To facilitate fine-scale phenotyping of whole specimens, we describe here a set of tissue fixation-embedding, detergent-clearing and staining protocols that can be used to transform excised organs and whole organisms into optically transparent samples within 1–2 weeks without compromising their cellular architecture or endogenous fluorescence. PACT (passive CLARITY technique) and PARS (perfusion-assisted agent release in situ) use tissue-hydrogel hybrids to stabilize tissue biomolecules during selective lipid extraction, resulting in enhanced clearing efficiency and sample integrity. Furthermore, the macromolecule permeability of PACT- and PARS-processed tissue hybrids supports the diffusion of immunolabels throughout intact tissue, whereas RIMS (refractive index matching solution) grants high-resolution imaging at depth by further reducing light scattering in cleared and uncleared samples alike. These methods are adaptable to difficult-to-image tissues, such as bone (PACT-deCAL), and to magnified single-cell visualization (ePACT). Together, these protocols and solutions enable phenotyping of subcellular components and tracing cellular connectivity in intact biological networks.
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We thank H. McBride, D.K. Newman and J. Allman for useful discussions on uses of tissue clearing across disciplines. We thank M. Brissova and A.C. Powers from Vanderbilt University for providing fixed human pancreas tissue used in Figure 1 and guidance with pancreatic markers and anatomy. This work was funded by grants to V.G.: the US National Institutes of Health (NIH) Director's New Innovator IDP20D017782-01; the NIH/National Institute on Aging (NIA) 1R01AG047664-01; the Beckman Institute for Optogenetics and CLARITY; the Pew Charitable Trust; and the Kimmel Foundation. Work in the Gradinaru Laboratory at Caltech is also funded by awards from the following (to V.G.): the NIH Brain Research through Advancing Innovative Neurotechnologies (BRAIN) 1U01NS090577; the NIH/National Institutes of Mental Health (NIMH) 1R21MH103824-01; the Human Frontiers in Science Program; the Mallinckrodt Foundation; the Gordon and Betty Moore Foundation through grant GBMF2809 to the Caltech Programmable Molecular Technology Initiative; the Michael J. Fox Foundation; Caltech-GIST; and the Caltech-City of Hope Biomedical Initiative. This work was also supported by grants to P.J.B. from the NIH (2 P50 GM082545-06; W.I. Sundquist, principal investigator) and gifts from the Gordon and Betty Moore Foundation and the Agouron Institute to support electron microscopy at Caltech; and by National Science Foundation (NSF) IIS-1253538 and DBI-1262547 grants to C.C.F. K.Y.C. and N.C.F. were supported by the NIH Predoctoral Training in Biology and Chemistry (2T32GM007616-36).
The authors declare no competing financial interests.
Integrated supplementary information
Supplementary Figure 1 Effects of bis-acrylamide crosslinker on clearing time and swelling of PACT-cleared sections.
(a) Representative images of the time course for PACT clearing of four 2 mm thick rat coronal brain slices, (displayed anterior to posterior, from left to right). Slices were embedded in either A4P0B0.05 or A4P0 and then cleared with 8% SDS-PBS (pH 7.5). The A4P0 slices were completely clear by 144 hours. Although some heavily myelinated brain sections seemed to resist clearing in A4P0B0.05-embedded sections initially, this effect did not persist, resulting in similar overall clearing time as slices embedded without bis-acrylamide. Likewise, tissue transparency was indistinguishable between conditions after their 48-hour incubation in RIMS. Unlike in 1 mm sections (Fig. 3), bis-acrylamide did limit tissue expansion in 2mm thick slices (A4P0: 38% total average linear expansion, A4P0B0.05: 28% average linear expansion). (b) RIMS formulation guide to optimize the RI to that of the cleared sample. RIMS formulated with 82% HistodenzTM (RI = 1.4655) should be broadly applicable to cleared brain tissue, while RIMS with a higher RI of 1.48-1.49 (RIMS-1.48, RIMS-1.49) is suggested for denser cleared tissue such as bone (Fig. 6).
