1.1 The Main Intercellular Junctions and Their Proteins

Tight junctions, adherens junctions and desmosomes are composed of transmembrane proteins that form extracellular adhesive contacts between cells while intracellularly, the junctional proteins are also linked with the cytoskeletal structural components of the cell (Hartsock and Nelson, 2008). Both adherens and tight junctions are closely associated with a circumferential belt of actin filaments. In this review we cover imaging techniques used to study the adhesion complexes in highly polarized epithelial cells and flat endothelial cells (Figure 1). Proteins in the tight junction barrier are known to regulate intercellular communication and paracellular transport between cells. These tight junctions between two neighboring cells are often 200–500 nm in length and 11–15 nm wide, while the intermembrane space at these junctions is only 10 nm (Farquhar and Palade, 1963). Adherens junctions are important for multiple functions including initiation and stabilization of cell-cell adhesion, regulation of actin cytoskeleton, intracellular signaling and transcriptional regulation. These junctions are 200–500 nm long and 35–50 nm wide with 20 nm gap between opposing membranes. Desmosomes provide strong adhesion between cells and mediate cell-cell contact. Desmosomes are 200–300 nm long and can span up to 100 nm in width while the intermembrane space varies between 20 and 25 nm. Reviewed in Adil et al. (2021) and more references are found in the legend of Figure 2.

Open in a separate window

FIGURE 1

Schematic overview of the organization of intercellular junctions in epithelial and endothelial cells. (A) Tight junctions, adherens junctions and desmosomes are present in polarized epithelial cells while no desmosomes are found in flat endothelial cells. Tight and adherens junctions are connected to the actin cytoskeleton while desmosomes connect to intermediate filaments. (B) Organization of intercellular junctions during the polarization of epithelial cells. In the initial stage of polarization, proteins like afadin and nectins are present. They recruit E-cadherin and junctional adhesion molecules (JAMs) to the lateral cell-cell contacts, called the primordial adhesions in immature epithelial cells. These mature to form belt-like adhesion junctions and tight junctions localized at the apical-basal membrane border. The mechanism behind the polarization is reviewed in (Martin-Belmonte and Perez-Moreno, 2012). The overall height of a polarized cell grown in vitro is ~ 15 μm, figures are not on scale. (C) Overview of the different microscopy techniques that are discussed in this review to study the structure and composition of the three types of intercellular junctions. LM: light microscopy; TEM: transmission electron microscopy; ET: electron tomography; VEM: volume electron microscopy; SXT: soft X-ray microscopy; SIM: structured illumination microscopy; STED: stimulated emission depletion; SMLM: single molecule localization microscopy; FRET: Forster resonance energy transfer; Immuno-EM: immuno electron microscopy.

Open in a separate window

FIGURE 2

Schematic overview of the different components of intercellular junctions in epithelial and endothelial cells. (A) Highly polarized epithelial cell-cell contacts are composed of tight junctions, adherens junctions and desmosomes from the apical to basal side. The transmembrane proteins of tight junctions are claudins, which largely determine the paracellular ion permeability between cells, and proteins like occludin and junctional adhesion molecules (JAMs) (Zihni et al., 2016; Otani and Furuse, 2020). The cytosolic plaque of the tight junctions is a complex protein network, with adaptor proteins containing multiple protein-protein interaction motifs that are connected to F-actin and microtubules. One of the main cytoplasmic tight junction proteins are of the zonula occludens (ZO) family. Adherens junctions consist of transmembrane cadherin proteins, such as E-cadherin, and cytoplasmic proteins that are members of the catenin family; p120-catenin binds to E-cadherin closest to the intercellular space while α-catenins anchor the cadherin-catenin complex to the actin cytoskeleton by binding to the armadillo proteins β-catenin and/or plakoglobin (Gumbiner 2005; Perez-Moreno and Fuchs, 2006; Meng and Takeichi, 2009). The main transmembrane proteins in the desmosomal complex are two types of cadherins: desmogleins and desmocollins. They form heterodimers that make up the fundamental adhesive unit of desmosomes (Harrison et al., 2011). The cytoplasmic side of the desmosomal cadherins are linked with plakophilin and plakoglobin, which in their turn bind both to desmoplakin to make up the connection to the intermediate filament (keratin) complex of the cell (Green and Gaudry, 2000; Garrod and Chidgey, 2008). (B) Tight junctions in flat endothelial cells are very similar to those in polarized epithelial cells. The major difference is that there is no microtubule to bind to in the endothelial cells. Adherens junctions consist of vascular endothelial cadherin (vascular-endothelial) in endothelial cells and do not contain plakoglobin as cytoplasmic protein. Transmembrane nectin protein attached to the cytoplasmic afadin can be found in endothelial cells (Wallez and Huber, 2008). The overall height of a polarized cell grown in vitro is ~15 μm, figures are not on scale.

