Methods xxx (2015) xxx–xxx

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Review Article

STED microscopy for nanoscale imaging in living brain slices Ronan Chéreau 1, Jan Tønnesen, U. Valentin Nägerl ⇑ Interdisciplinary Institute for Neuroscience, Université de Bordeaux, Bordeaux, France UMR 5297, Centre National de la Recherche Scientifique, Bordeaux, France

a r t i c l e

i n f o

Article history: Received 16 March 2015 Received in revised form 2 June 2015 Accepted 4 June 2015 Available online xxxx Keywords: Superresolution microscopy STED microscopy Brain slices Neurons Dendritic spines Axons

a b s t r a c t Stimulated emission depletion (STED) microscopy was the first fluorescence microscopy technique to break the classic diffraction barrier of light microscopy. Even though STED was conceived more than 20 years ago and acknowledged with the 2014 Nobel Prize in Chemistry, it has not yet been widely adopted in biological research, which stands to benefit enormously from the potent combination of nanoscale spatial resolution and far-field optics. STED microscopy is an ensemble imaging technique that uses a pair of lasers for controlling the excitation state of fluorescent molecules in a targeted manner over nanoscale distances. STED is commonly a point-scanning technique, where the fluorescence spot from the first laser is sharpened by way of stimulated emission induced by the second laser. However, recent developments have extended the concept to multi-point scanning and to additional photophysical switching mechanisms. This review explains the basic principles behind STED microscopy and the differences with other super-resolution techniques. It provides practical information on how to construct and operate a STED microscope that can be used for nanoscale imaging of GFP and its variants in living brain slices. We conclude by highlighting a series of recent technological innovations that are bound to enhance its scope and performance in the near future. Ó 2015 Published by Elsevier Inc.

1. Theoretical background 1.1. Progress in fluorescence microscopy Fluorescence microscopy has become the dominant method for imaging live biological tissue at the cellular and sub-cellular level. Groundbreaking developments in optical techniques, molecular labeling and image processing continue to increase its performance and scope at a rapid rate. A variety of optical techniques make it ever more feasible to probe and manipulate biological processes inside cells, progressively supplanting traditional biochemical and pharmacological approaches to study neuronal physiology at the molecular level. A wide range of synthetic small molecule fluorescent indicators and genetically encoded biosensors [1–4] has been engineered and, in a parallel development, multi-photon microscopy now allows for imaging of dendrites and spines in the intact brain over many days and even months [5,6]. ⇑ Corresponding author at: Interdisciplinary Institute for Neuroscience, Université de Bordeaux, Bordeaux, France. E-mail address: [email protected] (U.V. Nägerl). 1 Present address: Département des Neurosciences Fondamentales, Université de Genève, Geneva, Switzerland.

However, until recently the diffraction barrier of light microscopy made it impossible to resolve biological structures and events on a submicron spatial scale using visible light, as codified by the famous formula from Ernst Abbe dating back to 1873 [7]:

Dr  k=ð2NAÞ

ð1Þ

where Dr denotes the lateral spatial resolution of the microscope, k the wavelength of light, and NA the numerical aperture of the microscope objective. However, the advent of superresolution microscopy has decisively changed the situation, making it possible to non-invasively image synaptic structures and their molecular dynamics beyond this long-standing limit [8–11]. 1.2. Basic principle of STED microscopy In 1994 Stefan Hell proposed the groundbreaking idea of STED (stimulated emission depletion) for overcoming the diffraction barrier [12]. He envisioned this through the antagonistic action of two laser beams: one laser to excite the fluorophores (to the S1 state) in a small spot just like in regular confocal microscopy, and another one (the STED laser) to immediately de-excite them everywhere to the ground state (S0) except in the very center of

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the spot. The de-excitation is induced by stimulated emission at the long wavelength end of the spontaneous emission spectrum, where it can be readily filtered out from the fluorescence signal (Fig. 1A). The experimental proof-of-concept was delivered in 2000 [13], achieving sub-diffraction resolution along the optical axis, even though nowadays most STED systems use a doughnut-shaped quenching laser to improve the spatial resolution in the focal plane. In STED microscopy the spatial resolution does not only depend on k and NA, but also on the square root of the intensity of the STED laser (I):

. pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi Dr  k 2NA 1 þ I=Is

ð2Þ

where Is is the STED intensity at which half of the fluorescence is quenched, also called the saturation intensity. Accordingly, spatial resolution is theoretically unlimited in STED microscopy. A resolution record of around 6 nm was reported using fluorescent nitrogen vacancies in diamonds [14], an improvement by nearly two orders of magnitude over conventional light microscopy. However, for regular fluorophores and biological samples the achievable resolution typically lies between 30 and 70 nm. STED microscopy quickly drew the attention of neurobiologists because of its potential to image sub-cellular structures in live cells, like dendritic spines and synaptic vesicles, which cannot be properly resolved by conventional light microscopy. Another attractive feature is the fact that STED microscopy naturally shares several key benefits with confocal microscopy, such as depth penetration, optical sectioning and imaging speed, while placing few restrictions on the type of fluorophores that can be used. 1.3. Comparison to other superresolution microscopy techniques Electron microscopy (EM) remains unique in its ability to visualize cellular ultrastructure with nanometric resolution. However, in contrast to fluorescence microscopy, EM requires tissue fixation and hence cannot be used to visualize dynamic processes of living cells. EM also poses serious challenges to multi-target labeling and imaging of large volumes. In addition to STED microscopy, other forms of superresolution microscopy have been developed over the last decade, including photo-activated localization microscopy (PALM)/stochastic optical reconstruction microscopy (STORM) [15–17] and (non-linear) structured illumination microscopy (SIM) [18], all of which are widefield microscopy techniques. PALM/STORM does not break the diffraction barrier by using special optics or illumination patterns. Rather, it forms a super-resolved image from the positions of individual fluorophores, which can be localized with an accuracy of a few nanometers by calculating the center of mass of their positions captured on a sensitive CCD camera. PALM/STORM images represent therefore super-resolved maps of fluorophore positions that adequately sample the overall structure of interest (see [19]). For this pointillist strategy to work, the molecules must be so sparsely activated that their images do not overlap on the same image frame, otherwise the localization cannot be carried out unambiguously. The activation and localization of disparate sets of molecules is repeatedly performed until the map is dense enough. Thousands of image acquisitions are usually required to obtain a single super-resolved image, limiting the temporal resolution and placing considerable demands on image processing and analysis. A

bewildering number of acronyms (in addition to PALM/STORM) refer to various incarnations of the single-molecule based techniques, which essentially only differ in terms of the type of dye molecule and/or photophysical switching mechanism used to turn on or off the fluorescence. PALM/STORM offers nanometric resolution in the x, y, z planes and in multi-colors, and is well suited for imaging assemblies of intracellular or membrane proteins. However, due to its wide-field nature, it remains a challenge to use it in thick tissue samples. In fact, single-molecule detection typically is carried out in total internal reflection fluorescence (TIRF) mode to get a signal-to-noise ratio that is good enough. Compared to STED these stochastic techniques require much lower light intensities, reducing undesirable effects on the sample or the label. In PALM/STORM all emitted photons potentially contribute to the signal and the gain in resolution (the localization accuracy scales with the square root of the number of detected photons). By contrast, in STED the vast majority of S1 to S0 transitions do not contribute to the detected signal because they occur as stimulated emission at a wavelength that is deliberately discarded. This means that in PALM/STORM many photons need to be detected to localize a single molecule, whereas in STED only a low number of photons need to be detected (theoretically one per voxel is enough). The use of pulsed lasers makes STED more expensive and tricky than PALM/STORM, which uses simple CW lasers. Also, the need for overlaying two diffraction-limited PSFs makes STED technically much more challenging. Residual STED light in the center of the doughnut, or the slightest offset between the excitation and STED beams, will degrade contrast and resolution. On the other hand, PALM/STORM requires the detection of single-molecules and heavy image processing, neither of which is trivial nor immune to artifacts. However, it is fair to say that PALM/STORM approaches are in general easier to implement and operate than STED, which explains their widespread adoption by biological labs and the rapid development of commercial solutions. PALM/STORM has been used to visualize the distribution of synaptic receptors and scaffolding proteins at extremely high resolution (20–30 nm in two or three dimensions) in dissociated neurons [20,21] or ultrathin sections of fixed brain tissue [9]. In addition, STORM was used to discover that actin is organized in periodic ring-like structures inside axons [22], which was recently also observed by STED microscopy [23]. In SIM the sample is consecutively excited in wide-field mode with a shifting intensity pattern of high spatial frequency, typically a rotating grid. The resulting fluorescence image is the product of the spatial pattern of the illumination and the sample structure, which effectively shifts higher spatial frequency information of the sample structure into a range that becomes detectable by the bandwidth of the optical system [24]. The principle behind this effect is called heterodyne detection, which is used in many areas of physics. As the pattern of illumination is itself limited by diffraction in regular SIM, the maximal gain in recovering higher spatial frequencies in the sample structure cannot exceed a factor of two – therefore a maximum twofold resolution improvement can be obtained in SIM. Because it is a wide-field technique, SIM allows for relatively fast imaging (around 10 Hz) in multiple colors [25], but provides poor contrast in thick samples. In the non-linear variant of SIM, very strong excitation intensities are used to saturate the fluorescence, which effectively produces a finer grid structure [18]. However, these high light intensities are prohibitive in terms of bleaching and photo-toxicity, also considering that 10–20 different grid positions are needed to assemble a single superresolved image.

