Retrograde Flow Requires Actin Depolymerization Figure 7. Jasplakinolide treatment reduces barbed end density at the leading edge. Growth cones were dual-labeled with TRITC- phalloidin to show total filamentous actin (A, C, E, G, I) and Alexa 488 g-actin, which incorporates at barbed ends (B, D, F, H, J). (A–B) Barbed ends are concentrated at the leading edge of the growth cone peripheral domain and with intrapodia (red arrowhead) under control conditions. G-actin incorporation is eliminated in the presence of 5 uM cytochalasin D (D&K), while total F-actin is unaffected (C&K), demonstrating the specificity of this labeling procedure for uncapped barbed ends. 6 min treatment with jasplakinolide reduced barbed end density relative to control (L, compare J and F). After 6 min the change in barbed end density was significantly different from both control and 3 min time points (L, compare F&H vs J). A similar pattern was seen for total filamentous actin (L, compare E&G vs I). K) Control: 40 growth cones, CytoD: 32 growth cones. L) Control: 32 growth cones, Jasp: 39 growth cones, Jasp+bleb: 43 growth cones. Examples in E–J are regions similar in size to the dashed white box in A. Error bars are standard deviations. (*: significance level p,0.05 using a two sample Student’s t test, unequal variance). Scale bars: A&C 10 mm; F 5 mm. doi:10.1371/journal.pone.0030959.g007 which predict gradients of actin monomer in the cell. The shape of the G-actin gradient varies according to the location and mag- nitude of monomer sources (depolymerization sites) and sinks (polymerization sites); however, simulations to date predict lowest G-actin monomer concentration at the leading edge and a ,1.5- fold increase from front to rear [33]. We hypothesized that a decrease in G-actin concentration might contribute to the reduced rates of actin assembly after jasplakinolide treatment; however to our knowledge, quantitative assessment of actin monomer concentration gradients corrected for path-length errors has never been reported in growth cones. To address this problem, growth cones were stained with OG- 488 DNAse1, which reliably reports changes in G-actin content in cells [34]. Cells were treated with vehicle (0.005% DMSO) or jasplakinolide (500 nM) for six minutes prior to visualization of G-actin levels with OG-488 DNAseI. To correct for optical path- length differences and allow calculation of relative G-actin con- centrations, neurons were injected with 10 kD Texas-Red lysine- substituted dextran, an inert cytoplasmic volume marker that can be fixed in place by aldehyde cross-linking. The process of fixation and membrane extraction reduced the absolute intensity of fixed dextran by 17%; however, the measured volume profile of the growth cone was unchanged (Fig. S3). The intensity profiles of cytoplasmic volume and DNAseI were remarkably similar (Fig. 8A, compare dextran and DNAseI panels) suggesting that most of the DNAse1 intensity variation was due to differences in path-length (volume). However, when the DNAseI signal was divided by the dextran (volume) signal, a marked concentration gradient was revealed under control conditions (Fig. 8A, panel 3; intensity profile 8C). Specifically, the average concentration of actin monomer was significantly lower in the transition zone and central domain than in the peripheral domain (Fig. 8C). Within the overall inward facing negative gradient trend, a variety of G-actin concentration profile shapes in individual growth cones was noted (Fig. S4). The monomer gradient collapsed in growth cones treated with jasplakinolide showing that continuous turnover of actin filaments is required for maintenance of the gradient (Fig. 8B–C). These results suggest that in addition to reducing barbed end density, inhibition of filament turnover slows actin filament assembly by lowering monomer concentration in the peripheral domain. Two artifacts associated with the use of DNAseI to label actin monomers may affect the relative concentrations measured here. In addition to binding actin monomers, DNAseI also binds actin filament pointed ends [34,35,36]. Since the peripheral domain and transition zone contains a higher density of actin filaments and pointed ends than the central domain, DNAseI labeling intensity would be expected to be higher here and the actin monomer concentration overestimated. To investigate the magnitude of this artifact, we performed live cell extractions using Triton X-100 to permeabilize cells and remove all soluble G-actin before measuring residual DNAseI binding. DNAseI signals from the permeabilized growth cones were normalized by levels obtained from control growth cones that were fixed before permeabilization and DNAseI labeling (Fig. 8D). DNAaseI labeling in the live cell extracted growth cones was actually lowest in the peripheral domain and transition zone where pointed end density would be expected to be high and highest in the central domain which has lower actin filament density. These results suggest DNAseI labeled G-actin is highly extractable in peripheral regions where actin filament density is high. Lower DNAseI extractability in the central domain is in part due to the presence of mitochondria in this region which also bind DNAseI (Fig. 8 A–B, red arrowheads). Note that mitochondria also tend to exclude cytoplasmic volume; thus, their presence in the central domain would have the net effect of raising the measured DNAseI/ Dextran signal and overestimation of actual G-actin concentrations in the central domain. Taking the above into consideration, the DNAseI/Dextran intensity map shown in Fig. 8A is likely an underestimation of the actual G-actin concentration gradient between the peripheral and central domains. Discussion Depolymerization is a distributed process in the peripheral domain Actin network density is highest near the leading edge and markedly lower in proximal regions near the transition zone (Fig. 1A). We found that actin density within an ROI steadily decreased as the ROI moved centripetally with retrograde flow despite net ROI compression due to convergence of network flow. Since our approach measures net turnover, the difference between polymerization and depolymerization, we conclude from the consistent decrease in integrated ROI intensity that depoly- merization outweighs polymerization throughout the peripheral domain away from the leading edge. Our results agree with reports in which actin filament turn- over was measured with single particle tracking of actin speckles [37,38]; however, they contrast with other reports suggesting actin filament turnover is concentrated in the transition zone [23]. We note that the latter study used fluorescent phalloidin as a proxy for polymerized actin. Underestimation of actin filament density near the leading edge may account for this discrepancy since use of phalloidin as an actin probe tends to underestimate the density of newly assembled actin filaments near the leading edge and overestimate the density of older filaments in proximal regions due to the slow binding kinetics of phalloidin [21,39]. Coincident Deceleration of Retrograde Flow and Network Retraction Subsequent to actin turnover inhibition, coincident decreases in retrograde flow and leading edge actin network retraction were observed. Both live cell fluorescent images and electron micro- graphs show a buildup of actin filaments in the transition zone and central domain following turnover inhibition (Fig. 1C & Fig. 2). PLoS ONE | www.plosone.org 9 February 2012 | Volume 7 | Issue 2 | e30959