

<p>The ITAM region in the sequence is underlined and all shifts are plotted against their corresponding residue number (<b>A</b>) (Δδ^Cα–Δδ^Cβ) secondary chemical shifts. CD79a has an overall tendency for α-helical structure with an increased propensity in the regions Arg166 to Gly175 and Asp194 to Gly205. (<b>B</b>) (Δδ^Cα–Δδ^Cβ)_P–(Δδ^Cα–Δδ^Cβ) secondary chemical shifts. Negative values in the C-terminal part of the ITAM indicate decreased helicity in this region following phosphorylation. (<b>C</b>) Δδ^CO secondary chemical shifts. The Δδ^CO shift pattern agrees well with the pattern of (Δδ^Cα–Δδ^Cβ) indicating an overall tendency for α-helical structure (<b>D</b>) (Δδ_P^CO–Δδ^CO) secondary chemical shifts. Phosphorylation decreases the tendency for α-helical structure in the C-terminal part of the ITAM region.</p

Identification of the Binding Site of Chroman-4-one-Based Sirtuin 2‑Selective Inhibitors using Photoaffinity Labeling in Combination with Tandem Mass Spectrometry

Photoaffinity labeling (PAL) was used to identify the binding site of chroman-4-one-based SIRT2-selective inhibitors. The photoactive diazirine <b>4</b>, a potent SIRT2 inhibitor, was subjected to detailed photochemical characterization. In PAL experiments with SIRT2, a tryptic peptide originating from the covalent attachment of photoactivated <b>4</b> was identified. The peptide covers both the active site of SIRT2 and the proposed binding site of chroman-4-one-based inhibitors. A high-power LED was used as source for the monochromatic UV light enabling rapid photoactivation

Detection and relative quantification of wild-type and mutated embryonic MyHC.

<p>(A and B) Extracted ion chromatograms (XICs) for wild-type and (C and D) mutant embryonic MyHC tryptic peptides eluting at 28.6 and 39.9 minutes are shown as indicated. Note that panels A and C have been adjusted to equal intensity, as have panels B and D.</p

Formation of sarcomere structure of 6-day differentiated myotubes.

<p>(A) Triple staining was performed for α-actinin (green), F-actin (red), and myosin (magenta); (B) M-protein (green), F-actin (red), and myosin (magenta); (C) myomesin (green), F-actin (red), and M-band epitope of titin (magenta). The results were visualized with a Zeiss Axio Observer microscope (Carl Zeiss AG, Germany) at 63× magnification. All nuclei were counterstained with DAPI (blue). The repetitive well-structured sarcomere can be seen clearly in control myotubes and patient myotubes (insets). The bars represent 10 μm.</p

Co-immunofluorescence analysis of 6-day differentiated myotubes.

<p>(A) Triple staining was performed for α-actinin (green), F-actin (red) and M-band epitope of titin (magenta) (A); (B) myomesin (green), F-actin (red) and Z-disc epitope of titin (magenta); (C) myomesin (green), F-actin (red) and A/I junction epitope of titin (magenta). The results were visualised with a Zeiss Axio Observer microscope (Carl Zeiss AG, Germany) at 63× magnification. All nuclei were counterstained with DAPI (blue). The Z-disc can be seen clearly with α-actinin and Z-disc epitope of titin, and the M-band with myomesin, M-band epitope of titin (insets). Note the strikingly thinner myotubes in the patient compared to the cells in the control. The bars represent 10 μm.</p

Mutation determination and quantitative analysis of RNA.

<p>(A) Sequence chromatograms of the entire exon 5 and part of exon 6 of <i>MYH3</i> in the control (left panel) and the patient who was heterozygous for the <i>MYH3</i> mutation p.Thr178Ile (c.602C>T), which changes threonine at position 178 of MyHC to isoleucine. The p.Thr178Ile mutation is shown in the reverse orientation. (B) Analysis of PCR-amplified and <i>BsawI</i>-treated fragments derived from <i>MYH3</i> alleles in differentiating myoblasts from a control (left panel) and from the patient (right panel), using 2% agarose. Treatment of an amplified 308-bp fragment of the wild-type <i>MYH3</i> allele with <i>BsaWI</i> generated 210-bp and 98-bp fragments, whereas the c.602C>T mutation eliminated the cleavage site in the mutated <i>MYH3</i> allele. (C) Illustration of quantitative analysis of the relative expression of mutated and normal <i>MYH3</i> alleles in myoblasts and differentiating skeletal myoblasts from control and patient. This was based on treatment of reverse transcriptase (RT)-PCR-amplified <i>MYH3</i>-derived fragments with <i>BsaWI</i>. The fluorescent PCR products were separated in polyacrylamide gels and the intensities of the respective peaks were analysed. (D) Illustration of quantitative analysis of the relative expression of myosin heavy chain (MyHC) I (<i>MYH7</i>), MyHC IIa (<i>MYH2</i>), MyHC IIx (<i>MYH1</i>), embryonic MyHC (<i>MYH3</i>) and perinatal MyHC (<i>MYH8</i>) transcripts in differentiating skeletal myoblasts from control and patient, based on RT-PCR. (E) Levels of <i>MYH3</i> mRNA determined by RT-qPCR with specific Taqman probes and normalised to GAPDH mRNA. The levels of <i>MYH3</i> transcripts from cultured cells (D3, D6, D8, D10, and D15) derived from the patient were compared to those derived from five controls, taken on corresponding days. The number of experiments/differentiated myotubes is indicated in the bars. Bars represent mean ± SD. ***<i>p</i> < 0.001(significant difference compared with controls, one-way ANOVA).</p

Assignment of Saccharide Identities through Analysis of Oxonium Ion Fragmentation Profiles in LC–MS/MS of Glycopeptides

Protein glycosylation plays critical roles in the regulation of diverse biological processes, and determination of glycan structure–function relationships is important to better understand these events. However, characterization of glycan and glycopeptide structural isomers remains challenging and often relies on biosynthetic pathways being conserved. In glycoproteomic analysis with liquid chromatography–tandem mass spectrometry (LC–MS/MS) using collision-induced dissociation (CID), saccharide oxonium ions containing <i>N</i>-acetylhexosamine (HexNAc) residues are prominent. Through analysis of beam-type CID spectra and ion trap CID spectra of synthetic and natively derived N- and O-glycopeptides, we found that the fragmentation patterns of oxonium ions characteristically differ between glycopeptides terminally substituted with GalNAcα1-<i>O</i>-, GlcNAcβ1-<i>O</i>-, Galβ3GalNAcα1-<i>O</i>-, Galβ4GlcNAcβ-<i>O</i>-, and Galβ3GlcNAcβ-<i>O</i>- structures. The difference in the oxonium ion fragmentation profiles of such glycopeptides may thus be used to distinguish among these glycan structures and could be of importance in LC–MS/MS-based glycoproteomic studies