The amount of protein lost while clearing was measured by performing a BCA on the clearing buffer, which was collected and replaced periodically while 1 mm tissue samples were undergoing PACT. A standard curve of BSA protein concentration in each of the four different clearing buffers was generated. Standard curves were fit with a third order polynomial and used to extrapolate all protein loss measurements. (a) A representative case, in which the absorbance in arbitrary units (a.u.) of standard solutions at 562 nm is plotted against known BSA concentrations in 8% SDS-PBS (pH 7.5). (b) Graphs show single-trial, representative protein loss measurements for each hydrogel condition in each clearing buffer. Protein content was measured at 12 hours into clearing, at 24 hours, and then every 24 hours until the samples were clear, and normalized to the initial weight of the slice. Experiments were performed in triplicates, representative single trials for each combination are shown. (c) Time to clear for 1 mm sections PACT-processed with all hydrogel embedding and clearing buffer combinations (n = 3 for A4P1, A4P2 and Unhybridized. n = 4 for all others).
(a-c) Representative images of thick section clearing with addition of CuSO4 or 0.2% SB compared to regular PACT. 0.5 mm and 1 mm coronal Thy1-YFP mouse brain sections are shown after A4P1 hydrogel polymerization (a) and during clearing with 8% SDS-BB (pH 8.5) and subsequent 24 hour incubation in RIMS (b and c for 0.5 mm and 1 mm, respectively). (d) The control, CuSO4, and 0.2% SB treated 0.5 mm slices from (a-b) were immunostained for parvalbumin (see Table 4) and then transferred to RIMS, degassed, and mounted. Shown are 500 μm thick maximum intensity projections of endogenous YFP (cyan) and parvalbumin (red) staining (top) as well as lipofuscin (white) autofluorescence (top and bottom). Red blood cell-derived (e.g. lipofuscin-like) autofluorescence was excited at 561 nm and collected between 562-606 nm. (e) Visualizing endogenous fluorescence and immunostaining deep within thick tissue. A 1 mm thick Thy1-YFP mouse brain coronal slice was treated with 0.2% SB, A4P1-embedded, cleared with 8% SDS-BB (pH 8.5), immunostained for parvalbumin (see Table 4), and then transferred to RIMS and mounted. Endogenous YFP (cyan) and immunolabeled PV (red) were imaged throughout the slice (left) in a region of the cortex. A 100 μm thick maximum intensity projection (right) was taken at a depth of 500 μm to show representative imaging in the middle of the section. Signal range of the red channel was adjusted for better visualization of PV staining at depth. All sections were imaged on a Zeiss LSM 780 confocal with the Plan-Apochromat 10× 0.45 N.A. M27 air objective (w.d. 2.0 mm).
(a) Fluorescence image of Thy1-YFP expression prior to expansion-clearing. A 70 μm thick maximum intensity projection of five cells expressing YFP represents the standard for imaging pre-expansion. A bright-field image of the pre-expansion 100 μm brain slice is shown in the top right, with the location of the cells being imaged indicated by the red arrowhead. Noteworthy features that may differ between pre- and post- expansion-cleared tissue, such as cell bodies, branching processes, and large projections, are numbered 1, 2, and 3, respectively. (b) Fluorescence image of Thy-YFP expression after 4× expansion-clearing. A 340 μm thick maximum intensity projection of the same five cells in (a) is shown, with the same features labeled again 1, 2, and 3. Of note, a cell body (1) and the neuronal processes of an adjacent cell (2) are both partially obstructed by tissue lipids in (a), but can be easily identified in (b) after clearing and expansion. However, the 4× expansion that contributes to this increased visibility through tissue also causes some tissue destruction, as apparent in the multiple severed processes, such as (3). A bright-field image of the expanded slice embedded in agarose is shown at the top right. (c) Native YFP fluorescence from the same cell in pre-expanded (blue box) and post-expanded (yellow box) tissue is shown. (d) Equipment for sample processing: (1) 2% bis-acrylamide, (2) 40% acrylamide, (3) sodium acrylate, (4) ammonium persulfate (APS), (5) N,N,N’,N’-Tetramethylethylenediamine (TEMED), (6) 4-hydroxy TEMPO, (7) 20% SDS, (8) collagenase, (9) low gelling temperature agarose, (10) Entellan, and assorted, unlabeled, glass slides, spacers, and plastic dishes. All images were taken on a Zeiss LSM 780 confocal with the Plan-Apochromat 10× 0.45 N.A. M27 air objective (w.d. 2.0 mm).
