Further information and requests for resources and reagents should be directed to and will be provided by the Lead Contact, U. Valentin Nägerl ( valentin.nagerl@u-bordeaux.fr ).

Organotypic cultures were established from postnatal day 5 to 7 mice pups killed by swift decapitation. For most slice culture experiments, pups were produced by cross-breeding C57BL/6J (RRID: IMSR_JAX:000664) with homozygous Thy1-YFP-H (RRID: IMSR_JAX:003782) mice, and the hemizygous YFP-expressing pups were used for preparing slices containing YFP-labeled neurons. We also used C57BL/6J wild-type pups that did not express any fluorescent proteins. In a subset of experiments, ZsGreen-labeled astrocytes were imaged in slices obtained from cross-breeding tamoxifen inducible heterozygous GFAP-cre mice (Tg(GFAP-cre/ERT2)1Fki ()) and homozygous Ai6 (RRID: IMSR_JAX:007906) mice.

Mice were housed under 12 h light/12 h dark cycles at 20 to 22°C with ad libitum access to food and water in the animal facility of the Inserm Neurocenter Magendie at the University of Bordeaux, and monitored daily by trained staff. All animals used were free of any disease or infection at the time of experiments. Pregnant females and females with litters were kept in cages with one male. We did not distinguish between males and females among the perinatal pups used for organotypic cultures, as potential anatomical and/or physiological differences between the two sexes were considered irrelevant in the context of evaluating the SUSHI approach for imaging the extracellular space. No animals used had been involved in previous experiments.

All experiments were performed on mouse organotypic hippocampal slice cultures, with the exception of the data of Figure S3 C that were obtained from dissociated rat hippocampal neurons. Experimental procedures were in accordance with the French National Code of Ethics on Animal Experimentation and approved by the Committee of Ethics of Bordeaux. All procedures were in accordance with the guidelines of the European Directive 2010/63/UE.

Method Details

Organotypic slices of hippocampus Gähwiler, 1988 Gähwiler B.H. Organotypic cultures of neural tissue. Gähwiler et al., 1997 Gähwiler B.H.

Capogna M.

Debanne D.

McKinney R.A.

Thompson S.M. Organotypic slice cultures: a technique has come of age. Tønnesen et al., 2011 Tønnesen J.

Nadrigny F.

Willig K.I.

Wedlich-Söldner R.

Nägerl U.V. Two-color STED microscopy of living synapses using a single laser-beam pair. Organotypic slices were isolated from 5-7 day old mouse pups and cultured by the Gähwiler method (). Mouse pups were quickly decapitated and hippocampi were dissected out in cold GBSS (ThermoFisher Scientific) with 10 mM glucose (VWR) and 2 mM kynurenic acid (Sigma-Aldrich). Hippocampi were cut in 350 μm coronal slices using a tissue chopper (McIlwain), and individual slices were placed on 0.01% poly-L-lysine (Sigma-Aldrich) coated coverslips (12 mm x 24 mm; #1.5; CML). Slices were embedded for 30 min in 10 μL chicken plasma (Sigma-Aldrich) followed by 10 μl of 0.2% thrombin (VWR) in GBSS with 10 mM glucose, which coagulated to encapsulate the slice when mixed. The coverslips with the slices were transferred individually to flat-bottomed culturing tubes (110 mm x 16 mm; Dutscher) with 750 μl medium containing 50% Eagle’s basal medium (ThermoFisher Scientific), 25% HBSS, and 25% horse serum (ThermoFisher Scientific), supplemented with glutamine (ThermoFisher Scientific) and glucose. The tubes were sealed and placed in a roller drum at 10 rotations per hour at 35°C. Once a week 500 μL of medium was changed in the tubes. For experiments, a given coverslip with a slice was mounted in a perfused imaging chamber, and the slice was imaged from below through the glass coverslip, while it could be approached with electrophysiology electrodes from the top.

Primary hippocampal neuron culture Kaech and Banker, 2006 Kaech S.

