Sialic Acid Glycobiology Unveils Trypanosoma cruzi Trypomastigote Membrane Physiology.

Trypanosoma cruzi, the flagellate protozoan agent of Chagas disease or American trypanosomiasis, is unable to synthesize sialic acids de novo. Mucins and trans-sialidase (TS) are substrate and enzyme, respectively, of the glycobiological system that scavenges sialic acid from the host in a crucial interplay for T. cruzi life cycle. The acquisition of the sialyl residue allows the parasite to avoid lysis by serum factors and to interact with the host cell. A major drawback to studying the sialylation kinetics and turnover of the trypomastigote glycoconjugates is the difficulty to identify and follow the recently acquired sialyl residues. To tackle this issue, we followed an unnatural sugar approach as bioorthogonal chemical reporters, where the use of azidosialyl residues allowed identifying the acquired sugar. Advanced microscopy techniques, together with biochemical methods, were used to study the trypomastigote membrane from its glycobiological perspective. Main sialyl acceptors were identified as mucins by biochemical procedures and protein markers. Together with determining their shedding and turnover rates, we also report that several membrane proteins, including TS and its substrates, both glycosylphosphatidylinositol-anchored proteins, are separately distributed on parasite surface and contained in different and highly stable membrane microdomains. Notably, labeling for α(1,3)Galactosyl residues only partially colocalize with sialylated mucins, indicating that two species of glycosylated mucins do exist, which are segregated at the parasite surface. Moreover, sialylated mucins were included in lipid-raft-domains, whereas TS molecules are not. The location of the surface-anchored TS resulted too far off as to be capable to sialylate mucins, a role played by the shed TS instead. Phosphatidylinositol-phospholipase-C activity is actually not present in trypomastigotes. Therefore, shedding of TS occurs via microvesicles instead of as a fully soluble form

Characterization of model bile using fluorescence energy transfer from dehydroergosterol to dansylated lecithin

Fluorescence energy transfer from dehydroergosterol (DHE) to dansylated lecithin (DL) was used to characterize lecithin-cholesterol vesicles in the presence of the bile salt, sodium taurocholate. At lipid concentrations approximating physiological levels, exposure of fluorescently labeled vesicles to the bile salt led to a dose-dependent increase in the DHE-to-DL fluorescence ratio during the first 24 h after mixing. The initial changes in the fluorescence ratio correlated well with conventional turbidity measurements that quantify partial micellization of vesicles as a function of bile salt loading. In addition, fluorescence energy transfer from DHE to DL revealed cholesterol enrichment of vesicles and re-vesiculation of micelles at bile salt loadings for which vesicles and micelles coexisted. Samples containing the cholesterol-enriched vesicle fraction exhibited further increases in the DHE-to-DL fluorescence ratio during a 4-week observation period but only after a significant lag period of several days. The lag period decreased with cholesterol loading, and the increase in the fluorescence ratio always preceded the appearance of microscopic, birefringent, either needlelike or platelike, cholesterol crystals, in samples that were initially supersaturated with cholesterol. Cholesterol crystals were not observed, and the fluorescence ratio did not increase, for any sample that was undersaturated with cholesterol. Taken together, these results suggest that the latter changes in fluorescence are the result of cholesterol nucleation. Fluorescence energy transfer from DHE to DL is therefore a promising technique for the characterization of model bile and, possibly, provides a direct measurement of cholesterol nucleation.link_to_subscribed_fulltex

NECAP 1 Regulates AP-2 Interactions to Control Vesicle Size, Number, and Cargo During Clathrin-Mediated Endocytosis

