Tissue clearing is basically an extension of the conventional histology technique that literally makes tissue “transparent” by suppressing light scattering and light absorption in the fixed tissue sample. Since Lundvall and Spalteholz initialized clearing of human tissues with the use of organic solvents more than 100 years ago (Lundvall, 1905; Spalteholz, 1914), the field has achieved dramatic innovations in the last few decades, as dozens of protocols, categorized as organic solvent (hydrophobic reagent)-based protocols, hydrophilic reagent-based protocols, and hydrogel-tissue chemistry (Ueda et al., 2020a,b), have been recently established.

Optical clearing generally involves exchanging the surrounding medium (e.g., phosphate-buffered saline) with a solvent with optical properties similar to those of the biomaterial. This step is called refractive index (RI) matching because the RI is a preferred indicator of optical properties. However, this process is relatively complex physicochemically and not necessarily limited to RI homogenization, as dispersion of the fibrous structures of the extracellular matrix and the affinity (infiltration) of compounds into living tissues may also have significant effects (Ueda et al., 2020a; Yu et al., 2021). RI matching can also be combined with other processes to remove light-scattering and light-absorbing substances from the biological tissue, such as lipids (delipidation), the bone matrix (decalcification), and pigments (decolorization or bleaching). The requirement of incorporating these steps is dependent on the experimental purpose and target tissue type. Tissue clearing protocols incorporate these steps by combining various compounds and physical techniques (e.g., electrophoresis).

Organic solvent-based tissue clearing protocols originating from the Spalteholz reagent are represented by the benzyl alcohol/benzyl benzoate (BABB) method, 3D imaging of solvent-cleared organs (3DISCO), and the ethyl-cinnamate (ECi) method (Dent et al., 1989; Becker et al., 2012; Ertürk et al., 2012; Klingberg et al., 2017). Generally, these protocols have very strong clearing abilities over relatively short periods of time. However, it is necessary to understand the characteristics of the reagents regarding tissue shrinkage, signal retention of fluorescent proteins, safety, and compatibility with the microscope system.

The hydrophilic reagent-based clearing technique was initially tested with the use of several hydrophilic chemicals (e.g., sugars and alcohols) to transluce human skin and sclera in medical applications (Bakutkin et al., 1995; Zimnyakov et al., 1996; Tuchin et al., 1997). Since then, a wide variety of hydrophilic reagent protocols have been proposed with the advantages of ease of handling, safety, and preservation capacity of biomaterials, which include FocusClear^TM (Chiang et al., 2001), Scale (Hama et al., 2011, 2015), Clear^T (Kuwajima et al., 2013), SeeDB (Ke et al., 2013, 2016), FRUIT (Hou et al., 2015), CUBIC (clear, unobstructed brain/body imaging cocktail and computational analysis) (Susaki et al., 2014; Tainaka et al., 2018), FUnGI (fructose, urea, and glycerol for imaging) (Rios et al., 2019), RTF (rapid clearing method based on triethanolamine and formamide) (Yu et al., 2018), Ce3D (clearing-enhanced 3D) (Li et al., 2017), and TDE (2,2′-thiodiethanol) immersion (Aoyagi et al., 2015).

Hydrogel-tissue chemistry involves the preparation of a tissue-hydrogel scaffold by cross-linking hydrogel monomers to native biomolecules (Gradinaru et al., 2018). The initial formulation, called CLARITY (and its variations), uses acrylamide, while later versions, SWITCH (Murray et al., 2015) and SHIELD (Park et al., 2018), utilize glutaraldehyde and a polyepoxide, respectively. Due to the increased tissue rigidity caused by transformation, these protocols can be combined with harsh delipidation or re-probing procedures with sodium dodecyl sulfate (SDS) and physical electrophoresis. Furthermore, the use of a water-absorbing polymer for Expansion Microscopy (ExM) enables high-resolution imaging with a general microscopy setup (Chen et al., 2015; Ku et al., 2016). The expansion can also contribute to the final transparency of the sample. Other hydrophilic clearing reagents have also applied the same strategy (Kim et al., 2018; Murakami et al., 2018; Tainaka et al., 2018).

Although limited to fixed samples, tissue clearing facilitates observation of the 3D architecture of biological tissues with a light microscope at the cellular or higher resolution, thereby providing a powerful analytical approach for complex biological systems.

