Animals Wild-type mice (C57BL/6N and FVB/N, both males and females), Thy1-YFP mice (line H), and Th-cre (1Tmd/J) mice were used in the development and testing of novel clearing protocols and clearing reagents. Thy1-YFP mice were used to evaluate the maintenance of endogenous fluorescent signals throughout multi-week clearing steps and under long-term sample storage in RIMS. Periadolescent through adult wild-type rats (Long-Evans and Wistar, males and females) were used to optimize clearing protocols for larger tissue samples, and to depict the preservation of vasculature during lengthy perfusion-based clearing and antibody staining steps. For transcardial perfusion, subjects were deeply anesthetized with an overdose of Euthasol (100 mg/kg IP injection) prior to intracardiac perfusion first with heparinized PBS (10 U/ml heparin in 0.1 M PBS) containing 0.5% NaNO 2 and then with 4% PFA. For PACT, the brain and/or desired organs were excised and postfixed in 4% PFA for several hours prior to hydrogel monomer infusion and clearing steps. For PARS-based whole-body clearing, the intracardiac catheter was inserted into the left ventricle extending just beyond the aortic valve, stabilized inside the aorta with a loose loop of suture thread. For PARS-based whole-brain clearing, the descending aorta was ligated with a microclamp. For experiments involving the visualization of AAV9-CAG-eGFP transduced cells, young adult female C57Bl/6 mice were injected with virus via the retro-orbital sinus, and following a 6 month delay for viral transduction and eGFP expression, mice were euthanized for PACT and PARS studies.

Ethics Statement Animal husbandry and all experimental procedures involving mice and rats 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.

AAV Production and Systemic Delivery Lock et al., 2010 Lock M.

Alvira M.

Vandenberghe L.H.

Samanta A.

Toelen J.

Debyser Z.

Wilson J.M. Rapid, simple, and versatile manufacturing of recombinant adeno-associated viral vectors at scale. Pulicherla et al., 2011 Pulicherla N.

Shen S.

Yadav S.

Debbink K.

Govindasamy L.

Agbandje-McKenna M.

Asokan A. Engineering liver-detargeted AAV9 vectors for cardiac and musculoskeletal gene transfer. Pulicherla et al., 2011 Pulicherla N.

Shen S.

Yadav S.

Debbink K.

Govindasamy L.

Agbandje-McKenna M.

Asokan A. Engineering liver-detargeted AAV9 vectors for cardiac and musculoskeletal gene transfer. 12 vector genomes (vg) of either virus was delivered intravenously into young adult female C57Bl/6 mice via the retro-orbital sinus and the mice were euthanized 6 months later for assessment of native eGFP fluorescence by PARS. All imaging of PARS brain and liver tissue from AAV9-injected mice was performed after 2 weeks tissue storage in RIMS. By injecting mice with adeno-associated viral vectors carrying fluorescently labeled transgenes, we were able to observe the compatibility of PARS processing and RIMS mounting with more sparse, localized fluorescent labeling than that which is driven by the Thy1 promoter ( Figures 5 A–5C versus Figures 5 D and 5E). Sparse labeling of specific neuron types and glia as well as localized eGFP expression in distinct organs (e.g., liver and hippocampus Figures 5 D and 5E) was clearly visible. Single stranded ssAAV-CAG-eGFP vectors packaged into AAV9 or the AAV9 variant capsid, AAV9BD1, was generated and purified as described (). The AAV9BD1 capsid was modified from AAV9 (U. Penn) with the following mutations (VP1 numbering): (1) An N498Y mutation was made to reduce liver transduction (), (2) the amino acid sequence AAADSPAHPS (Chen et al., 2009) was inserted between AA588-589, and (3) a Y731F mutation was made (). 1 × 10vector genomes (vg) of either virus was delivered intravenously into young adult female C57Bl/6 mice via the retro-orbital sinus and the mice were euthanized 6 months later for assessment of native eGFP fluorescence by PARS. All imaging of PARS brain and liver tissue from AAV9-injected mice was performed after 2 weeks tissue storage in RIMS.