<div><p>AP-2 is the core-organizing element in clathrin-mediated endocytosis. During the formation of clathrin-coated vesicles, clathrin and endocytic accessory proteins interact with AP-2 in a temporally and spatially controlled manner, yet it remains elusive as to how these interactions are regulated. Here, we demonstrate that the endocytic protein NECAP 1, which binds to the α-ear of AP-2 through a C-terminal WxxF motif, uses an N-terminal PH-like domain to compete with clathrin for access to the AP-2 β2-linker, revealing a means to allow AP-2–mediated coordination of accessory protein recruitment and clathrin polymerization at sites of vesicle formation. Knockdown and functional rescue studies demonstrate that through these interactions, NECAP 1 and AP-2 cooperate to increase the probability of clathrin-coated vesicle formation and to control the number, size, and cargo content of the vesicles. Together, our data demonstrate that NECAP 1 modulates the AP-2 interactome and reveal a new layer of organizational control within the endocytic machinery.</p></div

PHear domain binding to the AP-2 β2-linker.

<p>(A) Affinity selection of Flag-tagged PHear, PHear–Ex, and NECAP 1 from HEK-293–T cells using purified GST or GST fused to the α-ear of AP-2. (B) The coat protein fraction stripped from purified brain CCVs was overlaid with purified GST or GST fused to PHear or PHear–Ex (overlay). (C) Schematic representation of the heterotetrameric AP-2 complex (top) and deletion variants of the two large subunits α-adaptin and β2-adaptin. The amino acid (aa) positions indicate the borders of the respective construct, and their binding properties to PHear and PHear–Ex are indicated. (D and E) Flag-tagged variants of α- and β2-adaptin as indicated were immunoprecipitated from HEK-293–T cell lysates, and equal aliquots were resolved by SDS-PAGE to give four replicates. One was used for Western blotting with Flag antibody. The other three were overlaid with either purified GST or GST fused to PHear or PHear–Ex. The hollow arrowhead indicates the migratory position of the IgG heavy chain, detected on the blot. (F) Affinity selection of endogenous clathrin and NECAP 1 from brain lysate using purified GST or GST-β2-linker. (G) HSQC spectra showing NMR titrations of ¹⁵N-labeled PHear with the β2-linker peptide. (H) Magnitude of the amide chemical shift changes of NECAP 1 PHear residues upon binding of the β2-linker peptide. (I) Molecular surface representation of PHear. Amino acids implicated in β2-linker binding by NMR are labeled and colored. Color shading represents the size of the amide chemical shift changes (red, Δδ>0.3; orange, 0.3>Δδ>0.2; yellow, 0.2>Δδ>0.1 ppm). (J and K) The degree of chemical shift change (Δδ) determined by NMR of I106 (J) and L47 (K) in PHear is plotted against the concentration of the β2-linker peptide added. Red lines are lines of best fit. Calculated affinities (<i>K_D</i>) are given for the peaks indicated. (L) Schematic representation of the AP-2 β2 subunit with a focus on the linker sequence. The trunk and ear regions are represented by circles at the N- and C-terminus, respectively. Peptide motifs for binding to the terminal domain of clathrin heavy chain (LLNLD and DLL) are highlighted in yellow. The bars indicate deletion variants of the β2-trunk+linker (aa 1–668). The numbers on each side represent the amino acid positions of the borders. Variants not binding to PHear and PHear–Ex are indicated by grey bars located above the amino acid sequence, while variants that do interact are indicated by black bars below the amino acid sequence. (M) Flag-tagged variants of β2-adaptin as indicated were immunoprecipitated from HEK-293–T cell lysates and equal aliquots were resolved by SDS-PAGE to give four replicates. One was used for Western blotting with Flag antibody. The other three were overlaid with either purified GST or GST fused to PHear or PHear–Ex. (A and F) Starting material (SM) represents 10% of the sample used in the binding assays.</p

The conserved NECAP 1 N-terminus contributes to AP-2 binding.

