A new filtering technique for removing anti‐Stokes emission background in gated CW‐STED microscopy

Stimulated emission depletion (STED) microscopy is a prominent approach of super‐resolution optical microscopy, which allows cellular imaging with so far unprecedented unlimited spatial resolution. The introduction of time‐gated detection in STED microscopy significantly reduces the (instantaneous) intensity required to obtain sub‐diffraction spatial resolution. If the time‐gating is combined with a STED beam operating in continuous wave (CW), a cheap and low labour demand implementation is obtained, the so called gated CW‐STED microscope. However, time‐gating also reduces the fluorescence signal which forms the image. Thereby, background sources such as fluorescence emission excited by the STED laser (anti‐Stokes fluorescence) can reduce the effective resolution of the system. We propose a straightforward method for subtraction of anti‐Stokes background. The method hinges on the uncorrelated nature of the anti‐Stokes emission background with respect to the wanted fluorescence signal. The specific importance of the method towards the combination of two‐photon‐excitation with gated CW‐STED microscopy is demonstrated. (© 2014 WILEY‐VCH Verlag GmbH & Co. KGaA, Weinheim)     

Inside Cover: Enhanced lateral resolution in continuous wave stimulated emission depletion microscopy using tightly focused annular radially polarized excitation beam (J. Biophotonics 9/2019)

STED microscopy is a tool that enables superresolution fluorescence imaging by overcoming the diffraction limitation, and has become more useful in various fields such as biology and material science. STED resolution enhancement can be useful in resolving and visualizing sophisticated details of structures of a sample. For this, the excitation focal spot reduction of CW STED microscopy is achieved by PSF engineering using radial polarization and annular aperture, and improved lateral resolution is obtained by STED effect. This leads to a performance improvement that can lower the depletion beam power required to achieve the same superresolution Further details can be found in the article by Geon Lim, Wan‐Chin Kim, Seunghee Oh, Hyungsuk Lee, No‐Cheol Parket ( e201900060 ).

     

STED Nanoscopy with Time-Gated Detection: Theoretical and Experimental Aspects

In a stimulated emission depletion (STED) microscope the region in which fluorescence markers can emit spontaneously shrinks with continued STED beam action after a singular excitation event. This fact has been recently used to substantially improve the effective spatial resolution in STED nanoscopy using time-gated detection, pulsed excitation and continuous wave (CW) STED beams. We present a theoretical framework and experimental data that characterize the time evolution of the effective point-spread-function of a STED microscope and illustrate the physical basis, the benefits, and the limitations of time-gated detection both for CW and pulsed STED lasers. While gating hardly improves the effective resolution in the all-pulsed modality, in the CW-STED modality gating strongly suppresses low spatial frequencies in the image. Gated CW-STED nanoscopy is in essence limited (only) by the reduction of the signal that is associated with gating. Time-gated detection also reduces/suppresses the influence of local variations of the fluorescence lifetime on STED microscopy resolution.     

Increasing fluorescence lifetime for resolution improvement in stimulated emission depletion nanoscopy

Super‐resolution microscopy (SRM) has had a substantial impact on the biological sciences due to its ability to observe tiny objects less than 200 nm in size. Stimulated emission depletion (STED) microscopy represents a major category of these SRM techniques that can achieve diffraction‐unlimited resolution based on a purely optical modulation of fluorescence behaviors. Here, we investigated how the laser beams affect fluorescence lifetime in both confocal and STED imaging modes. The results showed that with increasing illumination time, the fluorescence lifetime in two kinds of fluorescent microspheres had an obvious change in STED imaging mode, compared with that in confocal imaging mode. As a result, the reduction of saturation intensity induced by the increase of fluorescence lifetime can improve the STED imaging resolution at the same depletion power. The phenomenon was also observed in Star635P‐labeled human Nup153 in fixed HeLa cells, which can be treated as a reference for the synthesis of fluorescent labels with the sensitivity to the surrounding environment for resolution improvement in STED nanoscopy.      

Dual‐color STED super‐resolution microscope using a single laser source

STED (stimulated emission depletion) microscopy is one of the most promising super‐resolution fluorescence microscopies，due to its fast imaging and ultra‐high resolution. In this paper, we present a dual‐color STED microscope with a single laser source. Polarization beam splitters are used to separate the output from a supercontinuum laser source into four laser beams, including two excitation beams (488, 635 nm) and two depletion beams (592, 775 nm). These four laser beams are then used to build a low cost dual‐color STED system to achieve a spatial resolution of 75 nm in cell samples.     