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Fig. 1. Basic principle of STED microscopy. In STED microscopy stimulated emission is used to selectively quench the fluorescence. (A) Simplified Jablonski diagram of the energy transitions of GFP during a fluorescence cycle. A blue photon (488 nm) is absorbed; exciting the fluorophore from the ground state (S0) to the first excited singlet state (S1). Spontaneous emission of a green photon (510 nm) occurs, sending the molecule back to the ground state with an average delay defined by the fluorescence lifetime of the fluorophores (2.8 ns for GFP). This transition can be prevented by the process of stimulated emission, where the passage of a photon of longer wavelength returns the fluorophore to the ground state by giving off a photon of exactly the same wavelength and momentum as the incident photon. (B) Spatial restriction of fluorescence emission using a doughnut shaped STED PSF. The quenching of the excited molecules in a doughnut-shaped area effectively reduces the size of the fluorescence spot and increases the spatial resolution. The achievable resolution scales with the intensity of the STED laser beam. (C) Design of a home-built STED microscope for performing two-color superresolution imaging of green and yellow emitting probes in living brain slices. The 595 nm STED wavelength is generated by an optical parametric oscillator (OPO) pumped by a Ti:Sapphire laser emitting at 795 nm (200 fs pulses at 80 MHz). The STED beam passes through a 25 cm glass rod (not shown) and is then coupled into a 20–100 m long single mode fiber (SMF) for stretching the pulses to a couple of hundred picoseconds by dispersion. A helical vortex phase plate introduces the zero intensity in the center of the STED beam and a k/4 is used to create a doughnut-shaped intensity distribution with a deep central minimum at the focal plane. Excitation pulses from a 485 nm laser diode are externally triggered by the Ti:Sapphire laser pulses, and the pulse trains are overlaid in time using an electronic pulse delay. The excitation laser is coupled into a short (2 m) SMF in order to obtain a Gaussian beam profile at the output. The excitation and STED beams are brought to coincide in space after the short-pass dichroic mirror (DM1; cutoff at 590 nm). The laser beams are scanned across the sample using a telecentric scanner for the x and y dimensions and a fast nano-positioner for the z-axis. A brain slice is mounted in a perfusion chamber, and green and yellow fluorophore-labeled structures, respectively, can be imaged simultaneously. The inverted microscope configuration leaves plenty of room for placing pipettes and electrodes from above. The fluorescence signal is detected in a descanned fashion passing through dichroic mirrors; DM1, DM2 (499 nm long pass) and DM3 (514 nm long pass), separating the signal into two detection channels (avalanche photodiode, APD).