Clearing of cell reaggregates (spheroids and organoids) for whole-mount imaging was recommended in early protocols (Timmins and Nielsen, 2007). More recently, many of the clearing methods introduced in the previous section have been tested from simple whole-mount observations to advanced phenotyping of 3D cultured reaggregates (Costa et al., 2019; Table 1). Since most current clearing protocols are optimized for animal tissues and organs, clearing cell reaggregates with the use of any of these procedures is, in principle, much simpler. Although tissue clearing methods have been adopted at least to some extent, experience is required for further applications in broader organoid research.

A side-by-side comparison of protocols would be helpful to identify the clearing method that is most appropriate for a particular application. Boutin and Hoffman-Kim compared early phase hydrophilic clearing protocols (Clear^T2, ScaleA2, and SeeDB) and concluded that Clear^T2 is the most effective for clearing neural spheres (Boutin and Hoffman-Kim, 2015). However, in a later study, the authors also applied an updated Scale protocol (ScaleS) for clearing of cancer cell spheroids (Boutin et al., 2018b). Cheung and colleagues compared SeeDB, Clear^T2, and ScaleSQ for adaptation to an on-chip spheroid processing system and concluded that SeeDB and ScaleSQ were more effective to clear a sample than Clear^T2, although there were some drawbacks of increased autofluorescence and sample expansion (Grist et al., 2016). Schöler, Bruder and colleagues compared the performance of the organic solvent reagent BABB with several other types of clearing reagents and found that BABB-based clearing proved to be both the fastest and most efficient for clearing of human midbrain organoids (Renner et al., 2020). Garfa-Traoré and colleagues compared several clearing methods (TDE, CUBIC, and RapiClear^®) for murine intestinal organoids and obtained the best clearing and staining results with RapiClear^® (Lallemant et al., 2020). Rudolf and colleagues tested several hydrophilic reagents and mounting media (glycerol, ScaleS, Clear^T2, and CytoVista) for clearing of various spheroids and found that ScaleS and a high concentration of glycerol (88% RI = 1.459) provided the best clearing results, while preserving the fluorescent signals and maintaining sample integrity, although various factors (i.e., size, complexity, and composition) affected the clearing results (Nürnberg et al., 2020). Another study suggested similar performance of glycerol (>85%) and RapiClear^® for clearing of pancreatic tumor spheroids (Steinberg et al., 2020). Lorenzo and colleagues demonstrated the effectiveness of both CUBIC and CLARITY for clearing of tumor cell spheroids (Masson et al., 2015).

These mixed results indicate that there is no “gold-standard” protocol for clearing of all cell reaggregates. As with the clearing of large tissue samples, it is necessary to choose an appropriate protocol in consideration of the pros and cons. The complexity of a reaggregate can affect the efficiency of optical clearing (Nürnberg et al., 2020) and thus may occasionally require delipidation. For example, Paşca and colleagues clearly and quantitatively reproduced cortico-striatal projections in human induced PSC (iPSC)-derived cortico-striatal assembloids with the use of the latest CUBIC-L/R procedure (Miura et al., 2020). Chung and colleagues applied SHIELD technology to single-cell and cytoarchitecture combined with multiple labeling methods for analysis of organoids (Albanese et al., 2020). ExM (Chen et al., 2015) enables super-resolution imaging together with improved clearing and staining results. Brismar and colleagues applied ExM to probe labeling and for high-resolution imaging of tumor cell spheroids, and found that as compared to simpler clearing protocols, ExM improved antibody penetration and image resolution in deeper regions (Edwards et al., 2020).

Tailoring of key parameters (e.g., compound type, concentration, immersion time, and temperature) in the original protocol should also be taken into account. Correia and colleagues found that the molecular weight of polyethylene glycol in Clear^T2 reagent can affect clearing and imaging quality (Costa et al., 2018b). Molley et al. (2020) used the ScaleCUBIC-2 protocol for clearing of Matrigel-embedded microtumors with some modifications to timing, washing, and handling. Dekkers et al. (2019) designed a fructose-glycerol immersion method as a simple, non-toxic, optical clearing step for complete 3D imaging of fragile organoids. Later, the authors cleared human colonic organoids with their FUnGI clearing reagent that was originally developed for clearing of human cancer specimens (Rios et al., 2019; van Ineveld et al., 2020). Our group modified the delipidation stringency of our CUBIC protocol for clearing of large kidney organoids (Figure 1).

Other than clearing performance, the compatibility of the cell-labeling/probing methods and microscopy setup should also be considered, as discussed in the following sections.

