Author response: Rapid changes in tissue mechanics regulate cell behaviour in the developing embryonic brain Article Swipe

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Amelia J. Thompson , Eva K. Pillai , Ivan B. Dimov , Sarah K. Foster , Christine E. Holt , Kristian Franze · YOU? · · 2018 · Open Access · · DOI: https://doi.org/10.7554/elife.39356.019

Article Figures and data Abstract eLife digest Main text Data availability References Decision letter Author response Article and author information Metrics Abstract Tissue mechanics is important for development; however, the spatio-temporal dynamics of in vivo tissue stiffness is still poorly understood. We here developed tiv-AFM, combining time-lapse in vivo atomic force microscopy with upright fluorescence imaging of embryonic tissue, to show that during development local tissue stiffness changes significantly within tens of minutes. Within this time frame, a stiffness gradient arose in the developing Xenopus brain, and retinal ganglion cell axons turned to follow this gradient. Changes in local tissue stiffness were largely governed by cell proliferation, as perturbation of mitosis diminished both the stiffness gradient and the caudal turn of axons found in control brains. Hence, we identified a close relationship between the dynamics of tissue mechanics and developmental processes, underpinning the importance of time-resolved stiffness measurements. https://doi.org/10.7554/eLife.39356.001 eLife digest Neurons in the brain form an intricate network that follows a precise template. For example, in a young frog embryo, the neurons from the eyes send out thin structures, called axons, which navigate along a well-defined path and eventually connect with the visual centres of the brain. This journey requires the axons to take a sharp turn so they can wire with the right brain structures. Axons find their paths not only by following chemical signals but also by reacting to the stiffness of their environment. In an older frog embryo for instance, the brain is stiffer at the front, and softer at the back. As neurons from the eyes make their way through the brain, they turn to follow this gradient, moving away from stiffer areas towards the softer regions. Here, Thompson, Pillai et al. investigate when and how this stiffness gradient is established in frogs. To do so, a new technique was developed. Called time-lapse in vivo atomic force microscopy, the method measures how brain stiffness changes over time in a live embryo, while also taking images of the growing axons. The experiments show that the stiffness gradient arose within tens of minutes, just as the first 'pioneering' axons from the eyes began to grow across the brain. These axons then responded to the gradient, turning towards the softer tissue. Changes in the number of cells in the underlying brain tissue governed the formation of the gradient, with rapidly stiffening areas containing more cells than those that remained soft. In fact, using drugs that stop cells from dividing reduced both the mechanical gradient and the turning response of the axons. The technique developed by Thompson, Pillai et al. is a useful tool that can help elucidate how variations in stiffness control the brain wiring process. It could also be used to look into how other developmental or regenerative processes, such as the way neurons reconnect after injuries to the brain or spinal cord, may be regulated by mechanical tissue properties. https://doi.org/10.7554/eLife.39356.002 Main text During embryonic development, many biological processes are regulated by tissue mechanics, including cell migration (Barriga et al., 2018), neuronal growth (Koser et al., 2016), and large-scale tissue remodelling (Butler et al., 2009; Munjal et al., 2015). Recent measurements at specific time points suggested that tissue mechanics change during developmental (Koser et al., 2016; Iwashita et al., 2014; Majkut et al., 2013) and pathological (Murphy et al., 2011; Moeendarbary et al., 2017) processes, which might significantly impact cell function. Furthermore, several approaches have recently been developed to measure in vivo tissue stiffness, including atomic force microscopy (Barriga et al., 2018; Koser et al., 2016; Gautier et al., 2015), magnetic resonance elastography (Sack et al., 2008), Brillouin microscopy (Scarcelli and Yun, 2012), and