Saturated two‐photon excitation fluorescence microscopy for the visualization of cerebral neural networks at millimeters deep depth

Optical imaging is a key modality for observing biological specimen with higher spatial resolution. However, scattering and absorption of light in tissues are inherent barriers in maximizing imaging depth in biological tissues. To achieve this goal, use of light at near‐infrared spectrum can improve the present situation. Here, the capability of saturated two‐photon saturated excitation (TP‐SAX) fluorescence microscopy to image at depths of >2.0 mm, with submicron resolution in transparent mouse brain imaging, is demonstrated. At such depths with scattering‐enlarged point spread function (PSF), we find that TP‐SAX is capable to provide spatial resolution improvement compared to its corresponding TPFM, which is on the other hand already providing a much improved resolution compared with single‐photon confocal fluorescence microscopy. With the capability to further improve spatial resolution at such deep depth with scattering‐enlarged PSF, TP‐SAX can be used for exquisite visualization of delicate cerebral neural structure in the scattering regime with a submicron spatial resolution inside intact mouse brain.      

Low‐photobleaching line‐scanning confocal microscopy using dual inclined beams

Confocal microscopy is an indispensable tool for biological imaging due to its high resolution and optical sectioning capability. However, its slow imaging speed and severe photobleaching have largely prevented further applications. Here, we present dual inclined beam line‐scanning (LS) confocal microscopy. The reduced excitation intensity of our imaging method enabled a 2‐fold longer observation time of fluorescence compared to traditional LS microscopy while maintaining a good sectioning capability and single‐molecule sensitivity. We characterized the performance of our method and applied it to subcellular imaging and three‐dimensional single‐molecule RNA imaging in mammalian cells.      

Upgrade of a Scanning Confocal Microscope to a Single-Beam Path STED Microscope

By overcoming the diffraction limit in light microscopy, super-resolution techniques, such as stimulated emission depletion (STED) microscopy, are experiencing an increasing impact on life sciences. High costs and technically demanding setups, however, may still hinder a wider distribution of this innovation in biomedical research laboratories. As far-field microscopy is the most widely employed microscopy modality in the life sciences, upgrading already existing systems seems to be an attractive option for achieving diffraction-unlimited fluorescence microscopy in a cost-effective manner. Here, we demonstrate the successful upgrade of a commercial time-resolved confocal fluorescence microscope to an easy-to-align STED microscope in the single-beam path layout, previously proposed as “easy-STED”, achieving lateral resolution < λ/10 corresponding to a five-fold improvement over a confocal modality. For this purpose, both the excitation and depletion laser beams pass through a commercially available segmented phase plate that creates the STED-doughnut light distribution in the focal plane, while leaving the excitation beam unaltered when implemented into the joint beam path. Diffraction-unlimited imaging of 20 nm-sized fluorescent beads as reference were achieved with the wavelength combination of 635 nm excitation and 766 nm depletion. To evaluate the STED performance in biological systems, we compared the popular phalloidin-coupled fluorescent dyes Atto647N and Abberior STAR635 by labeling F-actin filaments in vitro as well as through immunofluorescence recordings of microtubules in a complex epithelial tissue. Here, we applied a recently proposed deconvolution approach and showed that images obtained from time-gated pulsed STED microscopy may benefit concerning the signal-to-background ratio, from the joint deconvolution of sub-images with different spatial information which were extracted from offline time gating.     

Live-Cell Superresolution Imaging by Pulsed STED Two-Photon Excitation Microscopy

Two-photon laser scanning microscopy (2PLSM) allows fluorescence imaging in thick biological samples where absorption and scattering typically degrade resolution and signal collection of one-photon imaging approaches. The spatial resolution of conventional 2PLSM is limited by diffraction, and the near-infrared wavelengths used for excitation in 2PLSM preclude the accurate imaging of many small subcellular compartments of neurons. Stimulated emission depletion (STED) microscopy is a superresolution imaging modality that overcomes the resolution limit imposed by diffraction and allows fluorescence imaging of nanoscale features. Here, we describe the design and operation of a superresolution two-photon microscope using pulsed excitation and STED lasers. We examine the depth dependence of STED imaging in acute tissue slices and find enhancement of 2P resolution ranging from approximately fivefold at 20 μm to approximately twofold at 90-μm deep. The depth dependence of resolution is found to be consistent with the depth dependence of depletion efficiency, suggesting resolution is limited by STED laser propagation through turbid tissue. Finally, we achieve live imaging of dendritic spines with 60-nm resolution and demonstrate that our technique allows accurate quantification of neuronal morphology up to 30-μm deep in living brain tissue.     