Selection of PACT and PARS Reagents 2 and 10 U/ml heparin, and then 4% paraformaldehyde (PFA, in 0.1 M PBS, pH 7.5). The excised whole brains were sliced into 1 mm and 3 mm sagittal sections and coronal sections, postfixed in 4% PFA at room temperature for 2–6 hours (postfixing whole brain and sections at 4°C overnight is also a valid option), and then sections were incubated at 4°C overnight in A2P0 (2% acrylamide and 0% paraformaldehyde in PBS), A4P0 (4% acrylamide and 0% paraformaldehyde in PBS), or A4P4 (4% acrylamide and 4% paraformaldehyde in PBS) hydrogel monomer solution, each containing 0.25% photoinitiator 2,2′-Azobis[2-(2-imidazolin-2-yl)propane]dihydrochloride (VA-044, Wako Chemicals USA). While still submerged in hydrogel monomer, the hydrogel-infused samples were degassed by bubbling nitrogen through the sample-hydrogel solution in the vacutainer or 5 ml Eppendorf tubes for 1 min. It should be noted that we experimented with several, more rigorous methods of replacing oxygen atmosphere with an inert gas, as was deemed necessary in the original and advanced CLARITY protocols ( Chung et al., 2013 Chung K.

Wallace J.

Kim S.Y.

Kalyanasundaram S.

Andalman A.S.

Davidson T.J.

Mirzabekov J.J.

Zalocusky K.A.

Mattis J.

Denisin A.K.

et al. Structural and molecular interrogation of intact biological systems. Tomer et al., 2014 Tomer R.

Ye L.

Hsueh B.

Deisseroth K. Advanced CLARITY for rapid and high-resolution imaging of intact tissues. To screen different hydrogel monomer formulations and clearing conditions, several adult C57 and Thy-1 eYFP mice (Jackson) were anesthetized with an overdose of Euthasol (100 mg/kg, IP injection) and transcardially perfused first with PBS containing 0.5% NaNOand 10 U/ml heparin, and then 4% paraformaldehyde (PFA, in 0.1 M PBS, pH 7.5). The excised whole brains were sliced into 1 mm and 3 mm sagittal sections and coronal sections, postfixed in 4% PFA at room temperature for 2–6 hours (postfixing whole brain and sections at 4°C overnight is also a valid option), and then sections were incubated at 4°C overnight in A2P0 (2% acrylamide and 0% paraformaldehyde in PBS), A4P0 (4% acrylamide and 0% paraformaldehyde in PBS), or A4P4 (4% acrylamide and 4% paraformaldehyde in PBS) hydrogel monomer solution, each containing 0.25% photoinitiator 2,2′-Azobis[2-(2-imidazolin-2-yl)propane]dihydrochloride (VA-044, Wako Chemicals USA). While still submerged in hydrogel monomer, the hydrogel-infused samples were degassed by bubbling nitrogen through the sample-hydrogel solution in the vacutainer or 5 ml Eppendorf tubes for 1 min. It should be noted that we experimented with several, more rigorous methods of replacing oxygen atmosphere with an inert gas, as was deemed necessary in the original and advanced CLARITY protocols () (e.g., 1. Placing the vacutainer containing the sample on ice; 2. Degassing the vacutainer with the house vacuum line while gently vortexing for several minutes; 3. Removing the sample from ice and bubbling nitrogen through the hydrogel monomer solution for several minutes; 4. Repeating steps 1–3 several times). However, we found that the brief 1 min exchange of oxygen for nitrogen supported adequate polymerization: residual oxygen may have hampered the complete hybridization between tissue and acrylamide monomers; however, our tissue-hydrogel matrix was sufficient for preserving tissue architecture and protein content. To polymerize the hydrogel-tissue matrix, the samples were transferred to a 37°C waterbath or heating block and incubated for 2–3 hr at this elevated temperature. The polymerized samples were washed briefly with PBS (0.1 M PBS, pH 7.5; all PBS wash steps) to remove excess hydrogel, transferred to 50 ml conical tubes, and incubated for 2–5 days at 37°C with shaking in either PBS, 0.1% Triton X-100 in PBS, or a clearing solution: 4% SDS, 8% SDS, 20% SDS, or 10% Deoxycholate, all prepared in 0.1 M PBS, pH 7.5. Images of 3 mm brain sections were taken at 24 hr and 48 hr ( Figure 1 A) to show the trade-off between greater tissue swelling for tissue-hydrogel matrices prepared with low PFA concentrations, and slower tissue clearing for tissue-hydrogel matrices prepared with high PFA (and acrylamide) concentrations; the A4P0 hydrogel formulation was selected for general use in subsequent PACT and PARS experiments. A 72 hr incubation of brain sections in the 8% SDS clearing solution resulted in superior tissue clearing ( Figure S1 A), and so the 8% clearing solution was selected for subsequent PACT and PARS experiments. Regarding clearing time, this parameter must be optimized in a case-specific manner. 24 hr clearing may be sufficient for small tissue samples or highly porous tissue, while larger, highly myelinated, or dense tissue sections and whole organs may require > 96 hr. Care must be taken to not overclear the samples and also to check periodically for excessive swelling if samples are to be stored long term since swelling does contribute to hydrogel softening and disintegration in the long run, risking sample loss. This is accelerated by elevated temperature and mechanical stress during sample preparation and handling. Gentle treatment of the tissue-hydrogel samples and the addition of antimicrobial agents to incubation solutions allow the hydrogel to remain stable for up to two weeks. Also, we found that an additional round of tissue crosslinking with 1%–2% PFA, or of tissue-hydrogel re-polymerization after clearing, was beneficial to counteracting both tissue expansion in mounting media and tissue disintegration.