RESOLFT (reversible saturable optical fluorescence transitions) refers to the general principle behind STED microscopy, where the diffraction barrier is broken by spatially controlling the ‘on’ and ‘off’ states of photoswitchable fluorescent proteins [26] or organic dyes [27] in the scanning doughnut configuration or in wide-field imaging [28,29]. This technique uses much lower light intensities than STED microscopy, but requires specifically engineered photoswitchable proteins, which can be difficult to express inside cells. Also, the slow switching kinetics and limited brightness of these probes still impose relatively long acquisition times.

1.4. Applications of STED in neurobiology The first biological applications of STED microscopy charted the distributions of synaptic proteins in fixed and immuno-stained cells [30–32]. Subsequently, STED microscopy was used to image the dynamics synaptic vesicles in nerve terminals [33] in dissociated neuronal cultures. Because of its point-scanning nature, STED microscopy lends itself to imaging in thick tissue samples, and has been used to resolve the morphological dynamics of dendritic spines and their

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actin content in living brain slices using YFP as label [34,35]. Recently, STED microscopy was used to reveal nanoscale morphological alterations in dendritic spines in a mouse model of fragile X syndrome (FXS) [36]. Moreover, the STED approach was combined with neurophysiological techniques to relate measurements of nanoscale spine morphology to the diffusive and electrical properties of synapses. This experimental strategy revealed the strong influence that spine morphology exerts on biochemical compartmentalization [37,38]. While changes in dendritic spine heads during activity-dependent synaptic plasticity have been intensely studied using conventional light microscopy [39,40], it was STED microscopy that revealed activity-dependent morphological changes in spine necks, indicating that these nanoscale structures are powerful adjustable determinants of synapse function [38]. In addition to brain slice preparations, STED microscopy has been successfully applied to nanoscale imaging of neuronal structures inside living animals, such as worms [41] and brains of anesthetized mice [42,43]. 2. Instrumentation and hardware requirements In this section, we describe the main hardware requirements for constructing a STED microscope, which are in general more stringent than for conventional confocal or two-photon microscopes. 2.1. Fluorophore considerations The first important step when designing a STED microscope is to decide on the fluorophores to be used, as this will define the wavelengths of the excitation and STED lasers, and consequently the specifications of all optical filters on the setup. Green is probably the most interesting spectral range for many applications in neurobiology, including fluorophores like GFP, YFP and Alexa-488. This is because most labeling tools (viral vectors, transgenic animals, antibodies) have been developed for these popular fluorophores, which in general have superior photophysical properties over red fluorophores [44]. The downside of working in the green range is still the limited availability of suitable pulsed STED laser sources (which should operate in the orange range). 2.2. STED lasers STED microscopy can be performed with continuous wave (CW) [45] or pulsed lasers [46,47]. CW lasers are easier to implement and cheaper. However, pulsed STED lasers are better suited for imaging in live samples, because they require much less average intensity to achieve the same gain in spatial resolution. However, a pulsed laser system requires temporal synchronization and a tunable delay to overlay the excitation and STED pulses arriving at the sample. Two-photon lasers, such as Ti:Sapphire lasers (e.g. MaiTai HP, Spectra-Physics, Darmstadt, Germany or Chameleon Ultra (II), Coherent, Utrecht, The Netherlands; repetition rate 80 MHz, pulse duration: 55% at 520 nm, versus 106 cps). If two-photon excitation is used, with its inherent low out-of-focus excitation, PMTs with large photosensitive areas are the appropriate choice for detecting non-descanned signals from deep within light scattering tissue [51,52]. More recently, hybrid