Coaligned Dual-Channel STED Nanoscopy and Molecular Diffusion Analysis at 20 nm Resolution

We report on a fiber laser-based stimulated emission-depletion microscope providing down to ∼20 nm resolution in raw data images as well as 15–19 nm diameter probing areas in fluorescence correlation spectroscopy. Stimulated emission depletion pulses of nanosecond duration and 775 nm wavelength are used to silence two fluorophores simultaneously, ensuring offset-free colocalization analysis. The versatility of this superresolution method is exemplified by revealing the octameric arrangement of Xenopus nuclear pore complexes and by quantifying the diffusion of labeled lipid molecules in artificial and living cell membranes.Since its first demonstration in (live) cell imaging (1), stimulated emission depletion (STED) fluorescence microscopy has been realized in many variants. Particularly, the key phenomenon employed in this method, namely switching fluorophores transiently off by stimulated emission, has been accomplished with laser pulses varying from picoseconds to nanoseconds in duration, and from kHz to MHz in repetition rate. Because continuous-wave beams are suitable as well (2), STED microscopy has been implemented with rather different laser systems, ranging from model-locked femtosecond to continuous-wave laser diodes (3,4). Although it underscores the versatility of STED to modulate the fluorescence capability of a fluorophore, this wide range of options may confuse adopters when balancing simplicity, applicability, and resolution gain. The situation is exacerbated when implementing pairs of excitation and STED beams for dual-color colocalization studies (5,6).Here we report on a simple arrangement providing dual-color STED nanoscopy (Fig. 1) and molecular diffusion quantification down to ∼20 nm in (living) cells. The presented dual-channel STED microscope utilizes a single fiber laser providing a 20-MHz train of 775 nm wavelength pulses of 1.2-ns duration. This compact laser source enables STED on fluorophores emitting in the orange to red range. Specifically, we applied this laser on the orange dyes Atto590 and Atto594 (excitation: 595 nm; detection: 620 ± 20 nm), and the red dyes KK114 and Abberior Star635P (excitation: 640 nm; detection: 670 ± 20 nm). Although the spectra of the dyes are partially overlapping, the individual color channels can be separated without data processing (see Fig. S1 and Fig. S2 in the Supporting Material). Both channels are recorded simultaneously within 50 ns, using temporally interleaved pulsed excitation in combination with time-gated detection (5,7,8).Open in a separate windowFigure 1Fluorescence nanoscopy of protein complexes with a compact near-infrared nanosecond-pulsed STED microscope. (A) STED reveals immunolabeled subunits in amphibian NPC; raw data smoothed with a Gaussian filter extending over 14 nm in FWHM. The diameter of the octameric gp210 ring is established as ∼160 nm. Scale bar, 500 nm. (B) Individual NPC image showing eight antibody-labeled gp210 homodimers as 20–40 nm sized units and a 80 nm-sized localization of the subunits in the central channel.Because in STED microscopy, the STED doughnuts firmly determine the position of the fluorescently active molecules, the use of a single doughnut for both fluorophores guarantees that the two color channels are almost perfectly coaligned. The use of the doughnut even counteracts misalignments of the confocal excitation and detection channels (Fig. 2, and see Fig. S3), making STED microscopy particularly powerful for colocalization measurements.Open in a separate windowFigure 2Determination of the colocalization accuracy. Xenopus A6 cells, labeled with an antiserum against multiple NUP subunits in the central NPC channel and two secondary antibodies decorated with the fluorophores Abberior STAR635P and Atto594 were imaged by STED microscopy. (A) Upon overlaying both channels, a high degree of colocalization is directly visible. Scale bar, 200 nm. (B) Quantification of the colocalization by cross correlation of much larger images (see Fig. S3). The correlation is maximal for zero displacement of the images, proving colocalization. (C) Confocal image of monocolored fluorescent beads taken with improperly coaligned excitation beams (left). Improper coalignment spoils the colocalization accuracy in confocal imaging; the two channels should be perfectly coaligned, but they show a false offset as indicated by the color difference. The offset is quantified by the cross correlation of the two channels (right). (D) The STED image of the same beads (left) not only shows 10-fold improved resolution over the confocal image in panel C, but also improved colocalization, again quantified by cross correlation (right). Thus, by predetermining the position of emission, the STED doughnut counteracts errors induced by imperfect coalignment of the two confocal color channels (for details, see Fig. S3). Scale bars = 100 nm.The cross section for stimulated emission is lower at 775 nm as compared to that found at somewhat shorter wavelengths (5), yet STED pulse energies of ∼7 nJ in the focus are sufficient to yield a resolution of ∼30 nm and ∼20 nm in the orange and red channels, respectively (see Fig. S4). In addition, due to the lower peak intensity, the 1.2 ns pulses are likely to induce less nonlinear absorption and hence less photostress as compared to their more commonly used <0.2 ns counterparts (8,9). On the other hand, the pulses are only 2–4 times shorter than the typical lifetime of the excited state, which lessens their STED efficiency. This slight reduction is neutralized here by detecting photons emitted ∼1 ns after excitation (5,7,8).The potential of this straightforward implementation of STED microscopy is evident when imaging immunolabeled nuclear pore complexes (NPCs) of cultured Xenopus cells. Contrary to the confocal recording, STED microscopy reveals subunits of this protein complex, specifically the typical eightfold symmetry of its peripheral transmembrane protein gp210, along with a set of proteins in the central pore channel (Fig. 1, and see Fig. S5 and Fig. S6). Unlike in stochastic superresolution imaging of gp210 (10), the color channels are inherently coaligned and simultaneously recorded simply by executing a single scan. Apart from a weak smoothing and background subtraction applied to enhance image contrast, the images are raw.Because fluorescence off-switching by STED is an instant process, STED microscopy can be employed to study fast spatial translocations, such as the diffusion of molecules on the nanoscale (3). To benchmark the performance of our setup, we analyzed the diffusion of a fluorescent glycerophospholipid analog (11) by fluorescence correlation spectroscopy (FCS) in membranes of living mammalian PtK2-cells (Fig. 3). STED allowed us to reduce the diameter of the probed area from the 250 nm-sized diffraction limit down to 19 nm (FWHM), representing σ = 8 nm in standard deviation of a Gaussian fit. The attained subdiffraction area is 2.5 times smaller as compared to what has been reported in living cells to date (4). In model membranes, the smallest diameter was 15 nm (σ = 6.4 nm).Open in a separate windowFigure 3Nanoscale molecular diffusion analyzed by STED FCS. (A) For moderate and larger STED beam power P_STED, the resolution scales inversely with its square-root, attaining 15 nm in FWHM of the distribution of fluorescence emission in space, describing the measurement area. Note the relatively small threshold power P_S = 1.4 mW, which implies that a large resolution gain is already attained for P_STED < 100 mW. (Inset) The resolution was determined by measuring the transit time of a fluorescent phospholipid-analog (DSPE-PEG-KK114) in a lipid model membrane through the detection area by FCS. (B) In living mammalian Ptk2-cells, the transit time of the lipid analog scales linearly with the detection area, revealing a diffusion constant D_lat = 0.33 μm²/s, and showing that this lipid analog diffuses largely freely in the plasma membrane down to <20 nm scales.In both measurements, the molecular transit time depends linearly on the probed area, indicating that the labeled lipid molecules diffuse essentially freely down to spatial scales of 20 nm. Accordingly, the anomaly exponent α was close to 1 with values of α > 0.85, showing only minor deviations from free diffusion (see Fig. S7). Because the diameter is inversely proportional to the square-root of the STED beam power, the resolution can be adapted to a particular application need (Fig. 3, A and B).In summary, our arrangement provides up-to-date STED microscopy resolution in offset-free colocalization recordings. The ready-to-use near-infrared laser pulses keep undesired single and multiphoton absorption low and leave the visible spectrum amenable for further studies.     