PACT Immunohistochemistry To immunostain PACT-processed tissue, cleared samples were washed with 4–5 changes of PBS over 1 day to remove residual SDS. Then, the samples were incubated with primary antibodies (1:200–400) in PBS containing 2% normal donkey serum, 0.1% Triton X-100 and 0.01% sodium azide at room temperature with shaking for 3–7 days. Six A4P0-polymerized and six A4P4-polymerized 3-mm sagittal sections were removed from these antibody incubations at 24 hr, 48 hr, and 72 hr in order to measure IgG penetration depth in cleared tissue (see Figure 1 B). For remaining sections, unbound primary antibody was removed via washing sections in 4–5 PBS buffer exchanges over the course of one day. Then, samples were incubated with secondary antibodies (Fab fragment secondary antibodies are preferred, 1:200–400, in PBS containing 2% normal donkey serum, 0.1% Triton X-100 and 0.01% sodium azide) at room temperature with shaking for 2–5 days. After washing with 4–5 changes of PBS over 1 day, the samples were incubated in RIMS solution (40 g of Sigma D2158 (Histodenz) in 30 ml of 0.02 M phosphate buffer with 0.01% sodium azide, pH to 7.5 with NaOH—which results in a final concentration of 88% Histodenz w/v) at room temperature until they became transparent. During long incubations (>4 days) of tissue in antibody, small-molecule stains, or RIMS, the solution was exchanged for fresh halfway through the incubation. As an extra precaution and to prevent bacterial growth, tissues may be transferred to fresh 50 ml conical tubes or staining jars with every buffer exchange. It is suggested that RIMS incubations and mounting be performed in a clean environment—either in a hood, or by decanting RIMS into a fresh conical over flame to minimize bacterial contamination. The primary antibodies used for passive staining were chicken anti-tyrosine hydroxylase (TH) IgY, chicken anti-glial fibrillary acidic protein (GFAP) IgY (Aves Labs, Tigard, OR), rabbit anti-ionized calcium-binding adaptor molecule 1 (Iba1) IgG (Biocare medical, Concord, CA), rabbit anti-integrin b4, b5 IgG, and rabbit anti-beta tubulin IgG (Santa Cruz Biotechnology, Dallas, Texas). An AlexaFluor 647-conjugated donkey anti-mouse IgG (Jackson ImmunoResearch, West Grove, PA) was used for the antibody penetration experiment ( Figure 1 B). Nissl staining was performed with NeuroTrace 530 / 615 Red Fluorescent Nissl Stain (1:50 in PBS, Life Technologies, Grand Island, NY; samples were incubated at RT overnight and then washed in PBS prior to mounting. For small molecule staining with acridine orange, samples were placed in a 100 μg/ml solution of acridine orange for 10 min, followed by washout in PBS for one hour. The tissues were then placed in RIMS solution for 4 hr prior to imaging. All steps were performed at room temperature.

PARS Chamber Design Gage et al., 2012 Gage G.J.

Kipke D.R.

Shain W. Whole animal perfusion fixation for rodents. Jonkers et al., 1984 Jonkers B.W.

Sterk J.C.

