Simultaneous in vivo time-lapse stiffness mapping and fluorescence imaging of developing tissue

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 in the developing Xenopus brain, a stiffness gradient evolves over time because of differential cell proliferation. Subsequently, axons turn to follow this gradient, underpinning the importance of time-resolved mechanics measurements.

During embryonic development, many biological processes are regulated by tissue mechanics in vivo, including cell migration 1 , neuronal growth 2 , and large-scale tissue remodelling 3,4 . Recent measurements at specific time points suggested that tissue mechanics change during developmental 2,5,6 and pathological 7,8 processes, which might significantly impact cell function.
Furthermore, several approaches have recently been developed to measure in vivo tissue stiffness, including atomic force microscopy 1,2,9 , magnetic resonance elastography 10 , Brillouin microscopy 11 , and magnetically responsive ferrofluid microdroplets 12 . However, the precise spatiotemporal dynamics of tissue mechanics remains poorly understood, and how cells respond to changes in local tissue stiffness in vivo 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 minutes ( Fig.   1). A fluorescence zoom stereomicroscope equipped with an sCMOS camera (quantum yield 82%) were custom-fitted above a bio-AFM set-up (Supplementary Fig. 1), which had a transparent pathway along the area of the cantilever. To cope with the long working distances 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. (Fig.   1a, b) (see online methods for details).
We tested tiv-AFM using the developing Xenopus embryo brain during outgrowth of the optic tract (OT) as a model (Fig. 1c). In the OT, retinal ganglion cell 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 visual centre of the brain 13 . 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 2 . This gradient strongly correlated with axon turning, with the OT routinely turning caudally towards softer tissue 2 .
We therefore wanted to determine when this stiffness gradient first developed, whether its emergence preceded OT axon turning, and what the origin of that stiffness gradient was.
To answer these questions, we performed iterated tiv-AFM measurements of the embryonic brain in vivo at early-intermediate stages, i.e. 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 a ~150 µm by 250 µm raster at 20 µm resolution every ~35 minutes, producing a sequence of 'stiffness maps' of the area (Supplementary Fig. 2). 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 (Fig. 1d, see online methods for details) 14,15 . Simultaneously, we recorded optical time-lapse images of fluorescently labelled retinal ganglion cell axons growing through the region of interest (Fig. 1d, Supplementary Fig. 2a).
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 (Fig. 1d). 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 minutes after the onset of measurements ( Supplementary Fig. 2c), i.e., 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. At the beginning of each time point in the sequence of tiv-AFM maps, we calculated the size of the angle through which axons turned ('OT turn angle').
For each animal, minimum and maximum absolute values were rescaled to 0 and 1, respectively. The projected appearance of the stiffness gradient significantly preceded the projected onset of axon turning (Fig. 2b), indicating that axons turned after the stiffness gradient was established (Fig. 2a), which is consistent with a role for mechanical gradients in helping to guide OT axons caudally 2 . 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 16 .
We have previously shown that tissue stiffness scales with local cell body density 1,17 , and that in Xenopus embryo brains local stiffness gradients at later developmental stages (39/40) correlate with a gradient in cell density 2 . 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).
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 (Fig. 2c). 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 (Fig. 2d). Plotting the stage-specific gradient in cell body densities against the stiffness gradient revealed a strong linear correlation between them (Pearson's correlation coefficient p = 0.97) (Fig.   2e).
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 18 , which has previously been used to inhibit in vivo cell proliferation in the embryonic retina 19 . In BI2536-treated brains, the nuclear density rostral to the OT and thus the cell density gradient was significantly decreased compared to controls, particularly at later stages (Fig. 2c).
Blocking cell proliferation furthermore significantly attenuated both the stiffness gradient and the OT turn angle (Fig. 2f, g), suggesting that the gradient in cell densities is responsible for the stiffness gradient, which on the other hand instructs axon growth.

Tiv-AFM allows simultaneous time-lapse measurements of cell and tissue mechanics in vivo
and optical monitoring of fluorescently labelled structures at the surface of otherwise optically opaque samples at length and time scales that are relevant for developmental processes. It enabled us for the first time to trace the in vivo mechanical properties of the embryonic Xenopus brain as the embryo developed, and to relate mechanical tissue changes to a key event in axon pathfinding.
More broadly, tiv-AFM can also be easily adapted for in vivo applications in other small organisms, or alternatively in tissues ex vivo. It can be used to study cellular responses to a range of mechanical stimuli via the AFM in vivo, such as sustained compression 1,2 , or to track the temporal mechanical response of tissues or organs to different pharmacological treatments (such as the mitotic inhibitor used here). Additionally, the setup is very versatile and can be further expanded, for example, by combining it with calcium imaging to investigate how cellular activity is regulated by changes in tissue stiffness during development and pathology. Tiv-AFM will greatly expand the range of bio-AFM experiments possible, allowing for more scope both for a detailed characterisation of in vivo tissue mechanics during development and disease progression, and for testing how mechanics shapes cell behaviour and function.    Tissue stiffness changed throughout the time course, with significant changes already occurring between ~40-80 minutes after the start of the experiment. AFM measurement resolution, 20 µm; scale bar, 100 µm.