Banker G. Culturing hippocampal neurons. Primary rat hippocampal neurons were grown as Banker cultures in the proximity of supporting astroglia feeder cells (). For harvesting primary neurons embryonic day 18 fetuses were dissected from pregnant Sprague-Dawley female rats after CO 2 -inhalation anesthesia and swift decapitation, and hippocampi were isolated in HBSS without calcium, magnesium and bicarbonate (HBSS-), supplemented with 10 mM HEPES (ThermoFisher Scientific) and penicillin-streptomycin. After chopping with scissors, hippocampi were digested in HBSS- with 10 mM HEPES and 0.25% trypsin, followed by trituration in HBSS- with 10 mM HEPES. Primary neurons were plated on 18 mm poly-L-lysine coated glass coverslips at a density of 50,000 cells per coverslip. Three hours after plating, coverslips were flipped into 60-mm culturing dishes containing a previously established astroglia cell layer and grown in Neurobasal medium (ThermoFisher Scientific) supplemented with 2 mM L-glutamine (ThermoFisher Scientific) and 1 x NeuroCult SM1 Neuronal supplement (STEMCELL Technologies) at 37°C and 5% CO 2 . Astrocyte feeder layers were prepared from similar embryos 2-3 weeks prior, by triturating the cerebral hemispheres in HBSS- with 10 mM HEPES, 0.5% trypsin and 0.1% DNase. Astrocytes were plated at 20,000 to 40,000 cells per 60-mm dish and cultured in MEM (Fisher Scientific) containing 4.5 g/l glucose, 2 mM L-glutamine and 10% horse serum until co-culturing with the primary neurons. All imaging experiments of primary neurons were performed at days 17-21 post plating.

3D-STED microscope Tønnesen et al., 2011 Tønnesen J.

Nadrigny F.

Willig K.I.

Wedlich-Söldner R.

Nägerl U.V. Two-color STED microscopy of living synapses using a single laser-beam pair. Urban et al., 2011 Urban N.T.

Willig K.I.

Hell S.W.

Nägerl U.V. STED nanoscopy of actin dynamics in synapses deep inside living brain slices. The custom-built STED microscopy setup was constructed around the body of an inverted microscope (Leica DMI6000 CS), as previously described (). The inverted microscope provided ideal imaging conditions by allowing imaging directly through the glass coverslip. We used either an oil objective (TIRF, HCX PL APO, 100X/1.47 NA, Leica), which is optimal for imaging within the first few micrometers of tissue beyond the glass interface, or a glycerol objective (HC PL APO 63X/1.30 NA, Leica) equipped with a correction collar to reduce spherical aberrations, extending depth penetration to tens of micrometers inside the tissue (). The excitation light was provided by a pulsed diode laser (λ = 485 nm, pulse width = 70 ps; PicoQuant, Berlin, Germany) triggered by TTL pulses from the STED laser source at 80 MHz. A variable delay was added to the TTL train to ensure that the synchronized STED and excitation pulses were optimally aligned in time. The STED beam was derived from Ti:Sa femtosecond laser source with a repetition rate of 80 MHz and emitting at 838 nm (MaiTai, Spectra-Physics), which was converted to λ = 597 nm by an optical parametric oscillator (OPO; APE, Berlin, Germany). The 597 nm wavelength of the STED beam corresponds to the far right emission spectrum of green/yellow fluorophores such as YFP and GFP, where it induces stimulated emission, but little direct excitation. The STED pulses were broadened in time to ensure effective de-excitation and maximal resolution enhancement. The femtosecond STED pulses were pre-stretched to around 2 ps by passing the beam through a 25 cm glass rod (SF6), and then broadened further to around 70 ps by passing it through a 20 m long polarization maintaining single-mode fiber (Pure silica core, NA 0.12, Schäfter & Kirchhoff). The pre-stretching was done to protect the optical fiber from photo damage and to avoid non-linear effects in the fiber by high peak powers. Wildanger et al., 2009 Wildanger D.

Medda R.

Kastrup L.