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photo-detectors (HPD; e.g. Leica HyD, Leica, Mannheim, Germany) have become available, combining desirable features from PMTs and APDs, such as large detection area, low dark noise, high quantum efficiency, high dynamic range and fast time response. 3. Handling and testing a STED microscope In the following section we present practical advice and tips on how to operate a home-built STED microscope, and explain the procedures for aligning it and evaluating its performance. 3.1. Fiber coupling To avoid non-linear (intensity-dependent) effects in the fiber and to protect the pulse-stretching fiber from potentially damaging high peak powers of the femtosecond pulses from the OPO, the beam first passes a 25 cm glass rod, which pre-broadens the pulses to a few picoseconds and substantially lowers peak powers but maintains average power [53]. A fiber optic beam coupler (e.g. APC coupler; Schäfter & Kirchhoff, Hamburg, Germany) is helpful for achieving high and stable coupling efficiency. First, the position of the coupler should be roughly adjusted without any fiber mounted. Next, a short, multi-mode fiber, which can be easily coupled, is mounted and transmission through this is optimized. Finally, the multi-mode fiber is carefully replaced with the long single-mode STED fiber, and coupling is optimized. Generally, a transmission of >50% through the 100 m single-mode fiber is acceptable. A working intensity of 50 mW or less should be used to protect the fiber during coupling, and high powers should be sent through the fiber only when >50% transmission is achieved. Once the fiber is inserted into the coupler, it is not advisable to repeatedly disconnect and connect it, as this may damage the fiber ending, or introduce dust that will compromise coupling. The coupling of the beam into the fiber is very sensitive and may require daily optimization. 3.2. Alignment of beams and coupling into the microscope A critical step for STED imaging is the alignment of the excitation and STED beams along their common path. Rather than trying to align both beams independently from their source to the objective, we recommend ‘backward alignment’, which involves removing the objective and sending a collimated beam backward through the system, making sure it is well centered on all lenses (Fig. 2A). After backward alignment of the excitation, STED and emission paths, the forward beams are brought onto the exact same path by means of pinholes placed during the backward alignment. The pinholes can be left in place permanently for future reference (Fig. 2B). The co-alignment in forward directions can be checked by observing the overlap of the two incoming beams projected at a distance, i.e. observing their overlap on the room ceiling when working with an inverted microscope. On our setups, backward alignment is typically performed on a weekly to monthly basis. While it may initially be a somewhat tricky procedure, with some practice it becomes a manageable routine (30 lm) to get past the dead cells on the surface. However, the performance of oil objectives quickly degrades when the focus is moved more than 10 lm into the tissue because of the mismatch in refractive indices between oil/glass (n = 1.5) and brain tissue (n 1.37). As the mismatch between water (n = 1.33) and brain (n 1.38) is much smaller, water-dipping objectives (60X LUMFI, 1.1 NA, Olympus, Hamburg, Germany) are better-suited for imaging more deeply [50]. Moreover, the long-working distance of this objective leaves plenty of space for electrophysiological recording electrodes on an upright microscope, and the objective is equipped with a correction collar that can minimize spherical aberrations arising from the remaining refractive index mismatch. Glycerol objectives (PL APO, CORR CS, 63, glycerol; Leica, Wetzlar, Germany; NA = 1.3; n 1.45), also equipped with a correction collar, have been used to enhance resolution as deep as 120 lm below tissue surface, in organotypic brain slices [35], acute slices [38], and in vivo [42]. Another interesting option are silicone oil objectives (UPLSAPO60XS, Olympus), which combine a high NA (=1.3) with a refractive index that is very close to that of brain tissue (1.4 versus 1.38). This holds promise for resolution and image brightness, though it remains to be reported how these objectives perform for live tissue STED imaging.

4.2. Multi-color STED A straightforward way to achieve two-color contrast is to use spectrally overlapping fluorophores, such as GFP and YFP, which

can be excited at one wavelength (i.e. 485 nm) and quenched at another (i.e. 595 nm), while still being easily distinguishable on the basis of their emission spectra (Fig. 1C) [47]. The spectral crosstalk of the emission signal can be readily separated using standard linear unmixing algorithms. The single beam pair two-color approach provides inherently aligned excitation PSFs, which reduces problems of chromatic aberration. Also with respect to light exposure and setup complexity the scheme compares favorably to other solutions based on three or four beams to image green and red fluorophores [54,55]. Two-color imaging can also be performed with temporal separation of fluorophores emitting in the same range. The on/off switching behavior of photo-chromic fluorescent proteins can be used to read out their fluorescence sequentially [56]. While this approach gets away with only two laser beams, it does require widefield UV photo-switching and cellular expression of these specially engineered proteins.

