Comparing Established Organic and Water-Based Protocols for Clearing Cochleae

To compare established tissue clearing protocols for mouse cochleae, we selected three organic solvent-based (i/cDISCO, ECi, and PEGASOS [9, 15,16,17]) and two water-based protocols (ScaleS and FRUIT [20, 21]). We evaluated each protocol for transparency, immunostaining continuity, signal-to-background quality, tissue morphology, and reliability (Fig. 1 and Table 1). Transparency was assessed visually with cold white light illumination in a stereo microscope based on the visibility of blood vessels, stria vascularis pigmentation, and residual bone (Fig. 1a, row 2 and graphs). For comparing immunostaining continuity and fluorescence intensity after clearing, we immunostained the cochleae for myosin VIIa, a well-characterized marker for inner and outer hair cells in the organ of Corti. The immunohistochemistry outcome was characterized as follows: Immunostaining continuity was determined by evaluating the presence of specific immunofluorescence as well as its homogeneity from apex to base in reconstituted 3D images (Fig. 1 and Supplementary Fig S4a). In addition, we qualitatively assessed the strength of the specific fluorescence signal in relation to unspecific background fluorescence. Morphological preservation was evaluated qualitatively based on visible tissue alterations. We also monitored the reproducibility of the clearing protocols by evaluating whether the best result was always or only occasionally achieved in these categories by independent experiments, and categorized this as the “reliability” of each protocol.

Fig. 1

Comparison of cochlear transparency and immunofluorescence across different clearing methods. a Representative images of cochleae from 2–3 month old mice cleared with i/cDISCO (n = 10 independent experiments), ECi (n = 10), PEGASOS (n = 10), ScaleS (n = 8), and FRUIT (n = 4). Top row: cochleae after EDTA incubation. Middle row: cochleae after clearing, imaged under a stereo microscope with top illumination. Bottom row: assessment of optical transparency using white cold light illumination. Representative transmitted-light line profiles are shown to visualize relative changes in transparency across each sample. Lower values indicate reduced transparency due to residual absorption or scattering. Blue dotted lines indicate where the profile was acquired. b, top row: 3D light-sheet microscopy reconstruction of cleared cochleae immunolabeled for myosin VIIa to visualize inner and outer hair cells; scale bar: 300 µm. Bottom row: Higher-magnification views of the apical region of the organ of Corti showing inner and outer hair cells; views were rotated for display to allow easier comparison across protocols. Scale bar: 100 µm

Table 1 Qualitative evaluation of five clearing protocols

Of the organic solvent-based protocols, i/cDISCO resulted in outstanding transparency with almost invisible bone, stria vascularis and blood vessels (Fig. 1a, middle row). Prolonged incubation in DBE for >1 week removed the remaining pigments in the stria vascularis, rendering the cochlea completely transparent. In ECi-cleared cochleae, the bone appeared transparent, while pigments in stria vascularis and blood vessels remained visible (Fig. 1a, middle row). Among the organic solvent-based protocols, PEGASOS was least efficient in clearing the cochlea, leaving the bony shelf behind the apex still visible (Fig. 1a, middle row; Table 1).

To complement the visual assessment of cochlear transparency, we performed line profile-based analysis of brightfield images to visualize relative changes in transmitted light across the cleared tissue (Fig. 1a, bottom row). For each cochlea, grayscale intensity profiles were extracted along a straight line spanning the entire width of the sample from inverted brightfield images acquired under identical imaging conditions (Fig. 1a and Supplementary Fig. S3a). Line profiles confirmed that i/cDISCO-cleared cochleae exhibited relatively uniform transmitted light intensity across the selected tissue region, consistent with low residual absorption and scattering. By contrast, PEGASOS-, ScaleS-, and FRUIT-cleared cochleae showed more pronounced local decreases in transmitted-light intensity along the selected line profiles, reflecting some residual light absorption by bone and pigmented structures.

All organic solvent-based protocols facilitated homogeneous staining and preserved immunofluorescence in a comparable manner, resulting in efficient labeling of all myosin VIIa-expressing cells in the organ of Corti (Fig. 1b). While ECi only occasionally resulted in incomplete antibody penetration, this occurred more frequently in i/cDISCO and PEGASOS, with near absence of specific labelling in some experiments. i/cDISCO and ECi led to the brightest and most homogeneous staining of hair cells from apex to base (Fig. 1b and Supplementary Fig. S4a). PEGASOS cleared samples displayed highly variable immunostaining signals along the cochlear axis over the course of several experimental repetitions. In both the PEGASOS and the i/cDISCO protocol, in some cases unspecific fluorescence was observed. Particularly, prolonged incubation in DBE in i/cDISCO increased unspecific fluorescence in the stria vascularis, which got even stronger than the specific antibody staining over time (Supplementary Fig. S4 a).

