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Background The development of nanoscale secondary ion mass spectrometry (NanoSIMS) has revolutionized the study of biological tissues by enabling, e.g., the visualization and quantification of metabolic processes at subcellular length scales. However, the associated sample preparation methods all result in some degree of tissue morphology distortion and loss of soluble compounds. To overcome these limitations an entirely cryogenic sample preparation and imaging workflow is required. Results Here, we report the development of a CryoNanoSIMS instrument that can perform isotope imaging of both positive and negative secondary ions from flat block-face surfaces of vitrified biological tissues with a mass- and image resolution comparable to that of a conventional NanoSIMS. This capability is illustrated with nitrogen isotope as well as trace element mapping of freshwater hydrozoan Green Hydra tissue following uptake of 15 N-enriched ammonium. Conclusion With a cryo-workflow that includes vitrification by high pressure freezing, cryo-planing of the sample surface, and cryo-SEM imaging, the CryoNanoSIMS enables correlative ultrastructure and isotopic or elemental imaging of biological tissues in their most pristine post-mortem state. This opens new horizons in the study of fundamental processes at the tissue- and (sub)cellular level. Teaser CryoNanoSIMS: subcellular mapping of chemical and isotopic compositions of biological tissues in their most pristine post-mortem state.

Structural analyses of organelles and localization of proteins in their confines are essential to investigate inner workings of eukaryotic cells. Electron tomography (ET) is a three-dimensional electron microscopy method with which we can extract sliced views of organelles from any direction and quantify their structural parameters at nanometer-level resolution. This advanced electron microscopy tool is suited for characterization of convoluted membrane compartments and of cellular constituents of dimensions smaller than 100 nm. ET studies of plant cells fixed by rapid freezing have expanded our understanding of the biogenesis and functions of plant organelles. Here we describe how the molecular imaging capacity of correlative light and electron microscopy can be integrated with ET in studies of plant organelles.

Since the beginning of the 1980s, cryo‐electron microscopy of a thin film of vitrified aqueous suspension has made it possible to observe biological particles in their native state, in the absence of the usual artefacts of dehydration and staining. Combined with 3‐d reconstruction, it has become an important tool for structural molecular biology. Larger objects such as cells and tissues cannot generally be squeezed in a thin enough film. Cryo‐electron microscopy of vitreous sections (CEMOVIS) provides then a solution. It requires vitrification of a sizable piece of biological material and cutting it into ultrathin sections, which are observed in the vitrified state. Each of these operations raises serious difficulties that have now been overcome. In general, the native state seen with CEMOVIS is very different from what has been seen before and it is seen in more detail. CEMOVIS will give its full potential when combined with computerized electron tomography for 3‐d reconstruction.

The performance of the photosynthesis machinery in plants, including light harvesting, electron transport, and protein repair, is controlled by structural changes in the thylakoid membrane system inside the chloroplasts. In particular, the structure of the stacked grana area of thylakoid membranes is highly dynamic, changing in response to different environmental cues such as light intensity. For example, the aqueous thylakoid lumen enclosed by thylakoid membranes in grana has been documented to swell in the presence of light. However, light‐induced alteration of the stromal gap in the stacked grana (partition gap) and of the unstacked stroma lamellae has not been well characterized. Light‐induced changes in the entire thylakoid membrane system, including the lumen in both stacked and unstacked domains as well as the partition gap, are presented here, and the functional implications are discussed. This structural analysis was made possible by development of a robust semi‐automated image analysis method combined with optimized plant tissue fixation techniques for transmission electron microscopy generating quantitative structural results for the analysis of thylakoid ultrastructure. Significance Statement A methodical pipeline ranging from optimized leaf tissue preparation for electron microscopy to quantitative image analysis was established. This methodical development was employed to study details of light‐induced changes in the plant thylakoid ultrastructure. It was found that the lumen of the entire thylakoid system (stacked and unstacked domains) undergoes light‐induced swelling, whereas adjacent membranes on the stroma side in stacked grana thylakoid approach each other.