4.3. Gated detection The lifetime of the excited state of the fluorophore defines the time window of quenching for the STED light. It is on the order of a few nanoseconds for most fluorophores, including GFP and fluorescein-based dyes. The sample should be doused with STED photons only during the brief period when fluorophores are in the excited state to get maximal quenching and avoid unnecessary light exposure. However, in CW STED the lasers provide a constant stream of photons without any temporal relationship between excitation and the arrival of the STED photons. As a result, quenching efficiency is much lower in a CW than in a pulsed STED system, which can be prohibitive for live-cell imaging applications. On the downside, pulsed lasers are more expensive and tricky to

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implement than simple CW lasers, because they need to be synchronized with the excitation pulses. ‘‘Gated STED’’ is a hybrid solution, which combines pulsed excitation with CW quenching [57]. In this scheme gated detection is used to discard the fluorescence signal right after the excitation pulse, effectively biasing the detected signal towards photons that originate from a smaller spot. This is because very early in time the CW doughnut has not yet produced much quenching (and thus could not yet appreciably reduce the size of the fluorescence spot). Gated detection can be implemented with an APD coupled to a time-correlated single photon counting (TCSPC) module (SPC-730, Becker & Hickl GmbH, Berlin, Germany), making it possible to improve image resolution by filtering out the photons a posteriori based on their arrival times. The downside of gated detection is that it per design discards a fraction of the fluorescence signal, which negatively impacts signal-to-noise ratios and photon budget.

4.4. Two-photon excitation STED microscopy Two-photon microscopy has been the method of choice for imaging in thick tissue preparations and in vivo [5]. The use of wavelengths in the near infrared range (700–1100 nm) improves optical depth penetration because longer wavelengths are scattered much less in biological tissue. However, longer wavelengths also mean larger PSFs, which limits the resolution of two-photon microscopy to 400 nm, unless it is combined with STED microscopy. Two-photon excitation STED microscopy using CW STED lasers was demonstrated in fixed samples [58] and in acute brain slices [51], using red fluorophores and quenching in the near infrared range. Subsequently, two-photon STED (using pulsed two-photon excitation and pulsed single-photon quenching) was developed for imaging GFP [59], and two-photon STED time-lapse imaging of neural structures was demonstrated in acute brain slices [50,52]. Notably, the latter two studies achieved a resolution of 50 nm some 50 lm below tissue surface using long-working distance water objectives, facilitating the combination of STED microscopy with neurophysiological approaches.

4.5. 3D STED It is very important to have high spatial resolution in all three dimensions for imaging complex 3D structures like dendritic spines and astroglial processes. Higher 3D resolution inherently means better optical sectioning and guards against imaging artifacts from z axis projections. For instance, a dendritic spine may look like it has grown or changed shape, whereas in reality it has rotated in and out of the optical axis of the microscope. The standard focal doughnut geometry of the STED beam only improves the lateral spatial resolution, which means that the resolution along the z-axis remains confocal, according to the formula:

Dr z  2k=NA2

ð3Þ

Hence, the PSF of the STED microscope is extremely anisotropic (Drxy  50 nm versus Drz  500 nm). In the confocal case the PSF is also anisotropic, but the aspect ratio is closer to 1:2. To improve the axial resolution, the fluorescence needs to be quenched above and below the focus. This can be achieved by using an annular phase plate, which imposes a delay of p in the central part of the STED laser (corresponding to 50% of the light energy going into the back aperture of the objective). By overlaying this new beam (‘‘bottle beam’’) with the original doughnut beam the fluorescence can be quenched more or less isotropically around

