Aug 14, 2020
Human Pancreas PACT Optical Clearing and High Resolution 3D Microscopy
- Elizabeth Butterworth1,
- Wesley Dickerson1,
- Vindhya Vijay1,
- Kristina Weitzel1,
- Julia Cooper1,
- Eric W. Atkinson1,
- Jason E. Coleman1,
- Kevin Otto1,
- Martha Campbell-Thompson1
- 1University of Florida
- Human BioMolecular Atlas Program (HuBMAP) Method Development CommunityTech. support email: Jeff.spraggins@vanderbilt.edu
External link: http://doi:10.3791/56859
Protocol Citation: Elizabeth Butterworth, Wesley Dickerson, Vindhya Vijay, Kristina Weitzel, Julia Cooper, Eric W. Atkinson, Jason E. Coleman, Kevin Otto, Martha Campbell-Thompson 2020. Human Pancreas PACT Optical Clearing and High Resolution 3D Microscopy. protocols.io https://dx.doi.org/10.17504/protocols.io.9gbh3sn
Manuscript citation:
Butterworth, E., Dickerson, W., Vijay, V., Weitzel, K., Cooper, J., Atkinson, E.W., Coleman, J.E., Otto, K.J., Campbell-Thompson, M. High Resolution 3D Imaging of the Human Pancreas Neuro-insular Network. J. Vis. Exp. (131), e56859, doi:10.3791/56859 (2018).
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: November 18, 2019
Last Modified: August 14, 2020
Protocol Integer ID: 29923
Keywords: Bioengineering, Issue 131, Pancreas, islet, PACT, optical clearing, immunofluorescence, insulin, β-cells, glucagon, α-cells, nerves, Schwann cells, GFAP, confocal microscopy, Lightsheet microscopy
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Abstract
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.
Attachments
Materials
| Name of Material/Equipment | Company | Catalog Number | Comments | |||
| 10x phosphate buffered saline (PBS) | Fisher | BP399-1 | Buffers | |||
| Sodium phosphate dibasic anhydrous | Fisher | S375-500 | PB buffer (RIMS) | |||
| Sodium phosphate monobasic monohydrate | Sigma | 71507-250 | PB buffer (RIMS) | |||
| 16% paraformaldehyde (PFA) | Electron Microscopy Sciences | 15714-5 | Immersion fixation, hydrogel, storage solution | |||
| 40% acrylamide | Bio-Rad | 161-0140 | Hydrogel | |||
| 2% bis-acrylamide | Bio-Rad | 161-0142 | Hydrogel | |||
| VA-044 initiator | Wako Pure Chemical Industries, Ltd. | VA044 | Hydrogel | |||
| Sodium dodecyl sulfate (SDS) | Fisher | BP166-5 | Clearing buffer | |||
| Sodium azide | Sigma | S8032 | Sample storage buffer | |||
| 18 gauge needles | Fisher | 14-840-91 | Degassing hydrogel solution | |||
| N2 tank | AirGas | various | Degassing hydrogel solution | |||
| Triton X-100 | Sigma-Aldrich | 100 ml | Buffers | |||
| Goat, normal serum | Vector | S-1000 | Use as 2% in blocking buffer | |||
| Histodenz | Sigma | D2158-100G | RIMS | |||
| 8-well chamber slides | Ibidi | 80827 | Imaging | |||
| Laser scanning confocal microscope | Zeiss | 710 | Imaging | |||
| LightSheet microscope | Zeiss | Z1 | Imaging | |||
| Primary Antibody | Host | Vendor | Cat. # | Dilution | Comments | |
| CD45 | Rabbit | Bioss | bs-4820R-A488 | 1:100 | Did not work | |
| CD45 | Mouse | DAKO | M0754 | 1:200 | Did not work | |
| GFAP | Rabbit | DAKO | Z0334 | 1:50 | Worked | |
| Glucagon | Mouse | BD Biosciences | 565891 | 1:50 | Worked | |
| Glucagon | Rabbit | Cell Signaling | 2760S | 1:200 | Did not work | |
| Glucagon | Mouse | Abcam | ab10988 | 1:200 | Worked | |
| Insulin | Guinea Pig | DAKO | A0564 | 1:200 | Worked | |
| NCAM (CD56) | Mouse | DAKO | M730429-2 | 1:50 | Did not work | |
| NCAM (CD56)-FITC conjugate | Mouse | DAKO | M730429-2 | 1:50 | Did not work | |
| Peripherin | Rabbit | EnCor | RPCA-Peri | 1:100 | Worked | |
| PGP9.5 | Rabbit | DAKO | Z5116 | 1:50 | Did not work | |
| Secretogranin 3 | Rabbit | Sigma | HPA006880 | 1:200 | Worked | |
| Smooth muscle actin | Mouse | Sigma | A5228; C6198 (Cy5) | 1:200; 1:200 | Worked; Conjugated worked better than unconjugated | |
| Substance P | Rat | BioRad | 8450-0505 | 1:200 | Worked | |
| Tyrosine Hydroxylase | Rabbit | Millipore | AB152 | 1:200 | Worked | |
| Tyrosine Hydroxylase | Chicken | Abcam | Ab76442 | 1:100 | Worked, but weak staining | |
| Vasoactive Intestinal Peptide (VIP) | Rabbit | Immunostar | 20077 | 1:100 | Worked | |
| Vesicular Acetylcholine Transporter (VAChT) | Rabbit | Synaptic Systems | 139103 | 1:50 | Worked | |
| Secondary Antibody | ||||||
| Guinea pig IgG | Goat | ThermoFisher Scientific | Various | 1:200 | AlexaFluor conjugates | |
| Mouse IgG | Goat | ThermoFisher Scientific | Various | 1:200 | AlexaFluor conjugates | |
| Rabbit IgG | Goat | ThermoFisher Scientific | Various | 1:200 | AlexaFluor conjugates | |
| Rat IgG | Goat, Donkey | ThermoFisher Scientific | Various | 1:200 | AlexaFluor conjugates | |
| Chicken IgG | Goat | ThermoFisher Scientific | Various | 1:200 | AlexaFluor conjugates |