Low-power compact continuous-wave stimulated emission depletion microscopy

Stimulated emission depletion (STED) microscopy can break the optical diffraction barrier and provide subdiffraction resolution. According to the STED superresolution imaging principle, the resolution of STED is positively related to the power of the depletion laser. However, high-laser power largely limits the study of living cells or living bodies. Moreover, the high complexity and high cost of conventional pulsed STED microscopy limit the application of this technique. Therefore, this paper describes a simple continuous-wave STED (CW-STED) system constructed on a 45 × 60 cm breadboard and combined with digitally enhanced (DE) technology; low-power superresolution imaging is realized, which has the advantages of reducing system complexity and cost. The low-system complexity, low cost, and low-power superresolution imaging features of CW-STED have great potential to advance the application of STED microscopy in biological research.     

STED microscopy with continuous wave beams

We report stimulated emission depletion (STED) fluorescence microscopy with continuous wave (CW) laser beams. Lateral fluorescence confinement from the scanning focal spot delivered a resolution of 29-60 nm in the focal plane, corresponding to a 5-8-fold improvement over the diffraction barrier. Axial spot confinement increased the axial resolution by 3.5-fold. We observed three-dimensional (3D) subdiffraction resolution in 3D image stacks. Viable for fluorophores with low triplet yield, the use of CW light sources greatly simplifies the implementation of this concept of far-field fluorescence nanoscopy.     

Axial resolution enhancement of light‐sheet microscopy by double scanning of Bessel beam and its complementary beam

The side lobes of Bessel beam will create significant out‐of‐focus background when scanned in light‐sheet fluorescence microscopy (LSFM), limiting the axial resolution of the imaging system. Here, we propose to overcome this issue by scanning the sample twice with zeroth‐order Bessel beam and another type of propagation‐invariant beam, complementary to the zeroth‐order Bessel beam, which greatly reduces the out‐of‐focus background created in the first scan. The axial resolution can be improved from 1.68 μm of the Bessel light‐sheet to 1.07 μm by subtraction of the two scanned images across a whole field‐of‐view of up to 300 μm × 200 μm × 200 μm. The optimization procedure to create the complementary beam is described in detail and it is experimentally generated with a spatial light modulator. The imaging performance is validated experimentally with fluorescent beads as well as eGFP‐labeled mouse brain neurons.      