Wouterlood F.G. Transcardial perfusion fixation of the CNS by means of a compressed-air-driven device. To perform PARS, a simple apparatus was needed to perfuse and recirculate PACT reagents through the circulatory systems for several days-to-weeks. Using the traditional cardiac perfusion fixation technique () as a delivery method, we devised a PARS chamber that consisted of the following components: 1) a feeding needle catheter placed within the left ventricle of the subject, 2) a perfusate collection well (pipette box) to catch recirculating reagents that exit the vasculature through a lesioned right atrium, and 3) catheter tube that transfers recirculating reagents from the collection well back into subject vasculature via its passage through a peristaltic pump ( Figure S4 A). To confirm that this set-up was functional for whole-organism clearing, several different detergents including SDS at several different percentages, sodium lauryl sarcosine, and sodium deoxycholate, at various concentrations, were continuously perfused through whole mice and rats via the carotid artery for up to 2 weeks. As in PACT ( Figure S1 A), only SDS could effectively render tissue transparent for optical imaging. Likewise, when delivered using the PARS chamber set-up, 8% SDS could efficiently solvate lipids deep in tissue and accomplish uniform clearing of large tissue samples. Thus, the PARS chamber set-up was adopted for subsequent PARS experiments.

PARS Clearing and Staining For transcardial perfusion fixation of adult mice or rats, a feeding needle was inserted through the left ventricle and into the aorta, and loosely sutured in place to the vessel at the level of the aortic arch. Following perfusion with PBS and 4% PFA, as summarized for PACT, the fixed whole rodents were transferred into a custom-built perfusion chamber where the solutions inside the chamber are perfused into the rodent and recirculated via a peristaltic pump. The rodent was postfixed with 4% PFA through the same feeding needle into the aorta at a flow rate of 1 ml/min for 1–2 hr at room temperature. For clearing of rat brain and spinal cord, we systematically ligated the arterial circulation leaving the carotid arteries intact and removed tissue not directly perfused by these vessels. To prevent excess PFA-crosslinked polyacrylamide from occluding the vasculature, we first perfused PBS (0.1 M PBS, pH 7.5; all PBS wash steps) for 2 hr at RT to wash out the residual PFA and we infused 4% acrylamide (A4P0) in PBS at RT overnight. The next day, we again perfused PBS to remove any remaining PFA/acrylamide polymers/monomers in the vasculature. Before polymerization and without disconnecting perfusion lines, we placed the perfusion chamber into a ziplock bag and infused nitrogen gas into the perfusion chamber through a separate connection to degas the sample. The polymerization process was initiated by adding 200 ml of 0.25% VA-044 initiator with PBS and submerging the degassed perfusion chamber in a 37–42°C water bath for 2–3 hr. A lead weight was placed on top of the perfusion chamber to prevent it from tipping over. After polymerization, the solution was replaced with 8% SDS in 0.1 M PBS, pH 7.5 clearing buffer, and the mouse/rat was perfused for up to 2 weeks. For PARS IHC, the cleared mouse/rat was first perfused with 8 buffer changes of 200 ml PBS over a 2 day period to remove the residual SDS. Then, using the same antibody formulations described in the PACT protocol, a 3 day perfusion with a primary antibody cocktail, 1-day perfusion with PBS wash, a 3 day perfusion with the secondary antibody cocktail, and a 1 day PBS wash was conducted in order to stain the organs of the cleared mouse/rat.