These spatially defined adhesion complexes are also known as signaling hubs that cross-talk in order to coordinate tissue organization and function. Malfunction of one type of adhesion complex, for instance by deleting one of the crucial junctional proteins, not only affects the function and/or organization of that specific junction type but can also impair other intercellular junctions [reviewed in (Rübsam et al., 2018)]. Moreover, in some tissue types mixed types of junctions can be found, i.e., area compositae at the intercalated disc of cardiomyocytes. There desmosomal proteins can be found in the adherens junction area (Borrmann et al., 2006; Franke et al., 2006). In flat endothelial cells, tight junctions and adherens junctions can overlap and intercalate. Tight junctions in endothelial cells are very similar to the ones in polarized epithelial cells, with the major difference that there is no microtubule to bind to in the endothelial cells (Figures 1, ​,22).

In addition to junctions between two neighboring cells, tricellular junctions can be formed at the corners where three cells meet. Consequently, tricellular contacts require more complex junctions and are reflected in the components present, such as angulins and tricellulin in the tight junctions at these tricellular borders (Zihni et al., 2016) or another plakophilin (Pkp) isoform in tricellular junctions in keratinocytes (Keil et al., 2016; Rietscher et al., 2018).

3.1 2D Structural Information

3.1.1 Transmission Electron Microscopy

The technique of TEM is based on an electron beam that passes through the sample (thin sections of 60–80 nm of tissue or cultured cells) such that the beam will be absorbed and scattered by the sample, producing contrast that can be imaged (Winey et al., 2014) (Figure 4A and Figure 6). Using electrons as a source instead of light has the advantage that imaging can be performed at a much higher spatial resolution. The wavelength of an electron beam is much shorter than that of visible light and images with a resolution up to 2 nm can be obtained. Therefore, EM can be applied for high-resolution (ultra)structural analysis of whole tissues. Disadvantages are the limited sample depth the electrons can penetrate, imaging is done under vacuum conditions and there can be radiation damage. Therefore, microscope design, sample preparation, imaging and image processing must be carefully optimized. The quality of an image is mainly defined by three factors: contrast, resolution, and SNR [reviewed in (Franken et al., 2020)]. Staining of the sample with heavy metals is performed to increase the scattering events caused by the electrons when they interact with the sample, leading to a higher contrast of the image. Because the specimen chamber is under vacuum, samples need to be highly processed and are no longer in the natural state. Samples can be prepared by chemical fixation, dehydration, and staining with heavy metals which enhances contrast as mentioned above and protects somewhat against dehydration and radiation damage. Another way to preserve the tissue is cryo-fixation (cryo-EM); i.e. samples on a grid are frozen (either by plunging into a liquid cryogen or placing under high pressure, called plunge-frozen or high-pressure frozen respectively), using liquid ethane or ethane/propane to avoid the formation of ice crystals. Only thin samples (less than ≈10 µm) can be plunge-frozen, while specimens up to 300 µm can be high-pressure frozen, to achieve sufficient freezing of the sample, including the center (Dubochet et al., 1988). Cryo-EM can be performed without heavy metal staining and the contrast is then based on phase contrast only. The advantage is that it resembles the most native state and results in a high resolution. Thin samples are obtained by a process called ultramicrotomy, which can be done at room temperature or at sub-zero temperatures which can be challenging (Richter et al., 2007). In order to create thin sections from thicker samples, recent developments including cryo-EM of vitreous sections (CEMOVIS), or cryo-focused ion beam (FIB) milling can be used to create thinned areas within cryo-frozen specimens [reviewed in (Franken et al., 2020)]. Imaging of cryo-frozen samples require imaging in a cryogenic environment, which is technically demanding. High-pressure frozen samples can be stained by freeze-substitution, embedded in plastic resin, cut by ultramicrotomy and imaged at room temperature. More information about TEM of biological samples can be found in a book chapter (Mielańczyk Ł and Romuald, 2015).

