Optogenetic activation of EphB2 receptor in dendrites induced actin polymerization by activating Arg kinase

Abstract

Erythropoietin-producing hepatocellular (Eph) receptors regulate a wide array of developmental processes by responding to cell-cell contacts. EphB2 is well-expressed in the brain and known to be important for dendritic spine development, as well as for the maintenance of the synapses, although the mechanisms of these functions have not been fully understood. Here we studied EphB2's functions in hippocampal neurons with an optogenetic approach, which allowed us to specify spatial regions of signal activation and monitor in real-time the consequences of signal activation. We designed and constructed OptoEphB2, a genetically encoded photoactivatable EphB2. Photoactivation of OptoEphB2 in fibroblast cells induced receptor phosphorylation and resulted in cell rounding ------- a well-known cellular response to EphB2 activation. In contrast, local activation of OptoEphb2 in dendrites of hippocampal neurons induces rapid actin polymerization, resulting dynamic dendritic filopodial growth. Inhibition of Rac1 and CDC42 did not abolish OptoEphB2-induced actin polymerization. Instead, we identified Abelson tyrosine-protein kinase 2 (Abl2/Arg) as a necessary effector in OptoEphB2-induced filopodia growth in dendrites. These findings provided new mechanistic insight into EphB2's role in neural development and demonstrated the advantage of OptoEphB as a new tool for studying EphB signaling.

Keywords: EphB receptor; Optogenetics; Synaptogenesis.

Fig. 1.

Optogenetic activation of EphB2. (A) Schematic illustrations of OptoEphB2 domain structure and the photoactivation process. Blue light illumination induces Cry2 clustering, which results in receptor autophosphorylation (Y, tyrosine; pY, phosphotyrosine) and downstream signaling. ECD, extracellular domain; TM, transmembrane domain; ICD, intracellular domain; Myr, myristoylation signal peptide; FP, fluorescent protein. (B) TIRFM images of optoEphB2-expressing HEK293 cells before and after photoactivation (440 nm, three 250-ms pulses delivered 4.5 s apart), showing optoEphB2 clustering. (C) Left: western blot analysis of whole cell lysates collected from MEFs expressing OptoEphB2-mCherry or KDoptoEphB2-mCherry that were illuminated by blue LED light (∼10⁻² W/cm²), or left in the dark, for 1 min. Right: quantification of OptoEphB2 phosphorylation. Relative tyrosine phosphorylation was assayed in OptoEphB2 immuno-precipitates and quantified by dividing the phosphotyrosine signal by the mCherry signal. Error bars show s.e.m. (n=3). (D) Time-lapse TIRF images of MEFs expressing OptoEphB2-mCherry or KD-optoEphB2-mCherry showing kinase dependent cell-rounding after blue light illumination (10 mW/cm², 50-ms pulses at 3 pulses/min). Black dotted lines trace initial cell area and solid line traces the final cell area. (E) Time-lapse fluorescence images of a MEF cell activated by blue light illumination (100-ms pulses, 6 pulses/min) within the specified region of illumination (ROI, black circle). OptoEphB2 clustering and cell process retraction were spatially restricted to the ROI. Time stamps are relative to the start of blue light illumination.

Fig. 2.

Kinetics of OptoEphB2 activation. (A) Left: western blot (pTyr) of MEF cell lysates after specified time of blue light illumination. Right: quantification of normalized total pTyr immunoactivity in cell lysates (n=3). Solid line denotes exponential fit with a time constant of 49.7 s. (B) OptoEphB2 cluster density (# of cluster/cell area, n=4) in MEFs under blue light activation. The dotted line denotes exponential fit with a time constant of ∼15 s. (C) Quantification of MEF cell rounding kinetics. Cell area was normalized to the mean value prior to photoactivation. Both cells expressing OptoEphB2 (black line, n=6) and the control cells expressing Kinase-dead mutant (gray line, n=7) were show for comparison. (D) Testing of the reversibility of OptoEphB2-induced cell rounding. MEFs expressing OptoEphB2 were stimulated with multiple trains of blue light (6×100-ms pulses, 0.1 Hz), while cell morphology was monitored with TIRFM. Top panel shows selected frames in time-lapse data, showing MEF contraction and recovery after the first round of stimulations, as well as the re-contraction after the second round of stimulations. Bottom panel shows the quantification cell area over time, normalized to the average cell area prior to the first stimulation (2 min). Blue bars denote time for optical stimulation. All error bars denote s.e.m.

Fig. 3.

OptoEphB2 activation in dendrites induces dynamic filopodial protrusions. (A) Time-lapse images of neurons expressing OptoEphB2-Venus, or its mutant variants, and mCherry (as a volume marker). Cells were photo-activated via blue light illumination (50-ms pulses, 3 pulses/min) over the indicated ROI (white circles) in dendritic segments. Images were mCherry fluorescence. Time labels are relative to the start of photoactivation. (B) Images of maximum-intensity projection over time (5-min durations) from the dendritic segments shown in A. (C) Quantification of increased filopodial protrusions. Dendritic areas within the ROI were measured from maximum intensity projections and normalized to measurements before photoactivation. Error bars are s.e.m. (n=32 for OptoEphB2, n=16 for KD-optoEphB2). *P<0.05; t-test, comparing OptoEphB2 to KD-optoEphB2.

