Evidence for new C-terminally truncated variants of α- and β-tubulins

New C-terminally truncated α- and β-tubulin variants, both ending with an -EEEG sequence, are identified in vivo: αΔ3-tubulin, which has a specific neuronal distribution pattern (distinct from that of αΔ2-tubulin) and seems to be related to dynamic microtubules, and βΔ4-tubulin, corresponding to β2A/B-tubulin modified by truncation of four C-terminal residues, which is ubiquitously present in cells and tissues. Cellular α-tubulin can bear various carboxy-terminal sequences: full-length tubulin arising from gene neosynthesis is tyrosinated, and two truncated variants, corresponding to detyrosinated and Δ2 α‑tubulin, result from the sequential cleavage of one or two C-terminal residues, respectively. Here, by using a novel antibody named 3EG that is highly specific to the -EEEG C-terminal sequence, we demonstrate the occurrence in neuronal tissues of a new αΔ3‑tubulin variant corresponding to α1A/B‑tubulin deleted of its last three residues (EEY). αΔ3‑tubulin has a specific distribution pattern: its quantity in the brain is similar to that of αΔ2-tubulin around birth but is much lower in adult tissue. This truncated α1A/B-tubulin variant can be generated from αΔ2-tubulin by the deglutamylases CCP1, CCP4, CCP5, and CCP6 but not by CCP2 and CCP3. Moreover, using 3EG antibody, we identify a C‑terminally truncated β-tubulin form with the same -EEEG C-terminal sequence. Using mass spectrometry, we demonstrate that β2A/B-tubulin is modified by truncation of the four C-terminal residues (EDEA). We show that this newly identified βΔ4-tubulin is ubiquitously present in cells and tissues and that its level is constant throughout the cell cycle. These new C-terminally truncated α- and β-tubulin variants, both ending with -EEEG sequence, are expected to regulate microtubule physiology. Of interest, the αΔ3-tubulin seems to be related to dynamic microtubules, resembling tyrosinated-tubulin rather than the other truncated variants, and may have critical function(s) in neuronal development

De nouvelles fonctions extraciliaires pour les protéines ciliaires

Les protéines ciliaires ont initialement été caractérisées comme essentielles au bon fonctionnement des cils et sont impliquées dans les ciliopathies. Cependant, certains défauts cellulaires associés à leur dérégulation laissent entrevoir la possibilité de rôles extraciliaires pour ces protéines. En effet, de telles fonctions ont récemment été décrites pour ces protéines, notamment dans le transport vésiculaire, l’orientation du fuseau mitotique et le maintien de la stabilité chromosomique. Ces résultats soulèvent une importante question quant à la contribution de ces fonctions extraciliaires à l’apparition des manifestations cliniques associées aux ciliopathies, comme par exemple la polykystose rénale

Rôle de l'association entre le récepteur adhésif de type N-cadhérine et les microdomaines membranaires

L'adhérence cellulaire médiée par les cadhérines est impliquée dans de nombreux processus cellulaires dont la myogenèse. J'ai montré, dans des myoblastes murins, qu'une partie de ces récepteurs adhésifs est présente dans les microdomaines membranaires enrichis en cholestérol, également appelés lipid rafts. Cette association est localisée au niveau de la jonction inter-cellulaire et participe à la stabilisation de ces récepteurs. De façon intéressante, le complexe adhésif fonctionnel est présent uniquement dans les lipid rafts car l'association de la p120-caténine à la N-cadhérine est retrouvée exclusivement au sein de ces structures membranaires. Finalement, j'ai montré que la formation du complexe N-cadhérine/p120-caténine dans les lipid rafts est nécessaire pour l'activation de la GTPase RhoA en aval de ce récepteur adhésif lors de l'induction de la myogenèseCell-cell adhesion mediated by cadherins is involved in a wide variety of cellular functions such as myogenesis. I have shown, in murine myoblasts, that a fraction of these adhesive receptors is associated with membrane microdomains enriched in cholesterol also called lipid rafts. This takes place mainly at the cell-cell contacts and participates in cadherins stabilization. Interestingly, the functional adhesive complex is present only in lipid rafts since the association between p120-catenin and N-cadherin is exclusively found in these membrane structures. Finally, I have shown that the formation of the N-cadherin/p120-catenin complex occurs in lipid rafts and is required for RhoA GTPase activation downstream of this adhesive receptor during myogenesis inductionMONTPELLIER-BU Sciences (341722106) / SudocSudocFranceF

Reactive oxygen species regulate protrusion efficiency by controlling actin dynamics.