(a) Mice were perfusion-fixed, A4P0-embedded, PARS-cleared for 5 days with 8% SDS-BB (pH 8.5), and washed with 1× PBS at pH 7.5. Numbers correspond to the extracted organs in panel (b). (b) Extracted organs from the cleared mouse in panel (a), pictured before (top) and after (bottom) RIMS incubation for 3 days. Black arrowheads correspond to the adrenal gland on the kidney and to the ovaries on the fallopian tubes. Each square represents 0.5 cm2. Rodent husbandry and euthanasia conformed to all relevant governmental and institutional regulations; animal protocols were approved by the Institutional Animal Care and Use Committee (IACUC) and by the Office of Laboratory Animal Resources at the California Institute of Technology.
(a) For labeling thick tissue sections, camelid nanobodies are a promising alternative to traditional antibodies, either full immunoglobulins or their engineered formats (single-chain variable fragment (scFv), Fab, and F(ab’)2). A possible workflow for nanobody production consists of: inoculating 500 ml terrific broth with 5 ml overnight cultures and grow at 37 °C until IPTG induction at OD = 0.5, then lowering the temperature to 20 °C for 10 hours for nanobody expression. Cell pellets are then lysed, carried through alternating cycles of freeze-thaw with benzonase addition, followed by a final addition of 0.1% polyethyleneimine to the pellet lysate before pelleting debris and filtering the nanobody-containing fraction. The His-tagged GFAP fusion protein is purified by immobilized metal affinity chromatography on a Ni-NTA column. The His-tag must be removed prior to staining to avoid non-specific binding. (b) To stain for glial fibrillary acidic protein (GFAP) using a GFAP camelid nanobody, 1 mm thick PACT-cleared mouse brain sections were immunostained with 1:500 Atto 488 conjugated GFAP nanobody (see Table 4) at RT overnight with shaking. The stained sections were then washed 3 times in PBST over 1 hour, followed by a 1-hour incubation in RIMS. The transparent sections were RIMS-mounted and imaged on a Zeiss LSM 780 confocal with the Plan-Apochromat 10× 0.45 N.A. M27 air objective (w.d. 2.0 mm). (b, left) 850 μm thick 3D rendering of mouse internal capsule stained with GFAP nanobody. (b, right) Side view showing uniform labeling of GFAP nanobody throughout the entire 850 μm slice.
(a) neuTube. (b) Imaris. Computer screenshots depict the image processing workspace for each software during the 3D visualization (i) of labeled cells in Figure 10.
Supplementary Figures 1–7, Supplementary Methods and Supplementary Table 1 (PDF 8493 kb)
Zip file containing STL design file required for custom printing of Immersion chamber for LSFM. (ZIP 62 kb)
Zip file containing STL design file required for custom printing of Sample holder for LSFM. (ZIP 17 kb)
Zip file containing source file used to test the tracing functionalities of neuTube and Imaris. Picturing two labeled neurons in the mouse striatum across two fields of view, the test image, provided in TIFF file format, was acquired on an LSM 780 at 25× magnification and stitched in Zen (Zeiss) to produce a single channel, 8-bit, 300 MB image stack of size 3.15 × 108 voxels (1024 × 2048 × 150) covering approximately 0.08 mm3 (480 × 960 × 175 um3) of tissue. 3D image analysis of this test image generated the neuTube and Imaris traces depicted in Figure 10 and Figure S7. (ZIP 141502 kb)
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Treweek, J., Chan, K., Flytzanis, N. et al. Whole-body tissue stabilization and selective extractions via tissue-hydrogel hybrids for high-resolution intact circuit mapping and phenotyping. Nat Protoc 10, 1860–1896 (2015). https://doi.org/10.1038/nprot.2015.122
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