The morphology of the organ of Corti was preserved in i/cDISCO and ECi protocols without obvious distortions, while PEGASOS often caused deformation of the organ of Corti structure, resulting in wavy hair cell rows (Supplementary Fig. S4a). Notably, the ECi protocol demonstrated the greatest reliability in terms of specific antibody labelling, staining continuity throughout the sample, and signal-to-background quality between experiments. Further, specific fluorescence intensity and low background were both stable for up to several weeks in ECi solution.

One advantage of water-based clearing protocols is that they better preserve tissue morphology by avoiding dehydration-induced shrinkage. Particularly in organs with complex mixtures of soft and hard tissues such as the cochlea, water-based strategies might thus be preferable to avoid shrinkage artifacts reported previously. To test water-based techniques, we chose ScaleS and FRUIT [20, 21]. ScaleS was selected because it had previously been adapted for cochlear clearing, whereas FRUIT was included as an additional hydrophilic, fructose-based protocol using comparatively simple and non-toxic reagents. Both water-based approaches resulted in even lower levels of transparency than PEGASOS, the worst performing organic solvent-based protocol. ScaleS and FRUIT both failed to render bone, stria vascularis and blood vessels transparent (Fig. 1a, row 2). The FRUIT solutions were further challenging to handle due to their high viscosity and tended to become turbid within a few days, which limited imaging quality.

Immunostaining continuity, immunofluorescence signals and signal-to-background quality appeared variable in repeated experiments using both protocols. In more than half of the experimental trials, specific immunofluorescence was absent after ScaleS clearing, and only rarely presented strong staining from apex to base (Fig. 1 b and Supplementary Fig. S4a).

Thus, compared to organic solvent-based protocols, the water-based clearing methods did not reduce light scattering to the same extent, resulting in more pronounced blurry regions in the images. Additionally, we observed more unspecific antibody binding and autofluorescence of the cochlear bony shelf in water-based protocols. Similar as in i/cDISCO and ECi, we did not observe morphological alterations in ScaleS and FRUIT experiments (Fig. 1b and Table 1). Notably, in both ScaleS and FRUIT, the agarose did not remain as a solid block, which made handling and mounting for imaging difficult.

In summary, i/cDISCO and ECi achieved the best transparency combined with good immunostaining continuity, signal strength and morphological preservation (Table 1), at the expense of long protocol durations.

Optimization of Pre-processing Steps for Accelerated Tissue Clearing

To accelerate the clearing process, we reduced the time required for decalcification, decolorization and refractive index matching. One key step is to remove the rather thick and pigmented part of the bony shelf at the back of the cochlea which connects to the bulla (Fig. 2a + b and Supplementary Fig. S2). Removal of this bone at the apical part of the cochlea allows a drastic reduction of the decalcification and decolorization steps and is crucially done before fixation and decalcification to avoid breaking of the cochlea. Subsequently, the fixation is performed as described, involving the diffusion of solutions through a hole in the apical cochlea and punctured oval and round window membranes.

Fig. 2

Cochlear trimming to allow faster clearing. a Cochlea after dissection from the skull. The white arrows indicate the highly pigmented bone connecting the cochlea with the bulla. b Cochlea after removal of the pigmented bony shelf. The white dotted line gives an impression of how big the hole at the cochlear apex should be for the fixation solution to enter by diffusion. The yellow arrows indicate where the oval and round window are gently punctured to ensure diffusion of solutions. c, d White arrows depict where extra removal of the bony shelf at the front (c) and back (d) of the cochlea is recommended to further shorten the decalcification time. After carefully separating the excessive bone and vestibular organ with sharp forceps (dotted line), remaining blood vessels (black arrows) should be detached. e, f Fully trimmed cochlea for one-day calcification and fast clearing

We next optimized decalcification conditions by varying EDTA concentration, temperature and incubation time, using tissue handling, macroscopic bone softening and subsequent light-sheet imaging quality as practical endpoints. Increasing the EDTA concentration from 200 to 500 mM did not result in an obvious improvement in bone softening or subsequent clearing. In contrast, increasing the temperature during EDTA incubation strongly accelerated bone softening. At 4 °C, decalcification proceeded slowly and required several days to complete. We next compared 37 °C to room temperature (RT) and found that incubation at 37 °C for three days enabled sufficient decalcification for clearing and imaging. Prolonging the incubation in EDTA to 14 days did not further improve clearing results. Based on these observations, all subsequent experiments were performed using 200 mM EDTA at 37 °C.