• We examined the subcellular cadmium (Cd) localization in roots and leaves of wild-type Arabidopsis thaliana (ecotype Columbia) exposed to environmentally relevant Cd concentrations. • Energy-dispersive X-ray microanalysis (EDXMA) was performed on high-pressure frozen and freeze-substituted tissues. • In the root cortex, Cd was associated with phosphorus (Cd/P) in the apoplast and sulfur (Cd/S) in the symplast, suggesting phosphate and phytochelatin sequestration, respectively. In the endodermis, sequestration of Cd/S was present as fine granular deposits in the vacuole and as large granular deposits in the cytoplasm. In the central cylinder, symplastic accumulation followed a distinct pattern illustrating the importance of passage cells for the uptake of Cd. In the apoplast, a shift of Cd/S granular deposits from the middle lamella towards the plasmalemma was observed. Large amounts of precipitated Cd in the phloem suggest retranslocation from the shoot. In leaves, Cd was detected in tracheids but not in the mesophyll tissue. • Extensive symplastic and apoplastic sequestration in the root parenchyma combined with retranslocation via the phloem confirms the excluder strategy of Arabidopsis thaliana.

Temporary accumulation of glycogen in the epithelial cells of the developing mouse submandibular gland was examined under light microscopic histochemistry and electron microscopy. To avoid loss of water-soluble glycogen during histological tissue preparation, fixation with ethanol and embedding in hydrophilic glycol methacrylate resin was used for light microscopy, and high-pressure freezing/freeze substitution for electron microscopy. Glycogen was detected on periodic acid—Schiff stain, periodic acid—thiosemicarbazide-silver proteinate reaction, and the digestion test with α-amylase. On embryonic day 14, glycogen began to accumulate in the proximal portions of the developing epithelial cords. On embryonic day 17, marked glycogen particles were seen at the basal portion of the ductal epithelial cells and an abrupt increase of glycogen accumulation occurred in the secretory cells in the terminal bulbs. Ultrastructural observation indicated large clumps of glycogen particles localized in the basal portion of the terminal bulb cells. The initiation of glycogen accumulation preceded the formation of lumens in the ducts and terminal bulbs. Furthermore, proliferation analysis by bromodeoxyuridine labeling showed that this glycogen accumulation followed the cessation of the epithelial cell proliferation. Postnatally, glycogen accumulation in the terminal bulbs became gradually inconspicuous and completely disappeared by postnatal day 3, but that in the ducts was retained until around postnatal day 12. Temporary glycogen accumulation after the cell proliferation and before/during the lumen formation and secretory granule formation suggests significant involvement of the carbohydrate metabolism in the organogenesis of the submandibular gland.

In this study, a self-cooling laboratory system was used for pressure–shift freezing (PSF), and the effects of pressure–shift freezing (PSF) at 150 MPa on the quality of largemouth bass (Micropterus salmoides) during frozen storage at −30 °C were evaluated and compared with those of conventional air freezing (CAF) and liquid immersion freezing (LIF). The evaluated thawing loss and cooking loss of PSF were significantly lower than those of CAF and LIF during the whole frozen storage period. The thawing loss, L* value, b* value and TBARS of the frozen fish increased during the storage. After 28 days storage, the TBARS values of LIF and CAF were 0.54 and 0.65, respectively, significantly higher (p < 0.05) than the 0.25 observed for PSF. The pH of the samples showed a decreasing trend at first but then increased during the storage, and the CAF had the fastest increasing trend. Based on Raman spectra, the secondary structure of the protein in the PSF-treated samples was considered more stable. The α-helix content of the protein in the unfrozen sample was 59.3 ± 7.22, which decreased after 28 days of frozen storage for PSF, LIF and CAF to 48.5 ± 3.43, 39.1 ± 2.35 and 33.4 ± 4.21, respectively. The results showed that the quality of largemouth bass treated with PSF was better than LIT and CAF during the frozen storage.

Electron microscopy (EM) offers unparalleled power to study cell substructures at the nanoscale. Cryofixation by high-pressure freezing offers optimal morphological preservation, as it captures cellular structures instantaneously in their near-native state. However, the applicability of cryofixation is limited by its incompatibility with diaminobenzidine labeling using genetic EM tags and the high-contrast en bloc staining required for serial block-face scanning electron microscopy (SBEM). In addition, it is challenging to perform correlated light and electron microscopy (CLEM) with cryofixed samples. Consequently, these powerful methods cannot be applied to address questions requiring optimal morphological preservation. Here, we developed an approach that overcomes these limitations; it enables genetically labeled, cryofixed samples to be characterized with SBEM and 3D CLEM. Our approach is broadly applicable, as demonstrated in cultured cells, Drosophila olfactory organ and mouse brain. This optimization exploits the potential of cryofixation, allowing for quality ultrastructural preservation for diverse EM applications.