Since tissue clearing alone cannot label objects in a 3D structure, appropriate cell/structure labeling with a fluorescent protein (FP) or histological staining is also required for observation. Therefore, it is crucial to consider the compatibility of the tissue clearing technique with various labeling methods.

Conventional organic solvents show weak retention of FP signals. When using reagents in this category with FP labeling, more FP-compatible protocols (Schwarz et al., 2015; Pan et al., 2016; Qi et al., 2019) should be considered. Tanaka and colleagues recently improved the ECi method for clearing of FP-labeled organoids (Masselink et al., 2019). Alternatively, clearing with hydrophilic reagents or tissue-hydrogel chemistry methods that generally retain FP signals should be considered. In this regard, some methods strictly compared the performance of keeping FP protein signals with other clearing protocols. For example, Ce3D has a superior ability to FP preservation when compared to major clearing methodologies (BABB, DISCO, Scale, ScaleCUBIC, SeeDB, Clear^T, CLARITY, PACT, and SWITCH) (Li et al., 2017).

Whole-mount labeling of organoids with molecular probes (antibodies and small chemical dyes) has also been widely applied in many organoid studies. However, probe diffusion in 3D specimens is generally problematic, even in “small” cell reaggregates. For example, Smyrek and Stelzer investigated multiple whole-mount spheroid immunostaining parameters, including permeabilization, incubation time, and temperature, based on classical immunostaining procedures and proposed parameter guidelines (Smyrek and Stelzer, 2017). However, the signal intensity and homogeneity were varied and dependent on the antibody type and treatment protocol. Schöler, Bruder and colleagues very recently reported staining, clearing, and quantification analysis of relatively large (> 800 μm diameter) organoids (Renner et al., 2020). While the procedure was successfully automated, the protocol required a long incubation time (total 12 days for primary and secondary antibody staining) with multiple renewals of the staining reagents.

Various tissue clearing methods for large-scale 3D staining have been developed with improved efficiency and homogeneity. Recent developments of improved and more versatile 3D staining protocols, such as iDISCO, AbScale/ChemScale, SWITCH, and CUBIC-HistoVIsion (Renier et al., 2014, 2016; Hama et al., 2015; Murray et al., 2015; Susaki et al., 2020), have enabled large-scale 3D tissue staining. iDISCO, with optimized permeabilization and staining steps, enables immunostaining of whole mouse embryos and brains. AbScale/ChemScale utilizes urea to facilitate probe penetration via Scale clearing technology. SWITCH modulates the kinetics of probe binding to tissue by two procedures: SWITCH-off (inhibits probe binding) and SWITCH-on (facilitates probe binding). In our latest study on CUBIC-HistoVIsion, biological tissue was modeled as an electrolyte gel for screening of multiple essential 3D staining conditions that provides highly optimized 3D staining of cleared specimens of an entire organ and the whole body. However, the current protocol is not applicable to multiplex immunolabeling. Alternative protocols (e.g., Ce3D, SWITCH) can be considered to avoid this drawback. The incorporation of these recent 3D staining strategies can overcome the drawbacks of labeling entire organoids.

Several studies have applied these recent 3D staining protocols for labeling of whole organoids. For example, Takagi and colleagues applied AbScale immunostaining together with ScaleS clearing to patient-derived tumor organoids for in vitro evaluation of molecularly targeted drugs (Takahashi et al., 2019). Another study employed AbScale and ScaleS for staining and clearing of various human cell spheroids (Nürnberg et al., 2020). In addition, Ergun and colleagues applied an iDISCO-based whole-organoid procedure to stain vascularized tumors and neural organoids (Wörsdörfer et al., 2019). Moreover, for staining of human cerebral organoids, Chung and colleagues applied a modified eFLASH protocol that enables homogeneous staining of 8–10 whole organoids simultaneously in 1–2 days (Yun et al., 2019). The versatile staining ability of their SCOUT method further supports system-level analysis of the framework of 3D organoids (Albanese et al., 2020).

As in the case of FP, it should be noted that retention of the stained signals is dependent on the clearing reagents, as some may remove a portion of the staining target, while others may alter the binding affinity or antigenicity (Lallemant et al., 2020; Matryba et al., 2020). For example, the intensive comparison by Li et al. (2017) showed that the preservation degree of antigenicity toward major cell type markers in the lymph node is varied among tested methods. Antigen retrieval methods have been incorporated in some 3D staining and clearing protocols (Messal et al., 2021). After staining, cross-linking is occasionally required to preserve binding of the probe when clearing a specimen (Susaki et al., 2020).

