TITLE: Human Pancreas PACT Optical Clearing and High Resolution 3D Microscopy
AUTHORS: Elizabeth Butterworth, Wesley Dickerson, Vindhya Vijay, Kristina Weitzel, Julia Cooper, Eric W. Atkinson, Jason E. Coleman, Kevin Otto, Martha Campbell Thompson

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block">Using traditional histological methods, researchers are hampered in their ability to image whole tissues or organs in large-scale 3D. Histological sections are generally limited to <20 μm as formalin fixed paraffin section on glass slides or <500 μm for free-floating fixed sections. Therefore, extensive efforts are required for serial sectioning and large-scale image reconstruction methods to recreate 3D for samples >500 μm using traditional methods. In addition, light scatters from macromolecules within tissues, particularly lipids, prevents imaging to a depth >150 μm with most confocal microscopes. To reduce light scatter and to allow for deep tissue imaging using simple confocal microscopy, various optical clearing methods have been developed that are relevant for rodent and human tissue samples fixed by immersion. Several methods are related and use protein crosslinking with acrylamide and tissue clearing with sodium dodecyl sulfate (SDS). Other optical clearing techniques used various solvents though each modification had various advantages and disadvantages. Here, an optimized passive optical clearing method is described for studies of the human pancreas innervation and specifically for interrogation of the innervation of human islets.</div></div>

[STEPS]
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Use a new razor blade or scalpel to cut through the paraffin perpendicular to the surface of the tissue. Cut the paraffin at the edge of the tissue to finish loosening it. Use forceps or a spatula to gently loosen and remove the tissue to be optically cleared. Gently scrape excess paraffin from the tissue using a spatula (Figure 1A).
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Fill a glass container with xylene (about 30 mL for a 3 x 3 x 3 mm section of tissue) and incubate the tissue in this solution for 24 h at room temperature (RT).
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Fill another glass container with fresh xylene with the same volume as 1.2.1. Transfer and incubate the tissue in this solution for 24 h at RT.
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Place 100% ethanol into a conical tube with the same volume as 1.2.1. Transfer and incubate the tissue in this solution for 24 h at RT.
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Place 95% ethanol into a conical tube with the same volume as 1.2.1. Transfer and incubate the tissue in this solution for 24 h at RT.
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Place 70% ethanol into a conical tube with the same volume as 1.2.1. Transfer and incubate the tissue in this solution for 24 h at RT.
?. [Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)]
Rinse the tissue in 0.01 M phosphate buffered saline (PBS) and place in 0.01 M PBS in a conical tube to equilibrate for 24 h at RT. Ensure that the tissue is free of paraffin (Figure 1B, right panel).
?. [Prepare 4% Paraformaldehyde (PFA) Fixative]
Pipette 10 mL 16% PFA into a 50-mL conical tube, add 4 mL 0.1 M PBS, and add 26 mL distilled deionized water (ddH2O). Close the cap and mix briefly.
?. [Prepare 4% Paraformaldehyde (PFA) Fixative]
Larger volumes of 4% PFA can be made ahead and frozen in aliquots. Aliquots are good for 1 day at room temperature (RT), one week at 4 °C, and 1 month at -20 °C
?. [Pancreas Fixation]
Fix the pancreas sample (≤ 1 x 1 x 2 cm) in freshly prepared 4% PFA at 4 °C for 48 h. If the sample is larger than 1 x 1 x 2 cm, use a scalpel or razor blade to dissect into smaller pieces no more than 1 cm thick. Wash tissue sample in three changes of 0.01 M PBS for at least 15 min each wash and store in 15 mL centrifuge tube in 0.01 % PFA/0.01 M PBS or 0.5 % sodium azide/0.01 M PBS until use.
?. [Pancreas Fixation]
After fixation, section the tissue into 1 - 2 mm thick sections for further processing. Use a vibratome to assist in even sectioning. NOTE: The final number of sections will depend on the size of the starting sample.
?. [Embed Tissue in Hydrogel]
Prepare 200 mL of the 4 % acrylamide/1 % paraformaldehyde (A4P1) hydrogel monomer solution as follows:
?. Place a 15-mL conical tube in the ice next to the flask containing the hydrogel solution. Pipette 14 mL of monomer solution into the tube and add one piece of the 1 - 2 mm thick fixed tissue sample. Then cap the tube.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Remove oxygen from the hydrogel monomer solution using gaseous N28.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Once the oxygen is removed, quickly remove both needles and cover the cap with a paraffin film to prevent any further exchange of gasses between the tube and the environment. Place the degassed sample in an incubator at 37 °C for 3 h to polymerize the hydrogel.