Open in a separate window

FIGURE 6

Schematic overview of electron and X-ray based techniques for imaging the tight junctions, adherens junctions and desmosomes. Transmission electron microscopy (TEM) uses an electron beam that passes through the sample (thin sections of 60–80 nm of tissue or cultured cells; either fixed or cryo-frozen because of vacuum conditions). The electron beam is absorbed and scattered by the sample which in their turn are captured on a detector and a contrast image of the sample can be seen. Electrons can only penetrate a very thin sample. In freeze fracture EM (FFEM) frozen biological samples are physically broken apart and platinum-carbon is used to replicate and contrast the fracture plane, which is then analyzed by TEM, exposing the intercellular junctions between cells. The lateral (xy) resolution is 2 nm. For an example of an image, see Figure 4A. Electron tomography (ET) is an EM-derived technique and is based on a tilt series of 2D images acquired at different viewing angles, which are subsequently aligned and combined digitally into a 3D volume. Tilt series contain projections from a complete ±60–70° rotation of the object to include projections from all possible directions. This gives a 3D view through the depth of the specimen at a high resolution and the intercellular junctions can be reconstructed in 3D. The sample must be thin sections (tissue or cultured cells; limited to about 300 nm; fixed or cryo-frozen) to render them electron transparent. The lateral (xy) resolution is 2 nm. For an example of an image, Figure 4B. Volume scanning electron microscopy (volume-SEM) can image larger volumes with a greater sample depth compared to ET. The technique is based on the alternation between scanning the tissue surface with a focused electron beam followed by cutting off a section of the en-bloc tissue. The electron interact with the tissue surface and the backscattered electrons (BSE) are then detected by a detector. Cutting the section is done by either a diamond knife [serial block face (SBF)-SEM] or an ion beam [focused ion beam (FIB)-SEM], enabling the cutting of sections with a thickness of 25–50 and 5 nm respectively. The maximum sample width for SBF-SEM is around 1 mm and 20–100 µm width for FIB-SEM. The sample preparation often includes heavy metal staining for contrast, embedding in plastic resin and coating of the sample with a thin layer of conductive material (i.e. gold) to reduce the charging effects. The resulting 2D stack of images can be then 3D reconstructed to see the intercellular junction in a volume. The voxel resolution is up to 5 nm. For an example of an image, see Figure 4B. Soft x-ray tomography (SXT) uses soft X-rays as an illumination light source. Projections are taken from different angles around a sample (cells in suspension or on a grid; cryo-frozen) by rotating the sample along its long axis. SXT image the sample in the water window, meaning that X-rays are absorbed by carbon and nitrogen in biological tissue more than by oxygen in water, resulting in grayscale that can be used for quantitative analysis. Soft X-rays have a limited penetration into the sample thus only samples of 10–20 µm can be imaged. The voxel resolution is 10–50 nm. Figures are not on scale.

EM made a major contribution to the visualization of intercellular structures by unraveling the adhesions in high detail. Ideally, the connection of the junctions to the cell’s cytoskeleton and membrane elements are imaged together to observe how they make an active junctional complex (see below). Additionally, individual proteins can be localized in context of the 3D volume.