Hell S.W. A compact STED microscope providing 3D nanoscale resolution. Spatial resolution enhancement in three dimensions was achieved by splitting the STED beam into two beams by means of a polarizing beam splitter and imposing separate phase delays on their wave fronts. One beam was shaped to enhance lateral resolution by creating a “doughnut” PSF, while the other beam enhanced axial resolution by a “bottle beam” PSF (). The doughnut PSF was created by introducing a 2π delay vortex phase mask in the beam path (RPC Photonics, Rochester, NY, USA), while the bottle beam PSF was created by a 2 mm diameter annular π delay phase mask. The width of the bottle beam was adjusted by a telescope so that half of the beam (in terms of power) passed through the circular delay pattern to yield a symmetric PSF with zero intensity at its focus. By placing a rotatable λ/2-wave plate in front of the polarizing beam splitter that spilt the original beam, we could arbitrarily distribute the laser power between the doughnut and bottle beam. By recombining the doughnut and bottle beams using a second polarizing beam splitter, the compound PSF enhanced the spatial resolution in 3D. The 3D-STED beam was precisely aligned with the excitation beam using a 590 nm short-pass dichroic mirror and a piezo-positioner. The overlaid beams passed through an x-y beam scanner (Yanus scan head, TILL Photonics), whose mirrors were conjugated to the back focal plane of the objective, and scanned across the sample. The fluorescence emission was collected episcopically, descanned, isolated from the incident beams using a 499 long-pass dichroic mirror, and cleaned up using a 525 ± 25 nm band-pass filter. The signal was focused into a multimode optical fiber (62.5 μm core diameter) connected to an avalanche photodiode (APD; SPCM-AQRH-14-FC; PerkinElmer), with the fiber acting as a confocal pinhole to reduce out-of-focus fluorescence. The setup was designed for 3D-STED imaging of green and yellow fluorophores, such as YFP, GFP, calcein green, Alexa Fluor 488, Atto-488 and Atto-514. A second Ti:Sa femtosecond laser beam was co-aligned with the STED and excitation beam using a 680 nm short-pass dichroic mirror and used to acquire 2-photon images (at λ = 935 nm) for comparison purposes, and to perform 2-photon glutamate uncaging at λ = 745 nm. Tønnesen et al., 2011 Tønnesen J.

Nadrigny F.

Willig K.I.

Wedlich-Söldner R.

Nägerl U.V. Two-color STED microscopy of living synapses using a single laser-beam pair. We performed two-color 3D-STED at tens of micrometers inside living brain tissue using the glycerol objective mentioned above. Two fluorophores with partially overlapping excitation and emission spectra (e.g., calcein green and YFP) were imaged simultaneously using a single common excitation/depletion laser beam pair, thereby facilitating spatial and temporal alignment of the two imaging channels and minimizing photon exposure. The sole difference to single color imaging was that the emission signal was split by a 514 nm long-pass dichroic mirror, and the two parts were sent into separate photo detectors (). The imaging depth and z stack stepping intervals were controlled via a piezo-driven objective nano-positioner for z axis scanning (PIFOC, Physik Instrumente). The 2-photon laser intensity was controlled via a Pockels cell (350-80LA Electro-Optic Modulator, Conoptics) that was controlled either manually for adjusting imaging intensities or by voltage pulses (1 ms, variable amplitude) from the electrophysiology amplifier for glutamate uncaging. The STED beam intensity was controlled by a combination of a λ/2-plate and a polarizing beam splitter, while the excitation beam intensity was adjusted by an optical density filter mounted in a motorized linear translation stage. Microscope performance was evaluated by measuring the PSFs of the laser beams, and by measuring the spatial resolution for a defined set of imaging parameters (5 mW STED power at the sample, 20 nm x 20 nm pixels, and 20 μs dwell time). PSFs were visualized by reflecting the individual laser beams on 150 nm gold nano-spheres, while spatial resolution was analyzed by imaging 40 nm fluorescent beads adhering to the surface of a glass coverslip. These beads served as point sources, and their apparent size is a measure of the spatial resolution of the microscope. In addition to imaging beads on coverslip surfaces, gold and fluorescent beads were embedded in agar where the refractive index was adjusted to 1.36 by the addition of glycerol, matching roughly the refractive index of brain tissue. This helped us establish the optimal position of the correction collar of the glycerol immersion objective ( Figure S1 ). The spatial resolution was around 175 nm (x-y) and 450 nm (z) for confocal imaging (λ = 485 nm) and around 260 nm (x-y) and 760 nm (z) for 2-photon imaging (λ = 935 nm). In diffraction-unlimited STED mode, the spatial resolution was around 60 nm (x-y) and 160 nm (z).