To further shorten the decalcification time, we removed additional bone structures. This advanced trimming step requires more technical skills but can be omitted if maximal morphological preservation is prioritized over protocol speed. After a short EDTA pre-incubation at 37 °C (Supplementary Table S2), we removed approximately two thirds of the posterior bony shelf of the cochlea while aiming to preserve the underlying soft tissues (Fig. 2c + d). This optional pre-processing routine reduced the time required for sufficient decalcification from three days to 24 h. We further trimmed the semicircular canals and surrounding tissue including blood vessels (both for five- and seven-day protocols, Fig. 2d–f).

Shortening Whole-Mount Antibody Staining to Accelerate the Protocol

To accelerate the two best-performing clearing protocols, i/cDISCO and ECi, we systematically reduced individual protocol steps (Fig. 3a) and evaluated the impact on transparency, antibody staining and morphology in 3D light-sheet reconstructions.

Fig. 3

Schematic overview of original-length and fast cochlear clearing protocols and comparison of resulting transparency. a: Schematic depiction of the individual processing steps and step durations for the original-length protocols (i/cDISCO: appr. 44 days; ECi: appr. 26 days) and the optimized fast clearing protocols (fastDISCO and fastECi: 5 days). b, top row: cochleae from 2–3 month old mice prepared with the trimming strategies used for original-length and fast protocols. Middle row: the same cochleae after clearing with original and fast protocols, illustrating differences in optical transparency (see also Supplementary Fig. S3). The dashed black line outlines the i/cDISCO-cleared cochlea. Arrows in the ECi condition mark bone and blood vessels that remain visible after clearing. The blue dotted lines indicate the approximate position used for transparency assessment. Bottom row: line profile analysis of transparency (see also Supplementary Fig. S3)

For i/cDISCO, we reduced the fixation time in 4% PFA from overnight to 3 h, which was sufficient to preserve the overall cochlear structure for downstream clearing and light-sheet imaging under the conditions tested. The decolorization step using Quadrol, which reduces autofluorescence and absorption from heme [2, 9], was shortened from 2 days to 2 h. Under our experimental conditions, this shorter incubation reduced residual absorption to a level compatible with light-sheet imaging of adult mouse cochleae. Although residual pigmentation of the stria vascularis was occasionally observed after fastDISCO clearing, it did not obviously prevent visualization of the organ of Corti in our datasets (Fig. 3b, middle row). Extended storage of cleared cochleae in DBE for >1 week further reduced the remaining pigmentation in the stria vascularis, rendering the cochlea completely transparent. However, this seemed to correlate with increasing autofluorescence of the stria, similarly as described for the original length i/cDISCO (see Supplementary Fig. S4 a and S5 a).

The original iDISCO protocol includes optional methanol-free pre-treatment taking approximately 130 h [15]. The cDISCO protocol recommends prolonged antibody incubation of 14 days for each primary and secondary antibodies [16]. This, together, accounts for the long duration of the combined i/cDISCO protocol. Omission of the methanol-free pre-treatment resulted in a markedly reduced immunofluorescence signal intensity. Consequently, we retained the original pre-treatment but shortened individual incubation steps, allowing completion within one day while still producing sufficient staining quality for volumetric light-sheet imaging under the conditions tested. In addition, antibody incubation times were reduced to overnight incubation for each step, which still resulted in continuous labelling of the organ of Corti in the optimized fast protocols (Fig. 4). To further assess whether shortened antibody incubation affects staining uniformity along the cochlear spiral, we assessed fluorescence intensity distributions along the length of the organ of Corti. For representative cochleae, three-dimensional light-sheet datasets were rotated into a standardized top-view orientation, and fluorescence intensity profiles were extracted along the hair cell row from apex to base using a manually traced polyline. Intensity values were normalized to cochlear length (0 = apex, 1 = base) to allow direct visualization of spatial signal distribution. These apex-to-base intensity profiles demonstrated staining continuity along the organ of Corti in both fastDISCO (Fig. 4b, left) and fastECi protocols (Fig. 4b, right).