Overall, researchers see that desmosomes are electron-dense and surrounded by a fuzzy area while adherens junctions are less electron-dense with the same intermembrane distance. For tight junctions however, there are different observations. In chemically fixed intestinal mucosa of the rat, tight junctions were seen as electron-dense apical membrane contacts forming a continuous belt-like attachment over a 200–500 nm distance by bringing the adjacent cell membranes in very close proximity (kissing points) (Figure 4A1) (Farquhar and Palade, 1963). Later, FFEM showed that tight junctions form interconnecting strand networks between cells that vary in number and morphology among different tissues. The frozen biological samples (with possible chemical fixation prior to freezing) are physically broken apart, typically fracturing at weak hydrophobic sites such as membranes, and a platinum-carbon replicate of the fracture plane is then analyzed by TEM, exposing the intercellular junctions between cells (Figure 4A3) (Chalcroft and Bullivant, 1970; Claude and Goodenough, 1973; Staehelin 1973; van Deurs and Koehler, 1979; Furuse 2010). FFEM has a limited spatial resolution due to the inherent property of metal atoms to crystallize. Recently Krystofiak et al. used a combination of amorphous carbon replicas with phase-contrast EM (Danev et al., 2014) to overcome this (Krystofiak et al., 2019). With this approach, they found that tight junction strands have an antiparallel double-stranded morphology.

When looking at the connection with the cytoskeleton, early observations using TEM of chemically fixed tissue section demonstrated that tight junction-associated actin filaments occur predominantly at sites of intercellular membrane apposition. These actin filaments are decorated by the actin binding region of myosin and appear to insert directly into the submembrane tight junction space (Madara and Pappenheimer, 1987). However, TEM of quick-freeze, deep-etch, rotary shadow replicas made it possible to distinguish between two different actin populations in epithelial cells (Höflinger 2014). Actin at the adherens junctions is organized as a ring and is composed of filaments that run parallel to the plasma membrane. This differs from the actin filaments found just beneath the tight junction membrane; these are organized as a meshwork of filaments (Hirokawa and Tilney, 1982; Hirokawa et al., 1983). Later, similar TEM-based techniques showed different features of the adherens junctions in a single layer of epithelial cells (chemically fixed and frozen). When sections were cut perpendicular to the cell membrane it revealed that adherens junctions have a 15–25 nm space between both membranes (Figure 4A2). With freeze-etching, it became clear that they are made up of a cytoplasmic macromolecular complex that consists of rod-like structures extending from the extracellular surface into the intercellular space, presumably catenin proteins of which the extracellular part may cant at about 60° with the plasma membrane (Miyaguchi 2000; Sluysmans et al., 2017). The standard TEM of chemical fixed cultured cells grown on a grid confirmed that the actin filaments run parallel to the adherens junctions (Buckley et al., 2014).

Cryo-EM of desmosomes in high-pressure frozen tissue sections showed that the desmosomal intercellular space has been variously reported as between 20 and 35 nm wide (Al-Amoudi et al., 2004). It is characterized by the presence of a dense midline with cross-bridges extending to the plasma membrane. When injecting the extracellular space of desmosomes with the electron-dense tracer lanthanum, researchers saw that the cross-bridges extending to the plasma membrane appear to alternate on opposite sides of the midline and that the desmosomal cadherin extracellular domains are ordered in all 3 planes (x, y, and z) (Rayns et al., 1969). So, the high resolution of EM and related techniques, including tracers and freeze-etching, allows researchers to distinguish the extracellular domains of cadherins in the intercellular junctions (Miyaguchi 2000; Al-Amoudi et al., 2004).

An evolved technique of rotary shadowing EM to reveal the surface topography of a sample is platinum replica EM (PREM). PREM is a specific type of EM where a cell on a glass coverslip is extracted or unroofed, followed by fixation and critical point drying. Then platinum is evaporated on a 3D sample at an angle during rotation of the sample after which carbon is reinforced on it (Heuser 1981). This reveals the topography and makes it an ideal technique for the structural analyses of the cytoskeleton (Svitkina and Borisy, 1998; Svitkina 2016). PREM achieves the high resolution typical for EM and the cytoskeletal structures are well preserved with visibility of single filaments, even if they are densely packed. Sample preparation for PREM is fast and efficient making it an inexpensive and accessible technique. However, only thin samples can be imaged as only the surface is made visible. The technique requires samples to be attached to glass surfaces which is not ideal to study membrane structures. Because of its informative value for cytoskeleton studies, PREM has been used and revealed the double-stranded morphology of the tight junction intermembrane fibrils (Krystofiak et al., 2019). In combination with light microscopy and immunolabeling for vascular endothelial cadherin with immunogold, it revealed the relationship between the parallel extensive branched actin network and the junctions between two endothelial cells (see Figure 4A4) (Svitkina 2017; Efimova and Svitkina, 2018). Specifically, the vascular endothelial cadherin colocalizes with the Arp2/3 complex of the F-actin network at different adherens junction types and is positioned at the interface between two oppositely oriented branched networks from adjacent cells. It is also proposed that the actin cytoskeleton at adherens junctions is a dynamic push-pull system (Efimova and Svitkina, 2018).