PARS-CSF Methodology for Brain and Spinal Cord Clearing For applications restricted to brain and spinal cord mapping, we developed a within-skull PARS strategy that grants thorough clearing of the whole-brain and whole-spinal cord by direct infusion of hydrogel monomers and clearing reagents into the CSF via an intracranial brain shunt. Under specific circumstances (e.g., the pre-existing availability of a guide cannula in the subject from an in vivo pharmacological, neurobiological, or optogenetic study), PARS-CSF would permit whole-brain clearing and histology that is automatically optimized for the region near the existing cannula, and that requires less time and reagents as the equivalent whole-organ PACT and PARS procedures. 2 ) for two minutes ( Herein, we validated two routes for intracranial delivery of PACT reagents. To clear the spinal cord, a cannula may be lowered through the skull (by drilling a hole in the region of interest and using tweezers to create an opening in the dura), to the level of the subarachnoid space, directly above the dorsal inferior colliculus, (see Figure 3 A, right). The rat spinal cord sample (see Figure 3 B, right) could be cleared when PARS-CSF was conducted at elevated temperatures and for a longer period of clearing. To clear the whole brain, the cannula may be lowered through the skull, penetrating the dura, and placed in the region directly above the olfactory bulb (see Figure 3 B, left). The cannula (21G blunted needle BD) is cemented in-place on the skull surface using dental acrylic (C&B-Metabond, Parkell). The PARS procedure was then applied to this intracranial preparation: the catheter tubing was connected to the subdural cannula as opposed to the cardiac feeding tube, and all PACT reagents were infused at 1 ml/min using the same order and timeframe as in PARS. For whole-brain clearing, the subject was transcardial-perfusion fixed and decapitated, with only the head transferred to the PARS chamber and connected to the infusion lines ( Figure S4 A). The pipette box and catheter lines were prefilled with 4% acrylamide monomer solution (A4P0), the tubing was connected to the cannula, and A4P0 was intracranially infused at a 1 ml/min flow rate overnight at room temperature ( Figure S4 A, left). After flushing the brain of unbound PFA and acrylamide monomers (2 hr infusion of PBS), which was critical to ensure that the vasculature remained unpolymerized, the whole brain was degassed via transferring the PARS chamber into a ziplock bag and placing the chamber under an inert atmosphere (N) for two minutes ( Figure S4 A, right). The bagged-PARS chamber was then transferred to a 37–42°C water bath, and degassed PBS supplemented with the thermal initiator was infused through the brain for the entire 2–3 hr incubation. After formation of this whole-brain-hydrogel matrix, in-skull tissue clearing was accomplished via constant perfusion recirculation of 8% SDS through the cannula for 4 days, with the PARS chamber remaining in the 37–42°C water bath for the entire process. Finally, after extensive PBS washing (2–3 days), the catheter lines were disconnected, and the brain was removed, sectioned, and mounted in RIMS for imaging ( Figure 3 B). Using mice that were IV-injected with AAV9-eGFP, the PARS-CSF procedure for whole-organ clearing was validated with respect to the following conditions: (1) only limited bias in how well regions clear relative to the cannula placement, (2) no structural damage in regions near the cannula due excessive fluid pressure, either from too high flow rate or inadequate drainage of perfused liquids, causing high intracranial pressure, (3) preservation of subcellular structural morphology, and (4) good visualization of sparsely labeled cell populations and fluorescence. It may have particular relevance to scientists performing research that already involves the use of intracerebroventricular (ICV) cannulated mouse or rat subjects and that requires postmortem brain histology for each subject.

Antibody Penetration Schindelin et al., 2012 Schindelin J.

Arganda-Carreras I.

Frise E.

Kaynig V.

Longair M.

Pietzsch T.

Preibisch S.

Rueden C.

Saalfeld S.

Schmid B.

et al. Fiji: an open-source platform for biological-image analysis. Four transcardially perfused and 4% PFA postfixed adult mouse (4–12 weeks old) brains were cut into 2 mm sagittal slices, and these slices were PACT processed. Specifically, one half of each PFA-fixed brain was hybridized with A4P4 hydrogel, while the other half was hybridized in A4P0 hydrogel. All the samples were passively cleared with 8% SDS in PBS, as described in the PACT protocol, and the residual SDS was removed by PBS washing for 1 day. The samples were then incubated in primary antibody cocktails (donkey anti-mouse-IgG antibody, 1:200, in PBS containing 2% normal donkey serum, 0.1% Triton X-100 and 0.01% sodium azide) for a range of time-periods, spanning 24–72 hr. Samples were then washed with 4–5 buffer exchanges of 0.1 M PBS over 1 day and mounted in RIMS solution. Images were taken with a Zeiss LSM 780 confocal microscope using the W Plan-Apochromat 20×/1.0 DIC M27 (working distance 1.8 mm). The depth of antibody penetration ( Figure 1 B) was outlined on y-z projected images using Fiji () with Reslice and Z project plugins.