magnetically responsive ferrofluid microdroplets (Serwane et al., 2017). However, the precise spatiotemporal dynamics of tissue mechanics remains poorly understood, and how cells respond to changes in local tissue stiffness in vivo is largely unknown. To enable time-resolved measurements of developmental tissue mechanics, we here developed time-lapse in vivo atomic force microscopy (tiv-AFM), a method that combines sensitive upright epifluorescence imaging of opaque samples, such as frog embryos, with iterated AFM indentation measurements of in vivo tissue at cellular resolution and at a time scale of tens of minutes (Figure 1). A fluorescence zoom stereomicroscope equipped with an sCMOS camera (quantum yield 82%) was custom-fitted above a bio-AFM set-up (Figure 1—figure supplement 1), which had a transparent pathway along the area of the cantilever. To cope with the long working distance required for imaging through the AFM head, the microscope was fitted with a 0.125 NA/114 mm WD objective. The AFM was set up on an automated motorised stage containing a temperature-controlled sample holder to maintain live specimens at optimal conditions during the experimental time course. (Figure 1a,b) (see Materials and methods for details). Figure 1 with 1 supplement see all Download asset Open asset Experimental set-up for combined time-lapse in vivo AFM (tiv-AFM). (a) Schematic (not to scale) and (b) photograph of the experimental setup. An AFM with 100 µm z-piezo range is positioned above a temperature-controlled sample chamber containing the specimen. A custom-fit fluorescence zoom stereomicroscope with a long (114 mm) working distance and NA 0.125 objective, connected to a high quantum-efficiency sCMOS camera, is mounted on a custom-built support stand above the AFM head optimised for trans-illumination. The specimen is moved by a motorised x/y stage to allow AFM-based mapping of large areas. (c) (Top) Schematic of a Xenopus embryo, showing both how the brain is prepared for tiv-AFM and rostral-caudal (R/C) and dorsal-ventral (D/V) embryonic axes. All following images of embryonic brains in vivo will have the same orientation. (Bottom) Close-up diagram of the brain, showing the approximate region mapped by AFM (white dashed line), within which optic tract (OT) axons (blue) turn caudally. Also shown are the regions of interest (green boxes) used to calculate brain stiffness rostral and caudal of the OT, and hence the developing stiffness gradient. Red overlaid lines show calculation of the angle through which OT axons turn (turn angle). https://doi.org/10.7554/eLife.39356.003 We tested tiv-AFM using the developing Xenopus embryo brain during outgrowth of the optic tract (OT) as a model (Figure 1c). In the OT, retinal ganglion cell (RGC) axons grow in a bundle across the brain surface, making a stereotypical turn in the caudal direction en route that directs them to their target, the optic tectum (McFarlane and Lom, 2012). We previously demonstrated that by later stages of OT outgrowth (i.e. when axons had reached their target), a local stiffness gradient lies orthogonal to the path of OT axons, with the stiffer region rostral to the OT and softer region caudal to it (Koser et al., 2016). This gradient strongly correlated with axon turning, with the OT routinely turning caudally towards softer tissue (Koser et al., 2016). We therefore wanted to determine when this stiffness gradient first developed, whether its emergence preceded OT axon turning, and what the origin of the stiffness gradient was. To answer these questions, we performed iterated tiv-AFM measurements of the embryonic brain in vivo at early-intermediate stages, that is, just before and during turn initiation by the first 'pioneer' OT axons. The apparent elastic modulus K, which is a measure of the tissue's elastic stiffness, was assessed in an ~150 by 250 µm raster at 20 µm resolution every ~35 min, producing a sequence of 'stiffness maps' of the area (Figure 2—figure supplement 1). The applied force (F = 10 nN) and cantilever probe (r = 18.64 µm) were chosen to measure the stiffness mainly of the top ~20–30 µm of the tissue, within which RGC axons grow (Holt, 1989; Harris et al., 1987). To reduce noise, raw AFM data were interpolated and smoothed in x-, y-, and time dimensions using an algorithm based on the discrete cosine transform (Figure 2a,b, see Materials