Nanoscopy of Filamentous Actin in Cortical Dendrites of a Living Mouse

We demonstrate superresolution fluorescence microscopy (nanoscopy) of protein distributions in a mammalian brain in vivo. Stimulated emission depletion microscopy reveals the morphology of the filamentous actin in dendritic spines down to 40 μm in the molecular layer of the visual cortex of an anesthetized mouse. Consecutive recordings at 43–70 nm resolution reveal dynamical changes in spine morphology.The postsynaptic part of most excitatory synapses in the brain is formed by dendritic spines, which are small protrusions along the dendrites that are highly dynamic during development, but also undergo morphological changes in adulthood (1,2). A prime candidate for regulating these dynamics is the neuronal actin network (3). Filamentous (F-) actin is also important for anchoring postsynaptic receptors and modulating synaptic activities, e.g., through the organization of the postsynaptic density (3). Clearly, the actin dynamics of dendritic spines is best studied in vivo, e.g., in a living mouse, and with confocal and multiphoton microscopy because these techniques can provide three-dimensional optical sectioning several 100 μm inside brain tissue (4). However, because necks of dendritic spines are on the 50–150-nm scale, their details are beyond the 250–400-nm resolution afforded by these diffraction-limited techniques. Fortunately, the diffraction resolution barrier of lens-based fluorescence microscopy has recently been overcome by causing the fluorophores of nearby features to emit sequentially (5). One of the techniques relying on this principle, stimulated emission depletion (STED) microscopy, has recently resolved dendritic spines in the cortex of a living mouse (6). In that initial, in vivo superresolution study, the dendrites were only volume-labeled, and consequently, the spatial arrangements of specific cytoskeletal proteins could not be imaged. On the other hand, F-actin has actually been imaged in living brain slices (7), but in vivo imaging of these structures has not yet been attained.Compared to other superresolution or nanoscopy techniques, STED microscopy bears a number of advantages for imaging spines in the living brain. Implemented as a beam scanning confocal microscope, STED nanoscopy offers optical sectioning and measurements at greater depth. In addition, motion artifacts of the dynamic structures can be minimized by fast scanning. And last but not least, STED can be performed with standard fluorescent proteins. Therefore, we here apply STED nanoscopy to noninvasively uncover the actin cytoskeleton in the living mouse brain. In particular, we show that the 43–70-nm resolution obtained by STED visualizes rearrangements of the dendritic spines in vivo.We took on the challenge of labeling the actin cytoskeleton in the living mouse cortex. We utilized Lifeact-EYFP, a fusion protein consisting of a small peptide and the yellow fluorescent protein EYFP, which directly binds to F-actin without disturbing its polymerization (8). The labeling itself was accomplished by viral infection. To this end, adeno-associated viral particles (AAV) of serotype 2, facilitated by the neuron specific human synapsin promoter hSYN (9) and Semliki Forest viruses (SFV), were created to express Lifeact-EYFP in neurons. For virus injection, the mouse was anesthetized and the head was fixed in a model No. SG-4N head holder (Narishige International USA, East Meadow, NY). A 5-mm incision of the skin of the head enabled drilling a 0.5-mm-diameter hole into the skull. The hole was positioned 0.5 mm outside the prospective imaging center in the visual cortex. The AAVs were injected with a micropipette connected to a pressure generator (Tooheyspritzer; Toohey Company, Fairfield, NJ). Thus, we were able to inject ∼750 nL of concentrated AAV at an angle of 30° over a time of ∼5 min to the layer of pyramidal cells in the prospective imaging center. After injection and 5-min pause, the pipette was retracted with a 5-min break at the half-way point to allow the virus to diffuse into the tissue. The skin was closed with three stitches and the mouse kept on a heating plate in an anesthetic recovery box until wake-up.After 10 days the mouse was prepared for in vivo STED nanoscopy, according to Berning et al. (6) (see also the Supporting Material). At this point, the skin had completely healed and the mouse showed no sign of obvious behavioral abnormality. Optical access was provided by a glass-sealed hole of ∼2 mm in diameter, exposing the visual cortex (Fig. 1 a). STED nanoscopy was performed with an upright beam-scanning microscope similar to that described by Berning et al. (6), with short optical paths and good vibration-damping (Fig. 1 b and see the Supporting Material). The coaligned excitation and STED beams were focused onto the mouse brain using a 1.3 numerical-aperture glycerol immersion lens. The correction collar of the lens allowed compensation of