RIMS for PACT and PARS Samples As a final step in tissue preparation for imaging, sample mounting comprises immersing the tissue section in a medium that will help to align the refractive indices of the objective, lens immersion media, and tissue, which confers higher-resolution and -imaging depth. In order to circumvent the use of FocusClear, a prohibitively expensive reagent, in mounting our PACT and PARS samples, we elected to formulate our own mounting solution alternative. Based on the principles of tissue optical clearing, we rationally screened two major groups of chemicals (sugar alcohols and radiocontrast agents) that have the following desirable characteristics of an optical clearing agent: high water solubility, low viscosity, high density, low osmolarity, low autofluorescence, nonfluorophore quenching, biocompatible, low cost. We identified sorbitol, a sugar alcohol that is affordable and widely available. At 70-80% (w/v) solution, sorbitol can effectively clear 200 μm thick uncleared and up to 1 mm thick PACT-cleared brain sections with little quenching of fluorescence. The formulation (termed sRIMS, see below for description) was later optimized to include 0.02-0.05 M phosphate buffer (pH maintenance), 0.1% tween-20 (enhances tissue penetration), and sodium azide (preservative to inhibit bacterial growth). In addition to sugar alcohols, we also looked at radiocontrast agents, especially intravascularly delivered non-ionic iodinated contrast agents (also licensed for use as density gradient media) as their physical and chemical properties closely match that of an ideal refractive index matching media. Based on cost and availability, we tested iodixanol (Optiprep) and its monomer iohexol (Nycodenz, also available as Histodenz, a derivative of iohexol) and found iohexol to be superior than sorbitol in index matching much larger PARS-cleared samples. Next, we conducted a side-by-side comparison (see Figure S3A) between commonly used/commercially available mounting options: 80-90% glycerol and FocusClear, and our most promising mounting media candidates: sRIMS and RIMS, a mounting media optimized for our imaging set-up with standard confocal microscopy. RIMS solution was prepared via dissolving 40 g of Histodenz (Sigma D2158) in 0.02 M phosphate buffer with 0.01% sodium azide for a total volume of 30 ml, pH to 7.5 with NaOH, which results in a final concentration of 88% Histodenz (w/v) with RI = 1.46 (used throughout this work unless otherwise noted). Estimated cost to produce is $3/ml while FocusClear is $36/ml. We note that the refractive index (RI) of RIMS may be adjusted to match the specific tissue/imaging system: it is expected that the RIMS RI may range from 1.38 (30% Histodenz w/v) to 1.48 (95% Histodenz w/v) in order to obtain optimal sample resolution. Light transmittance in RIMS ( Figure S3 B) was measured with a Reichert AR200 Refractometer. For RIMS mounting, samples were first submerged in RIMS at room temperature until they become transparent. During this period, the cleared tissue initially shrinks for the first few hours (see Figure S3 D). Continued incubation in RIMS will lead to gradual tissue expansion over time until RIMS has fully penetrated the tissue (see Figures 4 C and S5 A); we observed that our largest samples (e.g., rat whole brain) became transparent within one week of RIMS-immersion, after which their expansion ceased. We were able to limit tissue expansion, however, by postfixing the cleared and stained samples in 4% PFA for 1–2 hr at room temperature (small samples) or up to overnight (large samples) before proceeding to RIMS incubation (see Figure 4 C, right box; Figure S5 A, lower right box). Although postfixing PARS tissue curtailed gradual tissue volume expansion, the additional crosslinking also precipitated a slight decrease in tissue transparency ( Figure S5 A). Fluorescence intensity, cell phenotyping, or resolvable depth of imaging were not adversely affected, however. sRIMS: A Cost-Effective Sorbitol-Based Alternative to RIMS 70% sorbitol (w/v) (Sigma S1876) in 0.02 M phosphate buffer with 0.01% sodium azide, pH to 7.5 with NaOH; net cost of $0.2/ml. While RIMS outperformed sRIMS in our hands in terms of resolvable imaging depth, sorbitol is a commonly available chemical across scientific laboratories, and thus offers a convenient, cost-effective and superior alternative to glycerol-based mounting solutions.

Vasculature Preservation Li et al., 2012 Li T.

Bourgeois J.P.

Celli S.

Glacial F.

Le Sourd A.M.

Mecheri S.

Weksler B.

Romero I.

Couraud P.O.

Rougeon F.