Fig. 4

Myosin VIIa immunolabeling along the organ of Corti in original-length and fast clearing protocols. In representative cochleae from 2–3 month old (i/cDISCO, ECi, fastDISCO) or 7 month (fastECi) old mice, fluorescence intensity was extracted along a manually defined polyline tracing the hair cell row from apex to base and visualized using a color-coded profile (apex: yellow to base: purple). The same color coding is overlaid onto the top-view projections to enable spatial mapping of signal intensity to anatomical regions along the cochlear spiral. Cochleae were imaged with the apical side facing the detection objective, resulting in a gradual decrease in signal intensity toward the basal region due to light propagation through the tissue. a Original-length protocols. Left: i/cDISCO-cleared cochlea showing continuous myosin VIIa signal along the entire apical-to-basal extent of the organ of Corti. Right: ECi-cleared cochlea illustrating a representative example with local differences in the fluorescence signal along the cochlear spiral. b Fast clearing protocols. Left: fastDISCO-cleared cochlea, in this example with some unspecific fluorescence in the modiolus. Right: fastECi-cleared cochlea. In both cases, myosin VIIa signal is rather homogenously labelling hair cells along the full length of the organ of Corti. Scale bar: 300 µm

After immunolabelling, methanol was replaced by ethanol for dehydration without negatively affecting clearing performance. With these adjustments and the advanced trimming of the cochlear bone, the total duration of the i/cDISCO protocol was reduced to five days (fastDISCO). The protocol with simplified trimming requiring three days of decalcification takes seven days in total (see Supplementary Fig. S2).

In the original ECi clearing protocol, decalcification represented the most time-consuming step. As shown above, 24-h incubation in 200 mM EDTA at 37 °C turned out to be sufficient for decalcification for the extensively trimmed cochlea. We further adjusted downstream steps by shortening blocking and antibody incubation as we did for fastDISCO. Dehydration was performed as in fastDISCO using a graded ethanol series with one hour incubation time for each step. Again, the thoroughly trimmed cochlea requires a total protocol duration of five days for complete clearing, the less trimmed cochlea can be cleared within seven days with fastECi (Fig. 3 and Suppl. Fig. S2, S6b).

Validation of Morphological Preservation and Immunofluorescence of Cochlear Cells in Fast Protocols

To validate preservation of epitopes and morphology in the fast clearing protocols, we tested a panel of antibodies targeting key cochlear cell types. For hair cell labeling, vesicular glutamate transporter 3 (Vglut3), calretinin (Calb2) and parvalbumin were used as inner hair cell (IHC) markers, while oncomodulin reliably labeled outer hair cells (OHC). Myosin VIIa was used to robustly label both IHCs and OHCs (Figs. 4 and 5). In total, we combined three different markers per sample (Fig. 5 and Suppl. Fig. S5). In addition, SYTOX™ Green was used as a nuclear counterstain to support visualization of nuclei and tissue organization during image analysis. (for further details on antibody stainings see Supplementary Table S1).

Fig. 5

Immunolabeling of different hair cell markers in fast clearing protocols. Representative fastDISCO- (a) and fastECi-cleared (b) cochleae labeled with multiple hair cell markers. Inner hair cells are visualized using Vglut3, calretinin and parvalbumin, while outer hair cells are labeled using oncomodulin; myosin VIIa labels both inner and outer hair cells. Representative examples are shown from independent cochleae stained with different hair cell marker combinations: fastDISCO, n = 17; fastECi, n = 20; age of animals displaying calretinin immunolabelling: 1 month; other examples: 2–3 months. Scale bar in left panels: 300 µm. Scale in middle and right panels: 100 µm

Both fastDISCO and fastECi resulted in homogeneous staining along the entire apical-to-basal axis of the organ of Corti for all markers, allowing reliable detection of individual hair cells, but also their loss (Figs. 4a, b and 5a, b; Suppl. Fig. S5 and Fig. 7b).

Spiral ganglion neurons (SGNs) and associated nerve fibers were examined using antibodies against tubulin beta-III (TUBB3), calretinin and parvalbumin. TUBB3 labeled neurites of all SGN types, whereas calretinin and parvalbumin selectively labeled type I SGNs, which can be further divided into subtypes Ia, Ib and Ic [22]. Using calretinin and parvalbumin immunostaining, differentiation between SGN subtypes Ia, Ib and Ic was possible in fast-cleared samples (Fig. 6, Supplementary Movie 1). Notably, even in a sample with partial rupture at the basal SGN region, SGN-associated labelling remained detectable on both sides of the rupture (black rift between SGNs, Supplementary Fig. S5 b, Supplementary Movie 1). Yet, unspecific staining of SGNs was detected in some cochleae after fastDISCO clearing (Fig. 4b, left).