and methods for details) (Garcia, 2010; Garcia, 2011). Simultaneously, we recorded optical time-lapse images of fluorescently labelled RGC axons growing through the region of interest (Figure 2a, Figure 2—figure supplement 1). Figure 2 with 3 supplements see all Download asset Open asset Development of a stiffness gradient in the Xenopus embryo brain precedes axon turning. (a) Time-lapse stiffness maps obtained from a tiv-AFM experiment, showing outlines of fluorescently labelled OT axons (blue) and processed AFM-based stiffness maps (colour maps) overlaid on images of the brain. Colour maps encode the apparent elastic modulus K, a measure of tissue stiffness, assessed at an indentation force F = 10 nN. The time in minutes on each frame is taken from the timestamp of the first measurement in each successive stiffness map; the corresponding overlaid fluorescence images were obtained simultaneously. (b) Visualisation of fold-changes in brain tissue stiffness from one time point to the next, based on the interpolated and smoothed data shown in Figure 2a. Colour scale encodes the fold-change in K at each location on the stiffness map, expressed relative to the values at the previous time point, with the exception of t = 0 min, where all values were set to 1.Tissue stiffness changes throughout the time course, with large changes already occurring between ~40–80 min after the start of the experiment. (c) Plot of mean re-scaled values for the stiffness gradient (orange) and OT turn angle (blue). Stiffness values were binned to match the time points of the developmental stages at which cell body densities were assessed. Dashed lines denote linear fits (R2 = 0.99). (d) Boxplots of the extrapolated appearance times of the stiffness gradients and the onset of OT axon turning, relative to the start time of tiv-AFM measurements, with ladder plots for individual embryos overlaid (grey circles/dashed lines). Extrapolations are based on linear fits to the re-scaled data for individual animals (Figure 2c). Stiffness gradients appear significantly earlier than the onset of axon turning (p=0.03, paired Wilcoxon signed-rank test). (e) Scatterplot showing the time delay between extrapolated onsets of stiffness gradients and axon turning, calculated for individual animals. The average delay of 18 min is indicated by the blue line. Boxplots show median, first, and third quartiles; whiskers show the spread of the data; '×' indicates the mean. Error bars denote standard error of the mean. *p<0.05. AFM measurement resolution, 20 µm; all scale bars, 100 µm. N denotes number of animals. https://doi.org/10.7554/eLife.39356.005 To assess whether repeated AFM measurements of the Xenopus brain affect RGC axon growth, we first conducted time-lapse AFM measurements on one group of embryos while we exposed another group to the same conditions without making force measurements. At the end of the experiments (i.e., at stage 37/38), OTs were labelled with DiI (Wizenmann et al., 2009), and their elongation and turning angles measured. We did not find any significant differences between the groups (Figure 2—figure supplement 2), suggesting that repeated AFM measurements do not affect axon growth. We then performed tiv-AFM measurements of developing Xenopus brains. Early in the time-lapse sequence (i.e. prior to axon turning), the stiffness of the brain was similar on both sides of the OT. However, over the time course of the measurements a stiffness gradient arose, mostly due to rising stiffness of tissue rostral to the OT (Figure 2a,b). Visual inspection of the fold-change in tissue stiffness from one time point to the next indicated that significant changes in tissue mechanics were already occurring approximately 40–80 min after the onset of measurements (Figure 2b), that is before axons started turning caudally, suggesting that the tissue stiffness gradient was established prior to axon turning. To test this hypothesis, we quantified the temporal evolution of the stiffness gradient in a small region immediately in front of the advancing OT (Figure 2—figure supplement 1a). At the beginning of each time point in the sequence of tiv-AFM maps, we calculated the angle through which axons turned ('OT turn angle'). For each animal, minimum and maximum absolute values were rescaled to 0 and 1, respectively (Figure 2c). The projected appearance of the stiffness