<p>(A) Schematic representation of NECAP 1 and deletion variants. The conserved region is in grey, which includes the PHear domain in black, and the C-terminal WxxF-Ac motif is indicated. The numbers represent amino acid positions. (B) ¹H-¹⁵N HSQC of NECAP 1 Ex. (C) Affinity-selection of endogenous AP-2 from COS-7 cells using purified GST or GST fused to PHear, PHear–Ex, or NECAP 1. (D) Affinity-selection of amphiphysin I/II and AP-2 from brain lysate using purified GST or GST fused to PHear, PHear–Ex, or Ex. Relative amounts of bait proteins used are indicated (1×, 2×). (E) Flag-tagged PHear and PHear–Ex were immunoprecipitated from transfected HEK-293–T cells using Flag antibody, and co-immunoprecipitation of endogenous AP-2 was assessed by Western blot. (F) Affinity-selection of AP-2 from brain lysate using purified GST or GST fused to PHear, PHear–Ex, or PHear–Ex K154A, G156S. (G) Sequence alignment of NECAP Ex (mouse residues 129–178) from different species as indicated: magna, <i>Magnaporthe oryzae</i>; neurospora, <i>Neurospora crassa</i>; arabidopsis, <i>Arabidopsis thaliana</i>; oryza, <i>Oryza sativa</i>; Cbriggsea, <i>Caenorhabditis briggsae</i>; Celegans, <i>Caenorhabditis elegans</i>; anopheles, <i>Anopheles gambiae</i>; droso, <i>Drosophila melanogaster</i>; Xenopus, <i>Xenopus laevis</i>; Danio, <i>Danio rerio</i>; schistosoma, <i>Schistosoma japonicum</i>. The numbers indicate amino acid positions within the respective protein. (H) Affinity-selection of amphiphysin I/II and AP-2 from brain lysate using purified GST or GST fused to wild-type PHear (wt) or PHear mutants as indicated. In (C–F) and (H), starting material (SM) represents 10% of the protein amount used in each binding.</p

The number of vesicle formation sites and vesicle size are altered upon NECAP 1 KD.

<p>(A) Equal protein amounts of lysates from COS-7 cells transduced with control virus (shRNAmiR-control) or viruses to KD NECAP 1 (NECAP 1 nt162, NECAP 1 nt220) were processed by Western blot for the indicated proteins, CHC, clathrin heavy chain; EEA1, early endosomal antigen 1. (B) Immunofluorescence analysis of endogenous AP-2 in control and NECAP 1 KD COS-7 cells. The large panel is a magnification of the area boxed in the small AP-2 panel. The bar represents 20 µm in the small and 5 µm in large panels. (C) Quantification of (B). Repeated measures one-way ANOVA followed by Bonferroni's Multiple Comparison Test revealed a significant difference between control and KD cells, ***<i>p</i><0.0001, <i>N</i> = 4. (D) NECAP 1 KD COS-7 cells were processed for immunofluorescence to reveal endogenous AP-2 and clathrin light chains (clathrin). The bar represents 20 µm. In (B and D), GFP is expressed as part of the viral expression cassette to verify transduction. (E) Distribution of Cy3-labeled transferrin endocytosed for 1 min in control (shRNAmiR-control) and NECAP 1 KD (NECAP 1 nt162 and NECAP 1 nt220) COS-7 cells. The bar represents 20 µm. (F and G) 3D superresolution analysis of AP-2-labeled vesicle formation sites in control (F) and NECAP 1 KD cells (G). The large panels show a 4 µm² area with biplane signals rendered to a 250 nm particle size (“confocal”) to illustrate a traditional microscope image, and rendered to a 5 nm particle size (superresolution). The small panels show individual structures from the large panels as indicated in x/y and x/z orientation with signals rendered to a 5 nm particle size. The 5 nm spot size exceeds the localization precision of each point but shows the distribution and numbers of localizations. The scale bar represents 1 µm in the large and 100 nm in the small panels. (H) Quantification of the x, y, and z diameters of deeply invaginated CCPs in control and NECAP 1 KD cells. Unpaired two-tailed <i>t</i> tests revealed significant differences in size for all three dimensions, <i>x</i>-axis, **<i>p</i> = 0.0038; <i>y</i>-axis, ***<i>p</i><0.0001; <i>z</i>-axis, ***<i>p</i><0.0001; shRNAmiR-control: four cells, 40 vesicle formation sites; NECAP 1 nt 220: six cells, 60 vesicle formation sites. (I) Quantification of the number of AP-2 signals detected per vesicle formation site. An unpaired two-tailed <i>t</i> test revealed a significant difference, ***<i>p</i> = 0.0002; shRNAmiR-control: four cells, 40 vesicle formation sites; NECAP 1 nt 220: six cells, 60 vesicle formation sites. (J) Quantification of the x, y, and z diameters of deeply invaginated vesicle formation sites at the bottom (cell surface contacting the cover slip) or top (cell surface facing the culture medium) of NECAP 1 KD cells. Unpaired two-tailed <i>t</i> tests revealed no significant differences in size for all three dimensions, each location: three cells, 23 vesicle formation sites. (K and L) Size analysis of CCPs in control and NECAP 1 KD COS-7 cells measuring the width of CCPs detected by EM. (K) The electron micrographs show representative examples of structures found in control and KD cells. The bar equals 100 nm. (L) The bar graph shows the average width of CCPs binned for pits depths of 0–50 nm, 50–100 nm, and 100+ nm in control and NECAP 1 KD cells. Statistical analysis using a two-tailed Mann Whitney test revealed a significant difference in size between the two populations in each bin, 0–50 nm, *<i>p</i> = 0.0286; 50–100 nm, *<i>p</i> = 0.0370; 100+, *<i>p</i> = 0.0280. (M) Size analysis of CCVs in control and NECAP 1 KD COS-7 cells measuring the membrane-to-membrane diameter of CCVs detected by EM. The electron micrographs show representative examples of structures found in control and KD cells (bars equal 100 nm), and the bar graph shows the size distribution within the whole population of vesicles analyzed in each condition. Statistical analysis using a two-tailed unpaired <i>t</i> test revealed a significant difference in size between the two populations, ***<i>p</i><0.0001, <i>N</i> = 3.</p