Extracellular labeling and imaging For imaging, slices were transferred on their glass coverslip to a heated and perfused imaging chamber (32.5°C; 1-2 mL/min). The perfusion medium (artificial cerebrospinal fluid, ACSF) consisted of 125 mM NaCl, 2.5 mM KCl, 1.3 mM MgCl 2 , 2 mM CaCl 2 , 26 mM NaHCO 3 , 1.25 mM NaH 2 PO 4 , 20 mM d-glucose, 7.5 mM HEPES, 1 mM Trolox, 1 mM ascorbic acid (all from Sigma-Aldrich); 300 mOsm; pH 7.4. The extracellular dye, which was either Alexa Fluor 488 (0.57 kDa Hydrazide or 10 kDa dextran, ThermoFisher), calcein (ThermoFisher), Atto-514, or Atto-488 (both Atto Tec), was added from an ACSF-based 4 mM stock into the bulk ACSF while looping perfusion at a total volume of 20 to 30 mL, or pipetted directly into the chamber while pausing the perfusion. Both strategies worked equally well, with no discernible differences in slice viability for imaging sessions lasting tens of minutes to hours. The final fluorophore concentration was around 40 μM, corresponding to a 1:100 dilution of the stock. This could be adjusted on the fly by adding additional fluorophore or ACSF to the perfusion volume. For two-color imaging of ECS and fluorescently labeled cells, we kept the ECS fluorescence intensity around 50% lower than the intracellular label intensity, so that the two signals could be distinguished based on their relative intensity as well as spectral emission. Spectral detection of the two fluorophores worked well using a 514 nm dichroic mirror that split up the signal into two APDs. Direct injection of the dye into the tissue via patch pipettes provides an alternative ECS labeling strategy that is acute and local and uses up much less fluorophore than bath perfusion. For live-cell imaging, we commonly operated with a doughnut-to-bottle beam power ratio of 1:2, and an overall STED power of around 25 mW measured at the objective back aperture. The beam intensities were adjusted on the fly, individually or for both beams simultaneously, depending on the sample and imaging depth. Pixel sizes were typically 20 nm x 20 nm in STED, and 40 nm x 40 nm in confocal and 2-photon mode, while pixel dwell times were between 10 to 20 μs. When using the glycerol objective equipped with a correction collar we could obtain high-quality SUSHI images up to 50 μm below tissue surface. This depth agreed with the measurements of fluorescent beads embedded in agar described above, where spatial resolution was enhanced as deep as 70 μm in agar.

Electrophysiological whole-cell and field recordings All electrophysiology experiments were obtained by means of a 700B MultiClamp amplifier and a Digidata 1440A digitizer (both from Axon Instruments/Molecular Devices) and using the glycerol objective. Whole-cell patch-clamp recordings of excitatory postsynaptic currents (EPSCs) were obtained from CA1 pyramidal neurons, through glass pipettes with a tip resistance of 3.5 to 4 MΩ. The intracellular solution contained 120 mM cesium-methanesulfonate, 5 mM CsCl, 1 mM EGTA, 4 mM Mg-ATP, 0.3 mM Na-GTP and 10 mM sodium phosphocreatine (all from Sigma-Aldrich), and 5 mM QX-314 (Tocris), adjusted to 290 mOsm/L and pH 7.3. The sampling rate was 10 kHz. To evaluate the effect of extracellular dye labeling on electrophysiological properties of the neurons, ACSF with 40 μM Alexa Fluor 488 was washed in for 10 to 30 min before patch-clamping a CA1 pyramidal neuron. After 10 min of voltage-clamp at −70 mV, we recorded 3 min of spontaneous EPSCs. Control experiments were performed in an identical fashion, in the absence of Alexa Fluor 488 in the ACSF. Input resistance was calculated from the steady state response to a test pulse (−5 mV, 10 ms). Electrophysiological recordings of field potentials were obtained from the CA1 pyramidal cell layer in current-clamp mode using glass pipettes filled with ACSF and a tip resistance of around 2 MΩ. The sampling rate was 5 kHz. All electrophysiological recordings were performed at 32.5°C, as during the imaging experiments.