gradient preceded the projected onset of axon turning on average by 18 min (Figure 2d,e), indicating that axons indeed turned after the stiffness gradient was established, which is consistent with a role for mechanical gradients in helping to guide OT axons caudally (Koser et al., 2016). Based on the first time point at which we detected axon turning in each animal, our data suggested that a stiffness gradient of at least (0.9 ± 0.4) Pa/µm (mean ± SEM) was required for axons to change their growth direction. In line with this idea, RGC axons from heterochronic eye primordia transplants growing through Xenopus brains at stages before the stiffness gradient is established grow rather straight and do not turn caudally in the mid-diencephalon (Cornel and Holt, 1992). We have previously shown that tissue stiffness scales with local cell body density (Barriga et al., 2018; Koser et al., 2015), and that in Xenopus embryo brains local stiffness gradients at later developmental stages (39-40) correlate with a gradient in cell density (Koser et al., 2016). To determine if changes in cell densities are driving changes in tissue stiffness, and thus parallel the evolution of the stiffness gradient at earlier stages, we assayed cell densities using DAPI labelling of nuclei in whole-mounted brains with fluorescently labelled OTs, beginning at the morphological stage corresponding to the start of tiv-AFM measurements (33/34) and repeated for the two subsequent stages encompassing OT turning (35/36 and 37/38). While at the first stage cell densities on both sides of the OT were similar, a clear difference in nuclear densities rostral and caudal to the OT developed at later stages (Figure 3a). Cell densities at the two later stages were significantly higher in the region rostral to the OT (i.e. where tissue was stiffer) than caudal to it, and the overall magnitude of the cell density gradient significantly rose over time (Figure 3b). Plotting the stage-specific gradient in cell body densities against the stiffness gradient revealed a strong linear correlation between them (Pearson's correlation coefficient ρ=0.97) (Figure 3c). Figure 3 Download asset Open asset Changes in local cell body densities contribute to the emerging in vivo stiffness gradient in the Xenopus embryo brain. (a) Immunohistochemistry of nuclei (green) in whole-mount control Xenopus embryo brains at successive developmental stages. Stages shown correspond to the onset (stage 33/34), approximate middle (stage 35/36), and end (stage 37/38) of tiv-AFM measurements. OT axons are outlined in purple. (b) Local cell body densities were significantly higher rostral to the OT than caudal to it at both stage 35/36 (p=0.03, paired Wilcoxon signed-rank test) and stage 37/38 (p=0.04). (c) Gradients in local cell body density and tissue stiffness strongly correlate with each other (Pearson's correlation coefficient ρ = 0.97). Binned absolute values for the stiffness gradient (in Pa/µm) are plotted against the mean cell density gradient at each developmental stage. Dashed line denotes linear fit (R2 = 0.95). Boxplots show median, first, and third quartiles; whiskers show the spread of the data; '×' indicates the mean. Error bars denote standard error of the mean. *p<0.05. AFM measurement resolution, 20 µm; all scale bars, 100 µm. N denotes number of animals. https://doi.org/10.7554/eLife.39356.009 To test if local cell densities drive the evolution of the stiffness gradient during OT turning, we repeated both nuclear staining and tiv-AFM measurements on embryos treated with the mitotic blocker BI2536 (Lénárt et al., 2007), which inhibits Polo-like kinase 1 and has previously been used to inhibit in vivo cell proliferation in the embryonic retina (Weber et al., 2014). BI2536 also triggered mitotic arrest in brains of developing Xenopus embryos, as the number of phosphorylated histone H3 (pH3)-positive cells (Hugle et al., 2015) was significantly higher in treated compared to control brains, and the cross-sectional area of treated brains was significantly decreased at later stages, indicating a decrease in total cell number (Figure 4a–c). Inspection of the stage-dependent distribution of cells in the developing Xenopus brain suggested that in BI2536-treated brains, the nuclear density was decreased particularly rostral to the OT (Figure 4d, Figure 4—figure supplement 1). In tiv-AFM