Productive protrusions allowing motile cells to sense and migrate toward a chemotactic gradient of reactive oxygen species (ROS) require a tight control of the actin cytoskeleton. However, the mechanisms of how ROS affect cell protrusion and actin dynamics are not well elucidated yet. We show here that ROS induce the formation of a persistent protrusion. In migrating epithelial cells, protrusion of the leading edge requires the precise regulation of the lamellipodium and lamella F-actin networks. Using fluorescent speckle microscopy, we showed that, upon ROS stimulation, the F-actin retrograde flow is enhanced in the lamellipodium. This event coincides with an increase of cofilin activity, free barbed ends formation, Arp2/3 recruitment, and ERK activity at the cell edge. In addition, we observed an acceleration of the F-actin flow in the lamella of ROS-stimulated cells, which correlates with an enhancement of the cell contractility. Thus, this study demonstrates that ROS modulate both the lamellipodium and the lamella networks to control protrusion efficiency

N-Cadherin Association with Lipid Rafts Regulates Its Dynamic Assembly at Cell-Cell Junctions in C2C12 Myoblasts

Cadherins are homophilic cell-cell adhesion molecules implicated in cell growth, differentiation, and organization into tissues during embryonic development. They accumulate at cell-cell contact sites and act as adhesion-activated signaling receptors. Here, we show that the dynamic assembly of N-cadherin at cell-cell contacts involves lipid rafts. In C2C12 myoblasts, immunofluorescence and biochemical experiments demonstrate that N-cadherin present at cell-cell contacts is colocalized with lipid rafts. Disruption of lipid rafts leads to the inhibition of cell-cell adhesion and disorganization of N-cadherin–dependent cell-cell contacts without modifying the association of N-cadherin with catenins and its availability at the plasma membrane. Fluorescent recovery after photobleaching experiments demonstrate that at the dorsal plasma membrane, lipid rafts are not directly involved in the diffusional mobility of N-cadherin. In contrast, at cell-cell junctions N-cadherin association with lipid rafts allows its stabilization enabling the formation of a functional adhesive complex. We show that lipid rafts, as homophilic interaction and F-actin association, stabilize cadherin-dependent adhesive complexes. Homophilic interactions and F-actin association of N-cadherin are both required for its association to lipid rafts. We thus identify lipid rafts as new regulators of cadherin-mediated cell adhesion

H₂O₂ regulates the contractile machinery of the lamella.

<p>(A) Kymographs and F-actin flow maps computed from quantitative FSM analysis of time-lapse movies of control, blebbistatin (50 µM) and blebbistatin (50 µM)+H₂O₂ 500 µM-treated cells. White lines on kymographs indicate speckle translocation used to calculate flow velocities. Flow rates are color coded, ranging from slow flow in dark blue to fast flow in red. Flow maps have been averaged over 30 frames, i.e., 5 min. (B and C) Average F-actin flow rates measured at the leading edge (B) and 5 µm from the leading edge (C) of control, blebbistatin, H₂O₂ and blebbistatin + H₂O₂-treated cells. n ≥7 cells from at least four independent experiments. Five kymographs/cell were analyzed for each condition. Error bars represent s.e.m. ***, p<0.001 compared to control and blebbistatin (B). **, p<0.001 compared to control and ***, p<0.001 compared to control and H₂O₂ (C). (D) Average lamellipodium width of control, blebbistatin and blebbistatin + H₂O₂-treated cells. n≥7 cells from at least four independent experiments. Five kymographs/cell were analyzed for each condition. Error bars represent s.e.m. ***, p<0.001 compared to control and blebbistatin. (E) Immunolocalization of phosphorylated MLC (pMLC, green) and F-actin phalloidin staining (red) in starved PtK1 cells treated with H₂O₂ 500 µM for the indicated times. The scale bar is 10 µm. Red lines highlight the leading edge of the cells. (F and G) Fluorescence intensity of pMLC (F) and F-actin (G) in cells treated with 500 µM H₂O₂, measured from the cell edge (0 µm) into the cell center (10 µm). (H) pMLC/F-actin fluorescence intensity ratio in cells treated with 500 µM H₂O₂, measured from the cell edge (0 µm) into the cell center (10 µm). In (F-H), the data shown represent one experiment and are averaged from at least 13 cells for each condition. The experiment was repeated three times with similar results (<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0041342#pone.0041342.s002" target="_blank">Figure S2G</a>).</p

ERK is activated in response of H₂O₂ and contributes to H₂O₂-induced protrusion dynamics.