Fig. 4.

OptoEphB2 activation in dendrites induces actin polymerization. (A) Time-lapse images of mCherry-Lifeact in the ROI during photoactivation of OptoEphB2. (B) Maximum-intensity projection images of a dendrite undergoing two rounds of OptoEphB2 photo-activation. Neural cells (DIV11) expressing OptoEphB2-Venus and mCherry (shown) were photoactivated over the indicated ROI (dash line). Two rounds of photoactivation were spaced with 20 min of incubation in the dark. (C) Same as in B except the second round of photoactivation was carried out with the presence of CK666 (200 μM). (D) The Kolmogorov–Smirnov plot showing the cumulative probability of the cell area increase (based on maximum intensity projection, n =16 each) from the two-round activation protocol shown in B. (E) Images of mCherry-Lifeact in a dendritic filopodium before (-1:00) and after (8:00) OptoEphB2 photoactivation. (F) Quantification of Lifeact intensity in filopodia. Normalized intensity was calculated cell-by-cell by averaging the intensity in all filopodia (>5 per cell) along the ROIs before and after illumination, normalizing to the pre-illumination value, and averaging between cells. Error bars, s.e.m. (n=11 cells for optoEphB2, n=9 cells for KD-optoEphB2). *P<0.05; t-test.

Fig. 5.

OptoEphB2 interacts with Arg. (A) Rosette assay of whole cell lysates (MEFs expressing OptoEphB2-mCherry or KD-optoEphB2-mCherry) probed with a panel of purified SH2 probes (only Arg-SH2 is shown) as well as anti-mCherry and anti-pTyr. The top graph illustrates the lysates spotting pattern. Pervanadate- and PTP1B-treated samples were spotted at half volume and serves as positive and negative controls, respectively. (B) Quantification of Arg-SH2 and anti-mCherry (represent total OptoEphB2-mCherry expression) bindings to cell lysates. KD, KD-optoEphb2 lysate (with light stimulation). (C) Same as in A except lysates are from cells stimulated with ephrinB1-Fc ligand. Lysates were probed with Arg-SH2, anti-EphB2 and anti-pTyr. (D) Quantification of Arg-SH2 and anti-EphB2 bindings to cell lysates shown in C. (E) Fluorescence images of MEF cells co-expressing Arg-YFP and OptoEphB2-mCherry showing light-induced co-clustering of Arg and OptoEphb2, but not with KD-optoEphB2. Error bars are s.e.m.

Fig. 6.

OptoEphB2-induced dendritic filopodia requires Arg kinase. (A) Blue light induced co-clustering of OptoEphB2 and Arg in dendrites in neuron cells (DV11) expressing OptoEphB2-mCherry and Arg-YFP. (B) Left: images of maximum-intensity projection from neural cells (DIV11) co-expressing OptoEphB2-mCherry and KD-Arg (top) or KD-Src (bottom). Blue-light induced clustering of OptoEphB2 but no filopodial growth in KD-Arg cells. The inhibitory effect was not observed with KD-Src expression. Right: quantification of cell area increase (n=11, based on maximum-intensity projection) in the presence of KD-Arg or KD-Src. (C) Images of maximum-intensity projection from neural cells (DIV 11) co-expressing OptoEphB2-Venus and mCherry. Cells were either treated with GNF2, a specific Arg inhibitor, or DMSO, as controls. The images showed that GNF2 treatment, but not DMSO treatment, abolished OptoEphB2 induced dendritic filopodia growth. (D) Quantification of the effects of GNF2 treatment (n=16) in comparison to DMSO (n=27). Light-induced dendritic area increases were quantified with maximum-intensity projection images (as shown in C). The left panel showed the average values and the right panels showed the cumulative distributions. *P<0.05; t-test. Error bars are s.e.m.

Fig. 7.

OptoEphB2 can induced dendritic filopodia growth without activating Rac1 and CDC42. (A) Time-lapse fluorescence images of neural cell (DIV 10-11) expressing OptoEphB2-mCherry and dominant negative CDC42 (DN-CDC42-YFP, top) or dominant negative Rac1 (DN-Rac1-YFP, bottom). Fluorescence signals were from OptoEphB2. (B) Images of maximum-intensity projection before and after photoactivation (10-15 min) from cells shown in A. (C) Quantifications of cell area increases (based on maximum-intensity projection) for cells expressing DN-Rac1-YFP (n=15), DN-CDC42-YFP (n=13) and YFP only (n=11). Results were compared to control dataset of cells treated with actin polymerization inhibitor Cytochalasin D (n=4). *P<0.05; t-test. Error bars are s.e.m.