Structural evidence for a two-regime photobleaching mechanism in a reversibly switchable fluorescent protein.

International audiencePhotobleaching, the irreversible photodestruction of a chromophore, severely limits the use of fluorescent proteins (FPs) in optical microscopy. Yet, the mechanisms that govern photobleaching remain poorly understood. In Reversibly Switchable Fluorescent Proteins (RSFPs), a class of FPs that can be repeatedly photoswitched between nonfluorescent and fluorescent states, photobleaching limits the achievable number of switching cycles, a process known as photofatigue. We investigated the photofatigue mechanisms in the protein IrisFP using combined X-ray crystallography, optical in crystallo spectroscopy, mass spectrometry and modeling approaches. At laser-light intensities typical of conventional wide-field fluorescence microscopy, an oxygen-dependent photobleaching pathway was evidenced. Structural modifications induced by singlet-oxygen production within the chromophore pocket revealed the oxidation of two sulfur-containing residues, Met159 and Cys171, locking the chromophore in a nonfluorescent protonated state. At laser-light intensities typical of localization-based nanoscopy (>0.1 kW/cm(2)), a completely different, oxygen-independent photobleaching pathway was found to take place. The conserved Glu212 underwent decarboxylation concomitantly with an extensive rearrangement of the H-bond network around the chromophore, and an sp(2)-to-sp(3) hybridization change of the carbon atom bridging the chromophore cyclic moieties was observed. This two-regime photobleaching mechanism is likely to be a common feature in RSFPs from Anthozoan species, which typically share high structural and sequence identity with IrisFP. In addition, our results suggest that, when such FPs are used, the illumination conditions employed in localization-based super-resolution microscopy might generate less cytotoxicity than those of standard wide-field microscopy at constant absorbed light-dose. Finally, our data will facilitate the rational design of FPs displaying enhanced photoresistance