Lafaye P. Cell-penetrating anti-GFAP VHH and corresponding fluorescent fusion protein VHH-GFP spontaneously cross the blood-brain barrier and specifically recognize astrocytes: application to brain imaging. Rats were transcardially perfused with heparinized PBS, 4% PFA, and lastly additional PBS wash. Then, before any hydrogel monomer infusion or clearing as in PARS, the rat was perfused via its intra-aortic catheter with 100 ml Atto 488-conjugated anti-GFAP nanobody (1:100 in PBS) at room temperature overnight. The GFAP nanobody was prepared according to published methods (). It was conjugated to Atto 488 fluorescent dye prior to use. The brain was removed from the skull and incubated in 4% PFA at 4°C overnight to crosslink the nanobody. The brain was then cut into 1 mm coronal slices processed according to standard PACT protocols, including hydrogel monomer infusion, hybridization, and passive sample clearing in 8% SDS in 0.1 M PBS, pH 7.5 at 37°C for 3 days. Cleared samples were incubated in RIMS solution for one day and mounted in RIMS solution for imaging. Images were taken using Zeiss LSM 780 confocal microscope with LD LCI Plan-Apochromat 25×/0.8 Imm Corr DIC M27 multi-immersion objective. The above methods were repeated in mice for the labeling of vasculature with Alexa Fluor 647-conjugated anti-mouse IgG. Briefly, 4% PFA-fixed mice were transcardially perfused with Alexa Fluor 647-conjugated anti-mouse IgG overnight using the same intra-aortic catheter that was installed for 4% PFA fixation. The brain was excised, post-fixed in 4% PFA at 4°C overnight, sectioned into 1 mm coronal slices and PACT cleared in 8% SDS in 0.1 M PBS, pH 7.5 at 37°C for 3 days. Cleared samples were incubated in RIMS solution for one day and mounted in RIMS solution for imaging.

Sliced Tissue Expansion and Weight Gain Measurement PFA-fixed adult mouse (4–12 weeks old) brain was cut into six 1-mm-thick coronal slices. Slices from one half of the brain were stored in PBS, while slices from the other half of the brain were PACT-cleared for 4 days. Slices were weighed and imaged with a conventional camera before and after clearing. The size of the slices was outlined and calculated using Image J. The tissue expansion and weight gain were determined by calculating the change in size and weight of slices before and after clearing ( Figure 1 E), and normalizing them to the pre-PACT measurements.

Protein Loss Measurement The percentage of protein loss for each sample ( Figures 1 C and 4 E) was obtained by measuring the amount of total protein in the clearing solutions collected from PACT or PARS clearing with NanoDrop blanked with respective solutions, and normalized to the weight of the mouse (for PARS) or the slices (for PACT) before clearing.

Whole-Brain Tissue Morphology Preservation and Quantification To observe the effect of PARS processing and RIMS mounting on brain volume, PFA-fixed adult mouse brains were either immediately extracted (uncleared control) or PARS-processed; and then brains from these two groups were treated according to one of the following conditions: incubated in PBS for 1 day, incubated in PBS for 1 week, mounted in RIMS for 1 day, mounted in RIMS for 2 weeks, or postfixed and mounted in RIMS for 2 weeks. Cleared and uncleared brains from all conditions were then photographed to estimate their relative size change ( Figure 4 C), cut into sections to visualize clearing depth ( Figure S5 A), slide-mounted, and imaged via confocal microscopy in order to evaluate gross changes in tissue architecture (e.g., morphological deformations of major brain regions, structure integrity of ventricles and vasculature) ( Figure S5 B). The percentage of protein loss for each sample ( Figures 1 C and 4 E) was obtained by measuring via NanoDrop the protein concentration of clearing solution aliquots that were collected after PACT or PARS-processing of tissue. The net protein loss could then be estimated based on this concentration and the known total volume of clearing solution used during processing. The amount of protein lost by each sample was normalized to the weight of the mouse (for PARS) or the tissue slice (for PACT) before clearing so that protein losses across samples and across tissue processing conditions could be compared.