Synaptic vesicles can be released at extremely high rates, which places an extraordinary demand on the recycling machinery. Previous ultrastructural studies of vesicle recycling were conducted in dissected preparations using an intense stimulation to maximize the probability of release. Here, a single light stimulus was applied to motor neurons in intact Caenorhabditis elegans nematodes expressing channelrhodopsin, and the animals rapidly frozen. We found that docked vesicles fuse along a broad active zone in response to a single stimulus, and are replenished with a time constant of about 2 s. Endocytosis occurs within 50 ms adjacent to the dense projection and after 1 s adjacent to adherens junctions. These studies suggest that synaptic vesicle endocytosis may occur on a millisecond time scale following a single physiological stimulus in the intact nervous system and is unlikely to conform to current models of endocytosis. Neurons communicate with one another at junctions called synapses. When an electrical signal travels along a neuron and arrives at a synapse, vesicles filled with small neurotransmitter molecules fuse with the cell membrane and release the neurotransmitter. These chemicals rapidly bind to receptors on the downstream neuron that induce an electrical response in that cell. Vesicles can be consumed at prodigious rates, up to 500 a second, so the cell must recover the membrane rapidly and regenerate more vesicles filled with neurotransmitter. Experiments in the 1970s and 1980s suggested that when vesicles empty their contents into the synapse, they fuse completely with the membrane and are lost. To recover the membrane, the cell forms ‘pits’, by means of a coat protein called clathrin, which then bud off into the cell as new vesicles. It takes roughly 15–20 s for vesicles to be recycled in this way. By contrast, synapses with very high firing rates are thought to recycle vesicles through a faster process known as ‘kiss and run’, in which vesicles are not fully integrated into the membrane, but instead fuse transiently with it to form a reversible pore within about a second. However, these studies triggered vesicle release using conditions that are unlikely to occur naturally inside cells. Now, Watanabe et al. have used optogenetics to study vesicle recycling in response to single stimuli at the synapse between neurons and muscles in an intact living animal, the nematode C. elegans. The worms had been genetically modified to express a light-sensitive ion channel called channelrhodopsin in their motor neurons. Watanabe et al. used a single pulse of light to stimulate vesicle release, and then rapidly froze the worms before studying their synapses with electron microscopy. They found that vesicle recycling occurred at the edges of the synapse or at a specialized structure in the middle of the synapse. Vesicle recycling took less than 50 ms—much faster than anything previously observed. This ultrafast recycling is unlikely to occur via ‘kiss and run’ since recycling occurred at sites lateral to the sites of fusion and because the recycled vesicles were larger than the originals, implying that they had not simply re-formed after a brief fusion event. By using physiologically relevant stimuli in an intact animal, Watanabe et al. reveal that vesicles can be recycled at synapses much more rapidly than previously thought, suggesting that our current models of this process may need to be reassessed.

Ribbon synapses of cochlear inner hair cells (IHCs) are specialized to indefatigably transmit sound information at high rates. To understand the underlying mechanisms, structure-function analysis of the active zone (AZ) of these synapses is essential. Previous electron microscopy studies of synaptic vesicle (SV) dynamics at the IHC AZ used potassium stimulation, which limited the temporal resolution to minutes. Here, we established optogenetic IHC stimulation followed by quick freezing within milliseconds and electron tomography to study the ultrastructure of functional synapse states with good temporal resolution in mice. We characterized optogenetic IHC stimulation by patch-clamp recordings from IHCs and postsynaptic boutons revealing robust IHC depolarization and neurotransmitter release. Ultrastructurally, the number of docked SVs increased upon short (17–25 ms) and long (48–76 ms) light stimulation paradigms. We did not observe enlarged SVs or other morphological correlates of homotypic fusion events. Our results indicate a rapid recruitment of SVs to the docked state upon stimulation and suggest that univesicular release prevails as the quantal mechanism of exocytosis at IHC ribbon synapses.