the focal spot, improving the spatial resolution in all three dimensions [60]. 4.6. Adaptive optics In highly scattering biological samples, such as brain tissue, the shape of the STED doughnut gets distorted when imaging at higher depth, effectively degrading image contrast and spatial resolution. Adaptive optics can be used to correct for aberrations induced by the specimen [61], making it possible to maintain doughnut quality deeper inside tissue. Deformable mirrors or liquid crystal-based spatial light modulators (SLMs) can be used for this purpose, where the latter can even be used instead of a phase plate to generate more or less any desired intensity distribution of the STED light in the focal plane. Two recent studies used SLMs to generate and optimize the STED beam for 3D STED imaging in scattering medium [62,63]. Moreover, SLMs were recently used to perform the focal alignment of the excitation and STED beams using an automatic feedback loop [64]. 4.7. Parallelized STED In any point-scanning technique there are direct tradeoffs between imaging speed, signal intensity, field of view and spatial resolution. Because of the small pixel sizes (20 nm) and relatively long dwell times (50 ls) used in STED microscopy, it takes more than ten seconds to acquire a 10 lm  10 lm image, limiting the type of dynamic processes that can be recorded by time-lapse STED imaging. To reconcile high spatial with high temporal resolution, several schemes have been developed, which rely on parallelized image acquisition. The first such design quadrupled the number of excitation and STED focal points, providing accordingly fourfold faster image acquisition over single-point scanning [65]. Abandoning point-detectors, more recent schemes can offer greatly accelerated acquisition of superresolved images using camera-based detection of grid-like illumination patterns featuring 100 [66], 2000 [67] or more than 100,000 [68] doughnut-equivalent intensity minima. However, because these massively parallelized imaging approaches rely on widefield detection schemes, their usefulness for imaging in thick 3D tissue preparations are unfortunately limited. 4.8. STED imaging of nanoscale neural structures in brain slices STED microscopy is well suited for studying the morphological dynamics of neural structures in live and thick tissue preparations. Dendritic spines have been the subject of a growing number of STED publications [35,38,42,43,47,34], but microglial [50] and astrocytic processes [69] and axons (Fig. 4) are equally amenable to live-cell STED microscopy using small organic fluorophores introduced via a patch pipette or transgenic animals expressing variants of GFP in specific cell populations. The use of freely diffusing cytosolic fluorophores as a volume label renders the imaging largely insensitive to bleaching, because of continuous and rapid diffusional replenishment of dye molecules from areas outside of the field of view. The compatibility with different objective types (oil/glycerin/water) provides flexibility for finding a suitable combination of microscope design (upright/inverted), depth penetration and working distance (for placing pipettes), depending on the experimental preparation (organotypic or acute brain slices or in vivo) and other experimental constraints and necessities, essentially without sacrificing spatial resolution.

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Fig. 4. STED imaging in living brain slices. STED-images of (A) YFP labeled dendrite and (B) GFP labeled axon in live organotypic hippocampal slices. (C) Nanoscale dynamics of a dendritic spine (marked by arrow) following a spine specific two-photon glutamate uncaging based long-term potentiation (LTP) protocol (reprinted with permission from Tonnesen et al. [38]). (D) Dual-color live-STED imaging of GFP (green) and YFP-labeled (red) synaptic structures. A single beam pair STED microscope design (described in Fig. 1) was used to image the two fluorophores simultaneously (modified with permission from Tonnesen et al. [47]). (E) Two-color STED imaging can be used to dissect morphological interactions between astrocytes and synapses. The astrocytic processes were volume-labeled with Alexa Fluor 488 (green) via a patch pipette and neurons expressed YFP (red). Higher magnification (right) showing astrocytic processes in close apposition with dendritic spines (modified with permission from Panatier et al. [69]). (F) STED microscopy allows thin, closely associated structured to be discerned. In this zoom-in on the center part of (A), an ‘‘en passant’’ bouton on an axon (left) is contacted by four neighboring spines emerging from the same dendritic segment. Scale bars, 1 lm (A, B, D, F); 500 nm (C).

5. Outlook STED microscopy has come a long way since its conception more than 20 years ago. Having met its share of skeptics and naysayers, it is increasingly embraced by the scientific community.

Still, it is fair to say that STED microscopy is primarily celebrated for its potential for new discoveries, which has yet to be fulfilled. In fact, not many labs have adopted STED technology, largely because it is seen as too challenging and costly to implement for biologists, and because commercial solutions have been limited.

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