Analyzing ECS width and volume fraction To measure the geometric width of structures, we obtained the full width at half maximum (FWHM) of Gaussian fits to 3-pixel wide line intensity profiles. The widths of unlabeled neural ‘shadow’ structures appeared as drops in fluorescence signal, and here the fitting was done to downward Gaussian curves. As the added fluorophore rapidly distributed uniformly in the perfusion ACSF, variations in pixel intensities reflected physical displacement of the ACFS within a pixel area, rather than variations in fluorophore concentration. Pixel fluorescence intensity was therefore assumed to scale linearly with the ECS volume fraction in that pixel. To estimate the ECS volume fraction in an image, we assigned to each pixel a value between zero and hundred percent ECS, corresponding to pure ECS, measured in larger identifiable ACSF-filled voids in the neuropil. We then binarized the image, so that pixels with intensities > 50% of the maximal value were considered ECS. The ECS volume fraction was defined as the number of above-threshold pixels divided by the total number of pixels (‘binary’ method; Figure 3 and Figure S6 ). In an alternative approach, we estimated ECS volume fraction by calculating the average intensity of all pixels in a region of interest and dividing it by the maximal intensity value representing pure ECS (‘integrative’ method; Figure S6 ).

Hyper-osmolarity challenge Osmolarity was raised in the perfusion ACSF from 300 mOsm/L to 370 mOsm/L by the addition of NaCl. After 10 minutes of hyper-osmotic perfusion, the perfusion solution was changed back the normo-osmotic ACSF. To make the hyperosmotic solution, NaCl was added to the normal perfusion ACSF from a high molarity stock solution, preserving the concentrations of the other ingredients of the ACSF, including the 40 μM calcein.

Epileptiform activity To induce epileptiform discharges we blocked inhibitory GABA A -receptor transmission by adding 50 μM picrotoxin (PTX, Ascent Scientific) to the fluorescent perfusion ACSF. During the experiment, we recorded time-lapse SUSHI images and concurrent electrophysiological field recordings, as described above.

Two-photon glutamate uncaging Glutamate was uncaged from DNI-Glu-TFA (Femtonics), which was bath applied at 1 mg/mL in the perfusion ACSF. To reduce background neuronal activity, 2 μM TTX was added to the ACSF. The uncaging laser pulses were delivered at a wavelength of 745 nm at 1 ms duration and an intensity of 2.5 to 3.5 mW at the sample. The uncaging power was adjusted to produce consistent responses on dendritic spines of YFP-labeled CA1 neurons at the given imaging depth. Uncaging pulse duration, frequency and power were controlled by a Pockels cell via the electrophysiology amplifier and data acquisition software. To induce structural changes in the ECS we delivered 30 uncaging pulses at 1 Hz, preceded and followed by time-lapse SUSHI at 0.7 frames/second. Control experiments were obtained with identical settings in the absence of the caged glutamate. We analyzed the ECS intensity over the time-lapse frames in a 1 μm2 area centered on the uncaging spot. For each frame, the average intensity inside the area of interest was calculated. We calculated the standard deviation (SD) of the signal for each pixel across the time lapse frames in the area of interest. We compared the average SD of the 1 μm2 area around the uncaging spot to the SD of a control area of identical size within the same image frame (arbitrarily chosen at 2 μm to the right or left of the uncaging spot, avoiding any somas and large dendrites). The SD calculation of the time-lapse series was made using the ‘Z-Project’ function in ImageJ.

Two-photon laser lesions Two-photon laser lesion experiments were performed to induce cell migration. Organotypic slices from wild-type mice were perfused in closed loop with 10 mL ACSF. For SUSHI imaging, calcein was added to the ACSF at a final concentration of 200 μM. To inflict small lesions, the 2-photon laser scanned a 1.5 μm x 1.5 μm area using a wavelength of 810 nm and 80 mW laser power measured at the back aperture of the objective. The field of view was 100 μm x 100 μm, pixel dwell time was 30 μs, and pixel size was 48.5 nm x 48.5 nm.

Image processing Displayed images are single sections taken from z stacks or time-lapse series, as indicated. All morphometric measurements were done on raw images in ImageJ using the ‘Plot Line Profile’ function after drawing 3-pixel-wide straight lines through the area of interest. Gaussian fits were applied either directly in ImageJ, or after importing the line profiles into IGOR Pro. Fluorescence intensity within areas of interest was measured in ImageJ using the rectangular selection tool and the ‘Measure’ function. Displayed images were processed with a 1- or 2-pixel median filter in ImageJ to remove noise. No other image processing, such as deconvolution, was applied. Brightness and contrast were adjusted for each image using ImageJ’s ‘Brightness and Contrast’ function. Two-color images are shown as a merge of the filtered raw channels. The look-up tables (LUT) were grays (inverted) for ECS and orange hot for cellular structures. No spectral unmixing was done, as the fluorophore pairs were well distinguishable in the raw images.