TEM allows imaging of intercellular junctions and ultrastructure of tissue or cultured cells at a very high (nanometer) resolution. Junctions in a cell layer are imaged perpendicular to the cell surface layer and with some techniques (like freeze-etching and PREM), structures just beneath the cell surface can be exposed and imaged. This is a suitable technique for visualizing the morphology of junctions, unraveling the relationship between the actin cytoskeleton and the intercellular complexes—with PREM it is even possible to see individual filaments in a 3D manner—and for measuring the intermembrane space between cells or see the extracellular domains of cadherin proteins. PREM requires some technical handling of the samples and optimization of the sample preparation.

3.2 3D Structural Information

When imaging in 2D, the spatial relationship between the optical axis of the microscope and the sample is important to interpret the way one looks at the junctions. When imaging samples in 3D, however, this is (partly) overcome by combining multiple 2D images, either from different angles—like electron tomography (ET) and soft X-ray tomography (SXT)—or from a stack of images that represent a voluminous sample (scanning EM, SEM). When reconstructing intercellular junctions, they can be seen from all sides in relation to the surrounding cells.

5.1 Sample Preparation and Sample Type

One of the main considerations is the sample preparation and the type of sample you are dealing with. There is the possibility to perform in vitro, in vivo, and ex vivo imaging. Some techniques do not allow for in vivo imaging, for instance due to the destructive nature of the microscope, while all super-resolution techniques do allow live imaging. Intravital microscopy for instance provides imaging of cellular events in its native tissue environment as well as in a real-time setting. It has several compelling advantages to study intercellular junctions. For example, researchers visualized E-cadherin labeled cell-cell junctions in mouse pancreas using intravital microscopy combined with multiphoton imaging (Erami et al., 2016). However, imaging in live animals has many technical challenges while it also requires access to high-resolution microscopy techniques (reviewed in (Choi et al., 2015; Hickey et al., 2022). Smaller mammalian species might be considered, including larval zebrafish, because of their high optical transparency that enables direct visual access to biological processes (Ahrens et al., 2012; Høgset et al., 2020). Next to imaging in live animals, freshly excised organs can be studied, followed by optional processing (fixation, freezing or staining) a process termed ex vivo imaging. This processing can alter the native state of the tissue and the morphology, as well as the interpretation of functional studies. However, it is still one of the preferred methods for imaging intercellular junctions (see below) and possible with all discussed imaging techniques. Finally, in vitro imaging (imaging of cell cultures; imaging can be performed with all discussed techniques) has many advantages, including reduced cost, higher throughput, ease to manipulate and reduced need for animals. The most significant drawback is that cell culture models may fail to replicate in vivo biology since they lack the complex network of interactions or intercellular signaling, which occurs in tissues, organs or whole organisms. Recent advances do include co-cultures of multiple cell types and researchers are increasingly using 3D cell models to recreate organized, miniaturized versions of organs to study diseases (Martin 2020). All three types of imaging ways have their advantages and disadvantages as well as technical difficulties and necessary equipment (reviewed in (Hickey et al., 2022)).

In vivo imaging enables the study of dynamic cell behavior but is not possible with techniques that use electrons or X-rays. Another variable is the temporal resolution when imaging with confocal and super-resolution microscopy. Some techniques, like SIM, allow for a very fast imaging speed (sub-seconds) while others are much slower (including SMLM, seconds to minutes and STED, 10 ms to minutes). In living cells, direct in vivo visualization of protein-protein interactions is also possible with a technique called bimolecular fluorescence complementation (BiFC) (Hudry et al., 2011). For instance, this technique helped researchers to unravel the role that the tight junction claudin-2 plays in the entry of the reovirus into the cell (Zhu et al., 2021). Examples of which super-resolution techniques that can be used for tight junction and adherens junction visualization in live-cell or in fixed samples are reviewed in (Gonschior et al., 2020).