Fig. 6

Spiral ganglion neuron (SGN) immunostaining in fastECi. Representative fastECi-cleared cochleae immunolabeled for SGN subtypes. Anti-tubulin β-III (TUBB3) labels all SGNs, while calretinin and parvalbumin selectively label type I SGN subtypes. Calretinin shows strongest labeling of type Ia SGNs, parvalbumin labels type Ic SGNs, and type Ib SGNs are labelled with both markers. Upper panels: merged images. Lower panels: higher-magnification views of the indicated regions, illustrating distinguishable SGN subtypes; dotted lines outline the respective SGN subtypes. Representative example of n = 20 independent fastECi-cleared cochleae. Scale bar: 50 µm

In both, fast and original DISCO-based protocols, we occasionally observed autofluorescence in the stria vascularis, predominantly in channels corresponding to the excitation wavelengths of 647 nm and 790 nm. Particularly with the latter, we noticed an unspecific stria vascularis (auto-)fluorescence in almost all fastDISCO or i/cDISCO cleared cochleae as soon as samples were stored in DBE solution for more than 1–2 weeks (Supplementary Fig. S4a and S5a). However, when imaging was performed within one week after clearing, no such autofluorescence was detected.

To assess a potential improvement of stria vascularis transparency by attempting to wash out remaining blood from the vessels prior to clearing, we tested cochleae from transcardially perfused animals and compared these to non-perfused cochleae after clearing with the fastECi protocol (Supplementary Fig. S7). Since we observed no obvious difference in terms of tissue transparency, we assume that transcardial perfusion does not lead to substantial removal of blood from tiny blood vessels in the stria vascularis and is thus likely not superior to our standard procedure without transcardial perfusion.

Overall, morphological distortions, like wavy appearance of the basilar membrane, were not observed in fast protocols, similarly to original-length protocols, indicating that protocol shortening does not compromise tissue integrity.

Fast Clearing Protocols Support Hair Cell Row Segmentation and Hair Cell Counting

To assess whether the optimized fast clearing protocols support automated, object-based quantitative analysis, we performed three-dimensional segmentation of cochlear hair cells in a representative fastECi-cleared cochlea. Segmentation was based on combined nuclear counterstaining and immunolabeling for Vglut3 and oncomudulin, enabling identification of individual inner and outer hair cells within the intact organ of Corti (Fig. 7a–c).

Fig. 7

Segmentation of cochlear hair cell rows and cell counting is supported by fast clearing protocols. a Three-dimensional light-sheet microscopy reconstructions of a representative fastECi cleared cochlea immunolabeled for nuclei (green), Oncomodulin (magenta) and Vglut3 (yellow). A surface reconstruction of the cochlea is shown in purple. Selected z-planes (right panel) illustrate preservation of tissue architecture and signal quality throughout the sample depth. b Segmentation of inner and outer hair cell rows based on hair cell markers. The IHC row extends beyond the OHC marker signal, consistent with focal OHC loss in aged mice. c Higher-magnification view of hair cell detection using spot segmentation. Apical regions (top) show regular inner and outer hair cell rows while basal regions (bottom) show pronounced hair cell loss in a 9-month old C57BL/6 mouse. Representative segmentation example of n = 5 segmented fastECi-cleared cochleae

The resulting segmentation allows visualization of hair cell distributions throughout the cochlear spiral and enabled extraction of quantitative readouts, such as object counts along the apical-to-basal axis. While the present analysis is based on a single representative dataset and is not intended for statistical comparison, it demonstrates that the signal quality and structural preservation achieved with the fast clearing workflows are sufficient to support automated, annotation-free segmentation and quantitative analysis of cochlear cell populations.

Comparison and Overall Outcome of Original and Fast Protocols

In total, we systematically compared five established clearing protocols and optimized the two best-performing protocols. To enable an integrated comparison of all seven protocols, we assessed multiple attributes required for effective cochlear clearing and complemented these with criteria reflecting practical laboratory feasibility (e.g., toxicity of reagents or protocol duration). Seven categories were defined and scored on a scale from zero (lowest) to four (highest). Notably, we weighed all categories equally (see Table 2).

Table 2 Qualitative evaluation of seven clearing protocols

FRUIT, ScaleS, and PEGASOS achieved lower overall scores, although for different reasons. These included variability between clearing trials, morphological alterations, inconsistent immunofluorescence labeling or limited solution stability. In contrast, the original i/cDISCO and ECi protocols performed well across most criteria, with protocol duration representing their primary constraint. This limitation was effectively addressed by the optimization resulting in the respective fast protocols. Both fastDISCO and fastECi retained the key advantages of their original counterparts while substantially reducing protocol duration. Among all evaluated approaches, fastECi showed superior performance in terms of staining continuity, signal-to-background quality and reliability, resulting in the highest overall score and identifying fastECi as the most suitable protocol for routine cochlea tissue clearing.