?. [Tissue Clearing]
Prepare 500 mL clearing solution (4 % SDS at pH 8.5). To ~300 mL of ddH2O, add 50 mL 0.1 M PBS and 20 g SDS powder while stirring with a magnetic stir bar. Adjust the solution using sodium hydroxide and hydrochloric acid to pH 8.5. Add ddH2O until the final volume is 500 mL.
?. [Tissue Clearing]
After polymerization, pour away excess hydrogel and discard it into a chemical waste container. Use a paper towel to gently wipe away hydrogel from the sample and discard into a chemical waste container.
?. [Multiple Immunofluorescence]
Wash the samples on a shaker at 60 rpm at RT for one day with 40 mL 0.01 M PBS changing to fresh buffer often (4 - 5 buffer changes in total, 40 mL each wash, changing every h until the final wash). Let the final wash continue overnight at RT.
?. [Multiple Immunofluorescence]
Prepare PACT staining buffer. To 500 mL 0.01 M PBS, add 50 mg sodium azide and 0.5 mL TritonX-100. Mix well.
?. [Mounting Samples for Imaging]
Prepare the refractive index matched solution (RIMS) buffer.
?. [Mounting Samples for Imaging]
Transfer samples into RIMS. To do so, pipette 1 mL RIMs into a 2-mL flat-bottom tube. Use a spatula to remove the sample from wash buffer and dab the excess buffer off onto a paper towel, then place in the tube with RIMS solution.
?. [Confocal Imaging]
Select appropriate lasers for the excitation and emission spectra of the fluorophores used to stain the PACT samples. Adjust the settings of the acquisition software so that any overlap between channels is eliminated. Use separate tracks if necessary (when twofluorophores have similar excitation spectra).
?. [Embed Tissue in Hydrogel]
Place a flask on the ice in a bucket on the top of a magnetic stir plate. Make sure the flask is sitting flat and add a magnetic stir bar.
?. [Embed Tissue in Hydrogel]
Add the following in order: 147.8 mL cold (4 - 8 °C) ddH2O, 20 mL 0.1 M PBS, 20 mL cold (4 - 8 °C) 40 % acrylamide solution, 12.2 mL 16 % PFA solution, and 250 mg VA-044 initiator. Mix the entire hydrogel solution with a magnetic stir bar for at least 10 min and leave the solution on ice for the next step.
?. Incubate the sample in monomer solution for 3 days at 4 °C and protect from light. Aliquot any remaining monomer solution and store at -20°C for future use.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Prepare a bucket of ice and place the sample in the hydrogel monomer solution on ice.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Gather 2 - 3, 18-gauge hypodermic needles per sample, paraffin film, and a timer. Connect the tubing to the nitrogen tank so that the nitrogen can flow. While keeping the sample on ice, carefully pierce the cap of a conical tube containing the sample on one side and press one hypodermic needle through until it is under the surface of the liquid monomer solution.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Use another hypodermic needle to puncture the opposite side of the cap, but do not allow it to become submerged. NOTE: The second needle will vent the tube.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Connect the tubing from the nitrogen tank to the hypodermic needle submerged beneath the hydrogel and slowly turn on the nitrogen until it is bubbling steadily through the liquid.
?. [Degas the Monomer Solution and Polymerize the Hydrogel]
Allow the nitrogen to bubble through the liquid for 10 min.
?. [Tissue Clearing]
Wash the sample in 3 - 5 exchanges of 0.01 M PBS (discard wash fluid into the chemical waste) for 15 min each wash step. Transfer the sample into a 50-mL conical tube with 40 mL of clearing buffer.
?. [Tissue Clearing]
Incubate the sample in the clearing buffer at 37 °C and change sample to fresh clearing buffer every other day.
?. [Tissue Clearing]
Leave the sample in the clearing buffer for 2 - 8 weeks depending on the sample size (~8 weeks for a 3 mm x 3 mm x 3 mm sample) to ensure proper clearing.
?. [Tissue Clearing]
Monitor tissue clearing and stop when complete. Ensure that the sample is adequately transparent by holding it up to the light to check for proper clearing (usually some tan coloring will remain in the exocrine regions). NOTE: An over-cleared sample will appear frayed at the edges and the texture will be very soft when picked up with forceps. It is common for the sample to clear unevenly. Also, the tissue will not be fully transparent until placed in mounting media (Insert, Figure 1C).
?. [Multiple Immunofluorescence]
Incubate the sample with primary antibodies.
?. [Multiple Immunofluorescence]
Add 2% normal serum (same species as the secondary antibody) to the base PACT staining buffer in a 2-mL flat bottom tube (at least 1mL total volume is recommended per sample/tube).
?. [Multiple Immunofluorescence]
Add primary antibody to the 2 % serum/PACT staining buffer. Use approximately 5x the amount of primary antibody for PACT staining as would be used for standard immunohistochemistry (i.e. if an antibody is diluted 1:500 for standard immunohistochemistry, use 1:100 for PACT staining).
