STED nanoscopy of actin dynamics in synapses deep inside living brain slices

Abstract

It is difficult to investigate the mechanisms that mediate long-term changes in synapse function because synapses are small and deeply embedded inside brain tissue. Although recent fluorescence nanoscopy techniques afford improved resolution, they have so far been restricted to dissociated cells or tissue surfaces. However, to study synapses under realistic conditions, one must image several cell layers deep inside more-intact, three-dimensional preparations that exhibit strong light scattering, such as brain slices or brains in vivo. Using aberration-reducing optics, we demonstrate that it is possible to achieve stimulated emission depletion superresolution imaging deep inside scattering biological tissue. To illustrate the power of this novel (to our knowledge) approach, we resolved distinct distributions of actin inside dendrites and spines with a resolution of 60-80 nm in living organotypic brain slices at depths up to 120 μm. In addition, time-lapse stimulated emission depletion imaging revealed changes in actin-based structures inside spines and spine necks, and showed that these dynamics can be modulated by neuronal activity. Our approach greatly facilitates investigations of actin dynamics at the nanoscale within functionally intact brain tissue.

Figure 1

STED microscopy of hippocampal neurons labeled with Lifeact-YFP. (a) Schematic of the experimental setup. A red-shifted STED beam is superimposed on an excitation beam, thus enabling subdiffraction imaging. The imaged hippocampal slice is maintained in a heated chamber (32°C) and perfused with ACSF. PMF, polarization maintaining optical fiber; MMF, multimode optical fiber; VPP, vortex phase plate; DM, dichroic mirror; RM, resonating mirror; BP, bandpass filter; APD, avalanche photodiode; OBJ, glycerol objective lens; CC, correction collar; MM, micromanipulator. (b) Schematic of Lifeact-YFP binding to actin; low binding affinity leads to constant replenishment of potentially bleached YFP. (c) Schematic of the hippocampal formation of a mouse; CA1 pyramidal neurons were labeled with Lifeact-YFP. (d) Overlay of epifluorescence and transmitted light images of CA1 pyramidal neurons. (e) Two-photon image of a Lifeact-infected CA1 pyramidal neuron (4× zoom into a stretch of dendrite). (f) STED image of dendritic structures of CA1 pyramidal neurons. (g) High magnification of the boxed region in f. (h) Profile of pixel intensity across the neck of a spine following the marked line in g, showing an FWHM of the Lorentzian fit of 67 nm.

Figure 2

Glycerol objective lens with correction collar improves resolution of STED imaging deep inside brain slices. (a) STED images of dendrites belonging to hippocampal neurons recorded at various depths (from left to right: 10, 35, 63, and 78 μm) below the tissue surface. Tissue surface (z = 0 μm) was defined as the position of the first visible labeled structure. (b) Estimated spatial resolution at various depths inside brain tissue. Displayed are the thinnest spine neck diameters (FWHMs) as a function of imaging depth for both confocal (open circles) and STED imaging (open diamonds).The data bins are 20 μm wide, except for the superficial (0–10 μm) and deepest (90–120 μm) layers. The spatial resolution is estimated for each depth interval by averaging the five smallest FWHM values in that data bin. This leads to an average resolution of 190 nm for confocal microscopy (blue) and 60 nm for STED microscopy (red) at depths up to 90 μm. With the correction collar reaching its limit, the resolution starts to deteriorate at depths > 90 μm.

Figure 3

STED microscopy of Lifeact-YFP reveals dynamic organization of actin inside synapses. (a–d) Superresolved live-cell images of actin distributions inside dendritic spines reveal curvilinear bundles linking spine heads to parent dendrites. (e–j) Examples of actin-based substructures that can be seen inside spine necks (e–g) and spine heads (h–j). (k–p) Examples of rapid reorganization of actin distribution inside dendritic spines.

Figure 4

STED imaging of postsynaptic morphological plasticity after chemical LTP stimulation. (a and b) Zoom-in of dendritic spines observed under control conditions (a), as well as before and at subsequent times after chemical LTP stimulation (b). Spine neck diameters are indicated in each frame by arrows. (c) Changes in spine neck diameter after pure ACSF perfusion (control) or chemical LTP treatment. Spine neck growth is shown in red, shrinking in blue, and changes of <10% in gray (paired two-sided t-test, α-error = 0.05; control: p = 0.17, n = 34; LTP: p = 0.00096, n = 41). We checked for normal distribution by examining the residuals and performing a Lilliefors (modified Kolmogorov-Smirnov) test. We assessed the homogeneity of variances by using box plots and performing a Levene test of homogeneity. (d) Ratio of dendritic spines showing a measurable change in spine neck diameter after a certain amount of time (10–30 min) after pure ACSF perfusion (control, n = 74, left) or chemical LTP treatment (n = 64, right).