In vitro imaging of cultured cells is a standard approach in biological research. However, when fully polarized epithelial cells, in monolayer, are grown on a 2D support (such as a glass coverslip), the intercellular junctions are located 10–20 microns away from the growth substrate and are thus not suitable to be imaged with super-resolution techniques like SMLM (as explained above). The principle and important considerations for tight and adherens junction imaging are reviewed (Gonschior et al., 2020). Other approaches, including cells grown on a filter membrane followed by mounting on a coverslip [as described above (Maraspini et al., 2019)] can overcome this orientational problem during imaging. This still requires the cell-cell junctions to be imaged along the optical axis, which provides only limited information on their organization along the lateral membrane. The axial resolution of most light-based techniques is not enough when imaging deeper in the cells. Next to growing cells in 3D [like organoids (Martin 2020; Hickey et al., 2022)], imaging epithelial or endothelial tissue sections is another good approach. Tissue sections have the major advantage of having preserved complex interactions among cells and their microenvironment and therefore cells kept their cell polarity in the hierarchical architecture of the tissue. It also allows the researchers to image cell junctions in a side view (x/y) when cut at the right angle. With 3D imaging techniques (such as ET, SXT and volume-SEM), the junctions can be seen from different angles, reconstructed as one volume among cells and analyzed from different orientations.

Not only the way of sample imaging, but also the state in which it is imaged is a key step in the decision-making process. Some imaging techniques require fixation because of their destructive technique, for example when imaging with electrons and X-rays (Popescu et al., 2016). New developments are emerging to overcome this issue (Elphick et al., 2021). Chemical fixation sometimes followed with the embedding of the sample in plastic for EM processing, is a widely used technique but can affect the structure of the sample due to dehydration. To make sure the ultrastructure is preserved, cryo-freezing has become a more popular fixation method in recent years; it gives excellent preservation of the material in a near-native state compared to chemical fixation (Dahl and Staehelin, 1989; Shimoni and Müller, 1998). Sectioning cryogenic material (cryo-ultramicrotomy) for the preparation of ultrathin sections, as necessary for TEM, involves sophisticated tools and is technically demanding (Studer et al., 2008; Glaeser et al., 2016). Most fluorescent imaging techniques, either confocal or super-resolution, allow for imaging on living cells/samples within a controlled environment and the possibility of time-lapse imaging for days. However, possible phototoxic events after long illumination, as seen in super-resolution techniques like STED and SIM, need to be taken into consideration [reviewed and solutions in (Tosheva et al., 2020; Hickey et al., 2022)].

In some cases, it is very difficult to image the intercellular junctions in their native localization for several reasons. Therefore, researchers have engineered other techniques that allow them to image individual proteins/complexes at a high resolution. For instance, the cryo-EM image of isolated desmosomal fractions has been used to produce a molecular model of desmosomes (Sikora et al., 2020). Even though this has helped researchers to understand some biological questions, the physiological relevance of this type of set-up can be discussed.

5.2 Orientation of Imaging

Different techniques have different imaging depths, as discussed above (Table 1 and ​and2).2). Intercellular junctions are located at the lateral membrane perpendicular to the optical axis of most techniques (Figures 5, ​,6).6). Depending on the cell type studied the junctions are hundreds of nanometers to tens of micrometers away from the coverslip (Figure 1), thus they require an imaging technique with a certain illumination depth.