?. [Multiple Immunofluorescence]
Use a spatula to remove the sample from the wash buffer and dab the excess buffer off onto a paper towel, then place in the tube with a primary antibody solution.
?. [Multiple Immunofluorescence]
Incubate 2-4 days at RT on a shaker at 60 rpm. Wash samples thoroughly at RT on a shaker at 60 rpm in 0.01 M PBS changing to fresh buffer 4-5 times and leaving the final wash on overnight as in step 7.1.
?. [Multiple Immunofluorescence]
Incubate samples with secondary antibodies.
?. [Multiple Immunofluorescence]
Incubate at RT on a shaker at 60 rpm for 2 days and protect the sample from light.
?. [Multiple Immunofluorescence]
Wash the samples thoroughly, as in step 7.1, at RT on a shaker at 60 rpm in 0.01 M PBS changing to the fresh buffer 4 - 5 times and leaving the final wash on overnight, protect from light during washes.
?. [Multiple Immunofluorescence]
Add 2% normal serum (same species as the secondary antibody) to the base PACT staining buffer in a 2-mL flat bottom tube (1 mL total volume is recommended per sample/tube).
?. [Multiple Immunofluorescence]
Add secondary antibodies at a concentration of 1:200 (5 μl in 1 mL buffer).NOTE: Small format antibodies are preferred, as well as highly cross-adsorbed antibodies if using more than one primary antibody.
?. [Multiple Immunofluorescence]
Use a spatula to remove the sample from wash buffer and dab the excess buffer off onto a paper towel, then place in the tube with asecondary antibody solution.
?. [Mounting Samples for Imaging]
Weigh out 11 g of non-ionic density gradient medium (e.g., Histodenz) and carefully transfer to a 50-mL conical tube.
?. [Mounting Samples for Imaging]
Add ~5 mL 0.02 M phosphate buffer (PB)8 using a spatula to release air from the powder non-ionic density gradient medium.NOTE: The solution will be very viscous, mix well.
?. [Mounting Samples for Imaging]
Bring the volume to 10 mL using more PB, mix with a spatula and scrape the excess off the spatula into the tube.
?. [Mounting Samples for Imaging]
To image, place a small amount of RIMs into an 8-well coverslip bottom chamber slide. Only add just enough to coat the bottom, more will cause the sample to float making it more difficult to image on an inverted scope. Add the sample to the well and cap the slide for imaging.
?. [Mounting Samples for Imaging]
Gently tap the tube to submerge the sample in RIMS. Place samples in RIMS on the bench protected from light at RT for 2 - 4 days before imaging.
?. [Mounting Samples for Imaging]
Incubate RIMS at 37 °C until dissolved, invert and gently mix as needed
?. [Lightsheet Imaging]
Ensure that the samples have been equilibrated in RIMS for imaging for at least 24 h. Ideally, transfer the samples to fresh RIMS in the imaging chamber and allow to equilibrate in the chamber for at least a day.
?. [Lightsheet Imaging]
Mount the sample for imaging
?. [Lightsheet Imaging]
Select the smallest (black) capillary to mount the sample
?. [Lightsheet Imaging]
Use putty or a dish to hold the sample while applying super glue to the end of the capillary. Glue the tissue to the capillary touching as little a surface of the tissue as possible.
?. [Lightsheet Imaging]
Insert the sample for imaging.
?. [Lightsheet Imaging]
Image using 5X or 25X objectives suitable for optically cleared samples using the pivot scan option. If shadowing or blurring occurs, rotate the sample and try again, or let the sample continue to equilibrate in RIMS.NOTE: A 1 mm stack can generally be acquired in less than five minutes, depending on the settings.
?. [Lightsheet Imaging]
View and edit the image stacks using the image analysis software.
?. [Confocal Imaging]
Setup the z-stack
?. [Confocal Imaging]
Set up the image acquisition
?. [Confocal Imaging]
Select the optimal sectioning for the objective being used. If deconvolution is desired later, use a z-step smaller than optimal such as 1 μm.
?. [Confocal Imaging]
Use the z-stack correction.
?. [Confocal Imaging]
Begin nearest the surface of the tissue and increase the gain as the objective focuses through the z-plane and addcorrections. Do not change the laser settings for the correction!
?. [Confocal Imaging]
Acquire a test stack with one average, 512 x 512 resolution, and maximum acquisition speed. Check the image in 3D to make sure that there is equal brightness throughout the stack for each color before acquiring the final high-resolution zstack.
?. [Confocal Imaging]
Choose maximum acquisition speed, 16 bit, 4 or more averages, and 1024 x 1024 resolution or better.
?. [Confocal Imaging]
Zoom into the object to be imaged.NOTE: This will decrease acquisition time to reduce photo-bleaching and also decrease file size and downstream editing.