experiments, blocking cell proliferation significantly attenuated the increase of both the stiffness gradient and the OT turn angle over the time course of the experiment (Figure 4e,f), suggesting that the gradient in cell densities strongly contributed to the stiffness gradient, which in turn helps instruct axon growth. Figure 4 with 3 supplements see all Download asset Open asset Blocking mitosis in vivo reduces local cell density, decreases mechanical gradients, and attenuates both RGC axon turning and overall OT elongation. (a) Coronal sections of stage 37/38 control and BI2536-treated embryos stained for DAPI and phospho-histone3 (pH3). (b) Boxplot of normalized exposed brain area. The schematic demonstrates how regions were normalized. Normalized brain area is significantly lower in the BI2536-treated embryos compared with controls (p=5.7e-05, Wilcoxon rank-sum test), indicating a decrease in total cell number. (c) The number of pH3+ cells per 10 000 µm2 brain area in BI2536-treated embryos increased significantly over time if compared to controls (p=2.2e-04 at stage 37/38, Wilcoxon rank-sum test). (d) Immunohistochemistry of nuclei (green) in whole-mount mitotic inhibitor-treated Xenopus embryo brains at successive developmental stages. Stages shown correspond approximately to the onset (stage 33/34), middle (stage 35/36), and end (stage 37/38) of tiv-AFM measurements. OT axons are outlined in purple. (e) Plots of the fold-change over time in stiffness gradient (orange) and OT turn angle (blue) for both control and mitotic inhibitor-treated embryos. Blocking mitosis significantly attenuated the rise in both stiffness gradient (p=0.01, linear regression analysis) and OT axon turning (p=0.02). Solid and dashed lines denote linear fits for control and inhibitor-treated embryos, respectively. (f) Time-lapse AFM montages showing fold-changes in brain stiffness in representative control (top; OT axons outlined in blue) and mitotic inhibitor-treated embryos (bottom; OT axons in magenta). The colour scale encodes the fold-change in K at each location on the stiffness map, expressed relative to the values obtained at t = 0 min. (g) Boxplot of OT elongation at stage 37/38. Treatment with BI2536 significantly reduced OT elongation compared to controls (p=0.001, Wilcoxon rank-sum test). Boxplots show median, first, and third quartiles; whiskers show the spread of the data; 'o' indicates the mean. Error bars denote standard error of the mean. *p<0.05, **p<0.01, ***p<10−3. AFM measurement resolution, 20 µm; all scale bars, 100 µm. N denotes number of animals except in (b) and (c) where it denotes number of sections. https://doi.org/10.7554/eLife.39356.010 The absence of an increase in tissue stiffness near the advancing OT in BI2536-treated brains was furthermore accompanied by a decrease in OT elongation (Figure 4g). Similarly, OT elongation decreased when brains were softened by manipulating the extracellular matrix (Koser et al., 2016), confirming that tissue stiffness is involved in regulating axon growth in vivo. We obtained similar results using a different mitotic blocker, hydroxyurea/aphidicolin (HUA), which inhibits DNA replication (Gilman et al., 1980; Ikegami et al., 1978) and has previously been used to block cell division in Xenopus embryos (Harris and Hartenstein, 1991). HUA also decreased the gradient in nuclear densities (mainly by reducing nuclear densities rostral to the OT), decreased brain stiffness, and generated defects in OT outgrowth (Figure 4—figure supplement 2). In order to test if the mitotic blocker BI2536 perturbs neuronal mechanosensing, we cultured eye primordia on laminin-coated polyacrylamide substrates of different stiffnesses (Koser et al., 2016), exposed the outgrowing RGC axons to the drug, and quantified axon growth as a function of substrate stiffness. While RGC axon growth on stiff substrates with a shear modulus G'~5,500 Pa was not altered by the presence of BI2536 if compared to control conditions (Figure 4—figure supplement 3e), axons grew longer on soft substrates of G'~200 Pa when they were exposed to 50 nM BI2536 but not when they were exposed to 500 nM (Figure 4—figure supplement 3f), suggesting that the drug might partially impact axon growth on very soft substrates (although no significant differences were observed when axons were grown on slightly stiffer substrates of