Safety warnings
Paraformaldehyde, xylene, and acrylamide are toxic irritants. Handle reagents in a fume hood with appropriate personal protective equipment (lab coat, gloves, eye protection). Follow EHS guidelines for all chemical handling and disposal requirements.
Deparaffinization of Formaldehyde-fixed Paraffin Embedded Tissues (If Working with Fresh Tissues, Skip to Step 10)
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).
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).
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.
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.
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.
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.
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
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.
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
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.
2d
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
Embed Tissue in Hydrogel
Prepare 200 mL of the 4 % acrylamide/1 % paraformaldehyde (A4P1) hydrogel monomer solution as follows:
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.
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.
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.
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
Degas the Monomer Solution and Polymerize the Hydrogel
Remove oxygen from the hydrogel monomer solution using gaseous N28.
Prepare a bucket of ice and place the sample in the hydrogel monomer solution on ice.
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.
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.
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.
Allow the nitrogen to bubble through the liquid for 10 min.
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
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.
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.
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.
Incubate the sample in the clearing buffer at 37 °C and change sample to fresh clearing buffer every other day.
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.
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
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.
Prepare PACT staining buffer. To 500 mL 0.01 M PBS, add 50 mg sodium azide and 0.5 mL TritonX-100. Mix well.
Incubate the sample with primary antibodies.
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).
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).
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.
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.
Incubate samples with secondary antibodies.
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).
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.
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 a
secondary antibody solution.
Incubate at RT on a shaker at 60 rpm for 2 days and protect the sample from light.
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.
Mounting Samples for Imaging
Mounting Samples for Imaging
Prepare the refractive index matched solution (RIMS) buffer.
Weigh out 11 g of non-ionic density gradient medium (e.g., Histodenz) and carefully transfer to a 50-mL conical tube.
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.
Bring the volume to 10 mL using more PB, mix with a spatula and scrape the excess off the spatula into the tube.
Incubate RIMS at 37 °C until dissolved, invert and gently mix as needed
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.
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.
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.
Confocal Imaging
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).
Set up the image acquisition
Choose maximum acquisition speed, 16 bit, 4 or more averages, and 1024 x 1024 resolution or better.
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.
Setup the z-stack
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.
Use the z-stack correction.
Begin nearest the surface of the tissue and increase the gain as the objective focuses through the z-plane and add
corrections. Do not change the laser settings for the correction!
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.
Lightsheet Imaging
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.
Equipment
Lightsheet
NAME
Light Sheet Microscopy
TYPE
Zeiss
BRAND
2583000198
SKU
LINK
Mount the sample for imaging
Select the smallest (black) capillary to mount the sample
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.
Insert the sample for 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.
View and edit the image stacks using the image analysis software.