Single-Molecule RNA FISH Tissue samples were adhered to aminosilane-treated coverslips by dehydrating for 1 hr under light vacuum. Samples were permeabilized prior to hybridization according to the following protocol: First, samples were washed twice in 100% ethanol for 10 min at room temperature. Next samples were washed in 95% ethanol for 10 min at room temperature. Samples were then incubated in 70% ethanol for 2 hr at 4°C. After incubation, tissue was placed in a 0.5% sodium borohydride (w/v) 70% ethanol solution for 10 min at room temperature. Finally, the tissue was rehydrated with 3 washes of PBS. Hybridizations were performed overnight at 37°C in a hybridization buffer composed of 10% dextran sulfate (w/v, Sigma D8906), 10% formamide (v/v), 2× SSC containing 1 nM per each of 24 Alexa 594-labeled 20-mer oligo probes toward β-actin. The next day samples were washed in 30% Formamide 2× SSC at room temperature for 30 min followed by 4 washes with 2× SSC. After washing sample was mounted between two coverslips with Slowfade Gold + DAPI (Life S36938). Samples were imaged on a Nikon Ti Eclipse microscope with an Andor Ikon-M camera and a 60×/1.4NA Plan Apo λ objective with an additional 1.5× magnification. Images were acquired as z stacks with a 0.5 μm step size over 30 μm. Samples were excited by a 589 nm (SDL-589-XXXT), 532 nm (SDL-532-200TG) and 405 nm(SDL-405-LM-030) lasers manufactured by Shanghai Dream Laser. The smFISH images ( Figure 2 ) were analyzed using image analysis scripts written in MATLAB. To determine the average background of the sample, the images were median filtered using a 50 × 50 pixel kernel and the average pixel intensity of the center 200 × 200 pixel sub-image was used as the average background value of the image. The smFISH dots were found by applying a Laplacian of Gaussian filter, thresholding the image based on the average background value and comparing the resulting image with a dilated image to find local maxima. The error bars were calculated using the standard deviation of the resulting measurements.

Human Tissue Biopsy Preparation Human basal cell carcinoma skin tissue samples were obtained from patients undergoing excision of their cancers after appropriate informed consent and under approval of UCLA IRB #12-01195. Tissue samples were obtained from sections of tumors not necessary for diagnostic or margin control purposes and varied in size depending on the size of the original skin cancer. Biopsied tumor samples were processed utilizing the PACT methodology as described for rodent tissue, using 4% acrylamide solution (A4P0) to generate hydrogel support matrix for fixed tissue. Polymerized tissue-hydrogel matrices were then passively cleared for 2–7 days (i.e., depending on tissue thickness) in 8% SDS at 37°C. In general, a 3-mm-thick human skin section could be rendered transparent within 3–4 days. PACT-processed samples were immunolabeled with anti-pan-cytokeratin (AE1/AE3) Alexa Fluor 488 primary antibodies (eBiosciences) at 1:100 dilution for two days followed by a 1 day wash in PBS. All labeling and wash steps were performed at room temperature, and final PACT-processed biopsy samples were mounted in RIMS. Imaging was performed on a Zeiss 780 confocal microscope with a LD LCI Plan-Apochromat 25×/0.8 N.A. multi-immersion objective. Despite testing a range of clearing times (2–7 days in 8% SDS) for tissue-hydrogel samples, the subcutaneous layer (consisting primarily of adipocytes) resisted consistent clearing (yellow tissue, Figure S2 A) due to incomplete micelle solvation of all the packed lipids in adipocytes.

Scanning Electron Microscopy Samples were imaged on an FEI Quanta 200F environmental scanning electron microscope (ESEM) in ESEM mode. Thin slices of PACT-processed brain tissues were placed on the sample holder in the chamber and imaged at a voltage of 5 kV and working distance between 7.7–8.3 mm with a spot size of 3 or 4 using the gaseous secondary electron detector (GSED). Please note that stretching during cutting/SEM process will make the pores of tissue-hydrogel hybrid larger. The actual effective pore size of the SEM images should be smaller than we present here.

Quantification Methods Mean nearest neighbor distance (NND): 1-mm-thick coronal slices were stained with DAPI and imaged w/ the 10× 0.45 N.A. Plan-Apochromat objective. Twenty-four 3-μm-thick images were taken from different regions of the cortex thalamus and striatum. A 30 pixel rolling ball radius subtraction filter was used to remove the background. Individual images are thresholded and converted into binaries. A binary watershed segmentation was applied to divide cells that are clustered together. The resulting images were quantified with the analyzing particles option on ImageJ. The centroid of each cell was identified in the measurement, and the NND's were calculated by applying the “nnd” plugin on imageJ. GFP size quantification: 1-mm-thick coronal slices of AAV9-eGFP IV injected mouse brain were imaged with the 5× 0.25N.A. Fluar objective. A maximum projection of the Z-stack was used for quantification. The area of each GFP positive neuron was isolated and quantified with the analyzing particles option on ImageJ. Both nearest neighbor distance measurements and GFP size calculations were performed for three brain regions: cortex, striatum, and thalamus; in uncleared, PARS cleared, and PARS cleared then postfixed mouse brain slices (see Figure S5 A for representative brain slices and Figure S5 B for data results). The mean cell size and mean NND for all counted cells were computed for each region, and data were analyzed for statistically significant differences in cell size or in NND between regions.