spherical aberrations arising from focusing beneath the brain surface (7).Open in a separate windowFigure 1STED nanoscopy of the dendritic filamentous (F-) actin cytoskeleton in the visual cortex of a living mouse. (a) Clear view of the visual cortex through an optical window. (b) Upright STED imaging of the anesthetized mouse. (c) Dendritic F-actin in the molecular layer of the visual cortex at 4, 25, and 40-μm depths. Maximum intensity projection of a stack of five (xy) images taken in 500-nm axial (z) distances. (Right) Line profile at the marked positions; average of five lines of the raw data and Lorentz fit with full width at half-maximum (FWHM); all image data are raw.Fig. 1 c shows representative parts of dendrites in the molecular layer of the visual cortex. The combination of Lifeact-EYFP labeling and superresolution displayed the dendritic actin of the living mouse neuron in unprecedented detail. Most spines have an actin-rich bulbous end, i.e., a spine head. Sometimes, the dendrite shows small areas with high actin enrichment, which presumably constitute the beginning of filopodia outgrowths (see Fig. S1 in the Supporting Material). The STED image quality was maintained down to a depth of 40 μm below the cover glass. The actin filaments in the spine neck were 43–70-nm thin (see Fig. S2), which can also be interpreted as an upper estimate (poorest value) for the resolution obtained by STED. Note that the images were not processed after recording. All dendrites appeared normal, i.e., in comparison with the morphology of volume-labeled pyramidal cells of transgenic mice (6). The STED beam average laser power was 34 mW. For somewhat greater laser power, we occasionally saw swelling of the dendrites but they were never destroyed. The maximum applicable power depends on the thickness of the dendrite and most likely on the presence of mitochondria as well.Next, we raised the expression level of Lifeact-EYFP by replacing AAV with SFV infection (7,10). We injected 750 nL of SFV (see the Supporting Material) analog to the AAV protocol and allowed the mouse to wake up and recover. After one day, we recorded in vivo STED nanoscopy images of the visual cortex. The labeling was sparser than with the AAV, i.e., fewer cells expressed Lifeact-EYFP, but the signal was brighter and highly specific to neurons. Fig. 2 shows a STED image of a part of a dendrite in the visual cortex at depth <10 μm. The actin label is brighter in the spine heads than in the body of the dendrite, showing that Lifeact-EYFP is primarily attached to F-actin. STED recording over 12 min revealed morphological changes in the actin cytoskeleton. No changes were observed after fixation (see Fig. S4). Bleaching-corrected brightness changes in the spine head inherently reflect density changes in the actin network. In contrast to AAV, SFV shuts down host cell protein synthesis, which leads to cell death after >24 h (11,12); this was not improved by the less cytotoxic SFV(PD) variant (11). Therefore, we recorded in vivo nanoscopy images one day after viral transduction where most dendrites looked healthy. To confirm the viral transduction and verify the subtype of the infected neurons, we perfused the mouse with paraformaldehyde and imaged the brain slices of the region of interest (see Fig. S5). Whereas the AAV labeled mainly neurons of the pyramidal layer, the SFV infected sparsely neurons from all layers of the cortex.Open in a separate windowFigure 2Actin rearrangement in dendritic spines at 60-nm subdiffraction spatial resolution. Image stacks reveal dynamic changes of actin in the spines. (Arrows) Shape changes of spine heads. Maximum intensity projection of five slices of 500-nm axial (z) separation; all data are raw. Average power at back-aperture of objective lens: 2.4 μW excitation and 38-mW STED.The 4–5-fold lateral resolution improvement of STED over standard confocal and multiphoton microscopy is not sufficient to resolve single actin fibers, as with platinum replica electron microscopy (13). Future refinements of both labeling and STED imaging should make this goal achievable. The resolution along the optical (z) axis was kept diffraction-limited (∼500 nm) so that the total illumination dose remained small. At depth >40 μm, scattering and aberrations compromise the image quality. Nonetheless, the molecular layer of the sensory cortex is a highly interesting target for functional optical nanoscopy, because it is the site of the first stage of cortical sensory processing.In summary, STED microscopy can be applied to study subcellular protein structures at 43–70-nm resolution down to 40 μm in the brain of a living mammal. Specifically, we showed that the dynamic actin network responsible for the morphologic plasticity in the brain can be superresolved in the living mouse. Extending in vivo STED microscopy to other protein assemblies as well as to other cell types should provide basic insights into the working principles of the brain.     