G'~300 Pa, Figure 4—figure supplement 3d). However, similar to control conditions, axons exposed to different concentrations of BI2536 grew significantly longer on stiff substrates compared to softer substrates. Hence, RGC axons responded to substrate stiffness despite the presence of the mitotic blocker (Figure 4—figure supplement 3g). Our data show that, during early embryonic development, local tissue stiffness may change significantly within only tens of minutes (Figure 2), leading to heterogeneous stiffness distributions which, in the developing Xenopus brain, impact RGC axon growth. These stiffness heterogeneities were largely governed by differential cell proliferation (Figures 3 and 4), which is in agreement with previously published correlations between cell body densities and tissue stiffness (Barriga et al., 2018; Koser et al., 2016; Koser et al., 2015; Weber et al., 2017). Mitotic blockers almost completely abolished the rise of the stiffness gradient (Figure 4), which was accompanied by a decrease in turning angle of the OT, emphasizing the importance of the control of local cell proliferation for mechanosensitive cellular processes in vivo. As changes in substrate stiffness have been shown to promote cell proliferation in vitro (Georges and Janmey, 2005), an increase in cell density might lead to a mechanical positive feedback loop, facilitating further cell proliferation. Perturbing cell division, on the other hand, might alter not only local tissue stiffness but also topological in the tissue, which may also important signals regulating axon growth. cell rostral to the OT might decrease the of and more for axons to grow to the in OT turning angle (Figure Our of the correlation between local cell body density gradients and stiffness gradients that other may contribute to the stiffness gradient as (Figure to a cells et al., 2015) as as of the extracellular matrix et al., 2017). time-lapse measurements of tissue mechanics in vivo and optical of fluorescently labelled at the of opaque samples, at and time scales that are for developmental It for the first time to the in vivo mechanical of the embryonic Xenopus brain as the embryo developed, and to changes in tissue mechanics to a in axon As in all other AFM in tiv-AFM are applied to sample it to biological processes that within tens of away from the surface, such as OT elongation in developing Xenopus embryos. of the Xenopus brain at cellular resolution an which the maximum temporal resolution that can be of approaches may enable measurements within at higher However, tiv-AFM in its form revealed that significant changes in tissue stiffness in vivo within tens of minutes, and that these changes have significant for a biological the outgrowth of RGC axons along the developing embryonic brain. tiv-AFM can also be for in vivo in other small or in vivo. It can be used to cellular to a range of mechanical the AFM in such as (Barriga et al., 2018; Koser et al., 2016), or to the temporal mechanical response of or to different as the mitotic used the is very and can be further for example, by combining it with imaging to investigate how cellular is regulated by changes in tissue stiffness during development and will the range of bio-AFM experiments for more both for a of in vivo tissue mechanics during development and and for how mechanics cell and function. Materials and methods All and were obtained from et al., et al., control of sample Xenopus H3 of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide of polyacrylamide is used for OT stage of AFM raw mapping of stiffness maps brains and OT and local tissue stiffness gradient can be found at and at and for AFM-based stiffness µm to AFM of polyacrylamide AFM force for time-lapse AFM experiments model All

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Author response: Rapid changes in tissue mechanics regulate cell behaviour in the developing embryonic brain

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Amelia J. Thompson, Eva K. Pillai, Ivan B. Dimov, Sarah K. Foster, Christine E. Holt, Kristian Franze

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Stiffness, Brain tissue, Neuroscience, Embryonic stem cell, Dynamics (music), Xenopus, Anatomy, Computer science, Physics, Biology, Biophysics, Acoustics, Thermodynamics, Biochemistry, Gene

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