Researchers have developed many adjustments to confocal and super-resolution techniques to meet these requirements. However, the higher the lateral (xy) resolution, the more superficial the imaging must be. For instance, a technique with a very high lateral (xy) resolution is TIRF microscopy. The excitation light in TIRF is shined at a very high angle, causing it to internally reflect and no out-of-focus light is generated as only a very thin (~0.2 um) layer adjacent to the coverslip is excited, with a high SNR as a result (Fish 2009; Mattheyses et al., 2010). TIRF is often used as a setup during SMLM imaging which gives superb lateral (xy) resolution. However, only a limited field of view can be imaged and junctions within the tissue cannot be resolved with this technique (Tokunaga et al., 2008; Gerb and Jacob, 2011). Moreover, combination with other super-resolution methods with more specialized illumination geometries such as two-photon or light-sheet in addition to tissue-clearing and adaptive-optics based corrections could be promising to unravel the complex nature of junctions [as reviewed in (Bond et al., 2022)]. However, this has not been performed much for tight junctions, adherens junctions or desmosomes. Tissue clearing gives the opportunity to image with reduced light scattering and increased imaging depth with a range of microscopic and tomographic methodologies of larger and thicker samples (Almagro et al., 2021). Developments in the optics aspect of microscopy enabled researchers to image, with SIM, samples thicker than a single cell (Lin et al., 2021). This could be promising to image junctions in tissues instead of cell cultures with super-resolution microscopy.

If one wants to image deeper into the tissue and look at larger volumes, 3D imaging techniques using X-rays and electrons are ideal. Using volume-SEM, ET and SXT allow researchers to image deep into tissue, in intact organisms and to reconstruct the intercellular junctions in 3D. However, it is limited to fixed samples (chemically or cryo-frozen; as mentioned above).

5.3 Labeling and Controls

During imaging, researchers want to image the correct protein(s) and need to be sure that what they see is what they intended to see. Labels/probes need to bind the protein of interest correctly and these can be directly visualized or followed by targeting of an imaging label (in case of fluorescence microscopy) or gold particle (in case of EM). Therefore, applying appropriate controls when setting up an experiment is highly desirable [discussed in (Hickey et al., 2022)]. The easiest are techniques without the need for labelling to see subcellular structures, which include SXT and cryo-EM. The downside is that the distinction of the cellular components relies on the expertise of experienced scientists.

Antibodies are the most widely used probes, they are highly specific to their targets and there is a large selection available. However, they take up quite some space (15 nm in length) and their dual binding capability can introduce significant artifacts in super-resolution imaging [reviewed in (Fornasiero and Opazo, 2015)]. To reduce the size, antibody fragments or single-chain antibodies (nanobodies) were developed. As epitopes must be labeled at high density for accurate super-resolution imaging, the fluorophores must be as close as possible to their targets and nanobodies are perfect for this. However, one must make sure all endogenous proteins are imaged as antibodies have been known to miss some when detecting GFP-tagged proteins. Other options are aptamers, single-stranded DNA or RNA oligonucleotides that are selected in vitro for the desired targets, showing improved precision obtained in STED and SMLM when compared to antibody staining (Opazo et al., 2012). More small molecules and scaffolds are produced to make single localization imaging possible (Fornasiero and Opazo, 2015).

5.4 Post-Processing

Once images are taken, the data must be processed, stored and analyzed. This can get very complex, time- and computer-demanding depending on the technique used. 3D acquired data (from ET, SXT and volume-SEM) needs some post-processing, including corrections, stack alignments and sub-tomogram averaging, before analysis can be performed (Schur 2019; Pyle and Zanetti, 2021). EM and SXT data analysis is often carried out by specialists with experience in the identification and interpretation of biological features in the complex grayscale world of this type of imaging. Manual analysis is labor-intensive and slow. Automation with the use of machine learning is needed to accelerate the speed and efficiency of segmentations and structural analysis (Peddie and Collinson, 2014; Ekman et al., 2020; Lösel et al., 2020). Also, methods for good data storage and computing are evolving (Baldwin et al., 2018; Vescovi et al., 2020).

Evolution to nanometer resolution microscopy has been extremely fast in the last decade. The increased resolution imposes stringent conditions on data analysis and researchers must ensure that photophysical properties of the probes, the labeling, and the imaging strategies are correct and do not lead to misinterpretation of the data. New methods have been developed for proper analysis and quantification of imaged data by super-resolution techniques (Durisic et al., 2014). For the super-resolution techniques, SMLM data analysis is quite computational demanding, while SIM is less and STED is fairly straightforward (Bond et al., 2022).

The authors would like to thank the two reviewers for their helpful comments.