The acquisition of whole-organoid information requires a proper 3D imaging setup. Besides conventional line-scan imaging, such as confocal and two-photon microscopy, light-sheet fluorescence microscopy (LSFM) is especially useful for 3D observation of cleared whole specimens. LSFM excites the fluorescent signals within the cleared sample with a sheet-shaped illumination and acquires section images with a vertically positioned complementary metal-oxide semiconductor (cMOS) or charge-coupled device (CCD) camera. This setup enables collection of high-throughput 3D images of the entire sample with minimal photodamage. LSFM has thus been proposed as the optimal modality for high-speed, high-quality 3D imaging of cleared samples, including spheroids and organoids (Dodt et al., 2007; Pampaloni et al., 2007; Keller and Stelzer, 2008; Santi, 2011; Keller and Dodt, 2012; Costa et al., 2019). As compared with confocal and two-photon microscopy, experimental studies have reported the superior speed and depth of LSFM for imaging of 3D organoids (Lallemant et al., 2020).

Custom-built LSFM systems have been reported in earlier studies. For example, a series of studies reported organoid/spheroid clearing with LSFM imaging using a monolithic digital scanned laser light-sheet-based fluorescence microscope (mDSLM) (Keller et al., 2008; Schmitz et al., 2017; Smyrek and Stelzer, 2017) and a millimeter-scaled optofluidic lab-on-a-chip device, which integrates light-sheet illumination and a microfluidic channel, for imaging of cell spheroids (Paiè et al., 2016). Moreover, Lorenzo and colleagues improved the resolution of 3D spheroid images using a light-sheet setup with adaptive optics (Masson et al., 2015). A specific light-sheet setup, such as a dual-view inverted selective plane illumination microscope (Kumar et al., 2014; Eismann et al., 2020), open-top LSFM (Glaser et al., 2019), and single-objective LSFM (Li et al., 2014; Bouchard et al., 2015), have the potential to enable large-scale imaging for screening purposes owing to the compatibility of these systems for imaging of multiple organoids cultured in multi-well plates.

With the commercialization of LSFMs, the opportunities of 3D organoid imaging with clearing are beginning to expand. The ZEISS Lightsheet Z.1 has been one of the most popular commercial LSFMs so far, enabling high-throughput multi-view 3D imaging with an easy-to-use operation (Cora et al., 2019; Dekkers et al., 2019; Sakaguchi et al., 2019; Lallemant et al., 2020; Molley et al., 2020; Preusser et al., 2020). However, the RI range of the equipped objective lens is up to 1.48, which is incompatible with some transparent reagents with an RI of > 1.5 and thus can cause the formation of spherical aberrations when imaging large samples at higher magnifications. The recently released Lightsheet 7 improves this issue with an RI range of 1.33–1.58, allowing the imaging of samples up to 2 cm in size with the use of almost any clearing reagent.

Regardless of the type of microscope used, the design of the entire experimental workflow should be optimized by selecting an appropriate clearing method, microscope setup, objective lens specification, and image resolution. Compatibility of some organic solvents (e.g., dibenzyl ether in iDISCO) with a commercialized LSFM should be taken into account, due to their corrosive effects such as dissolution of glues used in the construction of objective lenses (McKey et al., 2020). A proper optical resolution should also be set to meet the experimental and analytical objectives while avoiding oversampling. A high numerical aperture (NA) objective lens with a short working distance can also interfere with volume imaging. RI discrepancies between objective lens coverage and clearing reagents can reduce imaging quality due to spherical aberrations.

Whole imaging of a large kidney organoid that was generated from human iPS cells is depicted in Figure 1 (Takasato et al., 2016). The sample was cleared with CUBIC reagent (final RI = 1.52) and 3D data were collected at a voxel resolution of 2.5 μm³ to detect cell coordinates for subsequent cellular architecture analysis. Low magnification macro-objectives (NA ∼ 0.1) provide reasonable image quality and data size (15.5 GB for raw 16-bit TIFF data, 1.6 GB for processed 8-bit TIFF data) for this purpose. Oversampling microscopic images with excessive magnification and higher NA objectives will produce redundant data sets. For example, if an image is captured at 2.5 times the voxel resolution, the data size will be an order of magnitude larger (2.5³ = 15.625), thereby burdening data storage and subsequent image analysis.