<p>(A) Cell lysates from starved PtK1 cells treated with 500 µM H₂O₂ alone or in combination with ROS scavenger (5 mM) for 0-15-30-45-60 min were immunoblotted with antibodies against pERK and ERK. In (B), the graph represents the averaged normalized pERK values. Data are from four and three independent experiments for H₂O₂ and H₂O₂+ROS scavenger, respectively. Error bars represent s.e.m. *, p<0.05 compared to 0 min H₂O₂ and 15 min H₂O₂+scavenger. (C) Immunolocalization of phosphorylated ERK (pERK, green) and F-actin phalloidin staining (red) in starved PtK1 cells treated with H₂O₂ 500 µM for the indicated times. The scale bar is 10 µm. (D and E) Fluorescence intensity of pERK (D) and F-actin (E) in cells treated with H₂O₂ 500 µM, measured from the cell edge (0 µm) into the cell center (10 µm). (F) pERK/F-actin fluorescence intensity ratio in cells treated with H₂O₂ 500 µM, measured from the cell edge (0 µm) into the cell center (10 µm). In (D)–(F), the data shown represent one experiment and are averaged from at least 13 cells for each condition. The experiment was repeated three times with similar results (<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0041342#pone.0041342.s002" target="_blank">Figure S2D</a>). Protrusion width (G), persistence of protrusion (H) and protrusion/retraction velocities (I) in starved Ptk1 cells incubated for 45 min with control media (+DMSO) or media containing 500 µM H₂O₂ alone or in combination with UO126, a MEK inhibitor. In (G)–(I), the data shown result from the analysis of at least 23 cells and 115 kymographs per condition. Error bars represent s.e.m. ***, p<0.001 compared to control and H₂O₂+UO126.</p

H₂O₂ modulates actin dynamics in PtK1 cells.

<p>(A) Single frames of actin fluorescent speckle time-lapse series of starved PtK1 control (top row) and treated with 500 µM H₂O₂ (bottom row). The scale bar is 10 µm. White arrows highlight the locations used to generate kymographs. (B) Kymographs of control and H₂O₂-treated cells depicted in (A). White lines indicate speckle translocation used to calculate flow velocities. (C) F-actin flow maps computed from quantitative FSM analysis of time-lapse movies of control and H₂O₂-treated cells. Flow rates are color coded, ranging from slow flow in dark blue to fast flow in red. Flow maps have been averaged over 60 frames, i.e., 10 min. (D and E) Average F-actin flow rates measured at the leading edge (D) and 5 µm from the leading edge (E) of control and H₂O₂-treated cells. n = 14 cells analyzed for control and H₂O₂ treatment. Error bars represent s.e.m. ***, p<0.001 compared to control.</p

H₂O₂ increases the formation of free barbed ends and F-actin turnover.

<p>(A) Free barbed end actin incorporation (green) and F-actin phalloidin staining (red) in starved PtK1 cells treated with 500 µM H₂O₂ for the indicated times. The scale bar is 10 µm. (B and C) Fluorescence intensity of free barbed end actin incorporation (B) and F-actin (C) in cells treated with 500 µM H₂O₂, measured from the cell edge (0 µm) into the cell center (10 µm). (D) Fluorescence intensity ratio of free barbed end actin incorporation relative to F-actin in cells treated with 500 µM H₂O₂, measured from the cell edge (0 µm) into the cell center (10 µm). In (B)-(D), the data shown represent one experiment and are averaged from at least 16 cells for each condition. The experiment was repeated four times with similar results (<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0041342#pone.0041342.s002" target="_blank">Figure S2B</a>). (E) F-actin turnover maps computed from quantitative FSM time-lapse movies of starved PtK1 cells treated or not with 500 µM H₂O₂. Turnover maps depict F-actin polymerization (red) and depolymerization (green) rates. Maps have been averaged over 6 frames, i.e., 1 min. The scale bar is 10 µm. Boxed regions are magnified in the bottom left of each panel. n = 14 cells analyzed for control and H₂O₂ treatment.</p