Bessel‐beam illumination in dual‐axis confocal microscopy mitigates resolution degradation caused by refractive heterogeneities

One of the main challenges for laser‐scanning microscopy of biological tissues with refractive heterogeneities is the degradation in spatial resolution that occurs as a result of beam steering and distortion. This challenge is particularly significant for dual‐axis confocal (DAC) microscopy, which achieves improved spatial‐filtering and optical‐sectioning performance over traditional confocal microscopy through off‐axis illumination and collection of light with low‐numerical aperture (NA) beams that must intersect precisely at their foci within tissues. DAC microscope image quality is sensitive to positional changes and distortions of these illumination‐ and collection‐beam foci. Previous studies have shown that Bessel beams display improved positional stability and beam quality than Gaussian beams when propagating through tissues with refractive heterogeneities, which suggests that Bessel‐beam illumination may enhance DAC microscopy of such tissues. Here, we utilize both Gaussian and Bessel illumination in a point‐scanned DAC microscope and quantify the resultant degradation in resolution when imaging within heterogeneous optical phantoms and fresh tissues. Results indicate that DAC microscopy with Bessel illumination exhibits reduced resolution degradation from microscopic tissue heterogeneities compared to DAC microscopy with conventional Gaussian illumination.

     

Inside Front Cover: Increasing fluorescence lifetime for resolution improvement in stimulated emission depletion nanoscopy (J. Biophotonics 5/2019)

A STED‐FLIM system is developed to observe the changes of fluorescence lifetime. The pictures show increased lifetime of fluorescent microspheres samples with laser illumination time in both confocal and STED imaging modes. Due to the saturation power of fluorophores is correlated with fluorescence lifetime, the lifetime increase is beneficial for the reduction of the saturation power, indicating the same imaging resolution can be achieved in a lower depletion power. Further details can be found in the article by Lu‐Wei Wang, Yue Chen, Wei Yan, et al. ( e201800315 ).

     

Visualization of the Immunological Synapse by Dual Color Time-gated Stimulated Emission Depletion (STED) Nanoscopy

Natural killer cells form tightly regulated, finely tuned immunological synapses (IS) in order to lyse virally infected or tumorigenic cells. Dynamic actin reorganization is critical to the function of NK cells and the formation of the IS. Imaging of F-actin at the synapse has traditionally utilized confocal microscopy, however the diffraction limit of light restricts resolution of fluorescence microscopy, including confocal, to approximately 200 nm. Recent advances in imaging technology have enabled the development of subdiffraction limited super-resolution imaging. In order to visualize F-actin architecture at the IS we recapitulate the NK cell cytotoxic synapse by adhering NK cells to activating receptor on glass. We then image proteins of interest using two-color stimulated emission depletion microscopy (STED). This results in <80 nm resolution at the synapse. Herein we describe the steps of sample preparation and the acquisition of images using dual color STED nanoscopy to visualize F-actin at the NK IS. We also illustrate optimization of sample acquisition using Leica SP8 software and time-gated STED. Finally, we utilize Huygens software for post-processing deconvolution of images.     

Oblique scanning 2‐photon light‐sheet fluorescence microscopy for rapid volumetric imaging

Light‐sheet fluorescence microscopy (LSFM) is a powerful tool for biological studies because it allows for optical sectioning of dynamic samples with superior temporal resolution. However, LSFM using 2 orthogonally co‐aligned objectives requires a special sample geometry, and volumetric imaging speed is limited due to physical sample translation. This paper describes an oblique scanning 2‐photon LSFM (OS‐2P‐LSFM) that eliminates these limitations by using a single objective near the sample and a refractive scanning‐descanning system. This system also provides improved light‐sheet confinement against scattering by using a 2‐photon Bessel beam. The OS‐2P‐LSFM hold promise for studying structural, functional and dynamic aspects of living tissues and organisms because it allows for high‐speed, translation‐free and scattering‐robust 3D imaging of large biological specimens.      

X10 expansion microscopy enables 25‐nm resolution on conventional microscopes

Expansion microscopy is a recently introduced imaging technique that achieves super‐resolution through physically expanding the specimen by ~4×, after embedding into a swellable gel. The resolution attained is, correspondingly, approximately fourfold better than the diffraction limit, or ~70 nm. This is a major improvement over conventional microscopy, but still lags behind modern STED or STORM setups, whose resolution can reach 20–30 nm. We addressed this issue here by introducing an improved gel recipe that enables an expansion factor of ~10× in each dimension, which corresponds to an expansion of the sample volume by more than 1,000‐fold. Our protocol, which we termed X10 microscopy, achieves a resolution of 25–30 nm on conventional epifluorescence microscopes. X10 provides multi‐color images similar or even superior to those produced with more challenging methods, such as STED, STORM, and iterative expansion microscopy (iExM). X10 is therefore the cheapest and easiest option for high‐quality super‐resolution imaging currently available. X10 should be usable in any laboratory, irrespective of the machinery owned or of the technical knowledge.     

Self‐referenced axial chromatic dispersion measurement in multiphoton microscopy through 2‐color third‐harmonic generation imaging

With tunable excitation light, multiphoton microscopy is widely used for imaging biological structures at subcellular resolution. Axial chromatic dispersion, present in virtually every transmissive optical system including the multiphoton microscope, leads to focal (and the resultant image) plane separation. Here, we experimentally demonstrate a technique to measure the axial chromatic dispersion in a multiphoton microscope, using simultaneous 2‐color third‐harmonic generation imaging excited by a 2‐color soliton source with tunable wavelength separation. Our technique is self‐referenced, eliminating potential measurement error when 1‐color tunable excitation light is used which necessitates reciprocating motion of the mechanical translation stage. Using this technique, we demonstrate measured axial chromatic dispersion with 2 different objective lenses in a multiphoton microscope. Further measurement in a biological sample also indicates that this axial chromatic dispersion, in combination with 2‐color imaging, may open up opportunity for simultaneous imaging of 2 different axial planes.