The authors thank Dr. Ann Fu and Joseph Canzano for technical assistance, Dr. Jennifer Treweek for invaluable advice, and Dr. Kristin Overton and Dr. Karl Deisseroth for training to MCT through the Stanford University CLARITY workshop. JDRF nPOD and the Organ Procurement Organizations provided tissue samples. This work was funded by JDRF (47-2014-1), Helmsley Charitable Trust (HCT 2015-PG-T1D052) and NIDDK 1OT2 TR001773 to MCT.

All experiments were conducted in accordance with University of Florida Institutional Review Board and Federal guidelines.
Caution: Paraformaldehyde and acrylamide are toxic irritants. Handle reagents in a fume hood with appropriate personal protective equipment (lab coat, gloves, eye protection).
1. Deparaffinization of Formaldehyde-fixed Tissues (If Working with Fresh Tissues, Skip to Step 2)
<ol>
	<li>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).</li>
	<li>Fill a glass container with xylene (about 30 mL for a 3 x 3 x&#160;3 mm section of tissue) and incubate the tissue in this solution for 24 h at room temperature (RT).</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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&#160;(Figure 1B, right panel).</li>
</ol>
2. Prepare 4% Paraformaldehyde (PFA) Fixative
<ol>
	<li>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.</li>
	<li>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 &#176;C, and 1 month at -20 &#176;C.</li>
</ol>
3. Pancreas Fixation
<ol>
	<li>Fix the pancreas sample (&#8804; 1 x 1 x 2 cm) in freshly prepared 4% PFA at 4 &#176;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.</li>
	<li>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.</li>
</ol>
4. Embed Tissue in Hydrogel
<ol>
	<li>Prepare 200 mL of the 4 % acrylamide/1 % paraformaldehyde (A4P1) hydrogel monomer solution as follows:
	<ol>
		<li>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.</li>
		<li>Add the following in order: 147.8 mL cold (4 - 8 &#176;C) ddH2O, 20 mL 0.1 M PBS, 20 mL cold (4 - 8 &#176;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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Incubate the sample in monomer solution for 3 days at 4 &#176;C and protect from light. Aliquot any remaining monomer solution and store at -20 &#176;C for future use.</li>
</ol>
5. Degas the Monomer Solution and Polymerize the Hydrogel
<ol>
	<li>Remove oxygen from the hydrogel monomer solution using gaseous N28.
	<ol>
		<li>Prepare a bucket of ice and place the sample in the hydrogel monomer solution on ice.</li>
		<li>Gather 2 - 3, 18-gauge hypodermic needles per sample, paraffin film, and a timer. Connect the tubing to the nitrogen tank&#160;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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>Allow the nitrogen to bubble through the liquid for 10 min.</li>
	</ol>
	</li>
	<li>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 &#176;C for 3 h to polymerize the hydrogel.</li>
</ol>
6. Tissue Clearing
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Incubate the sample in the clearing buffer at 37 &#176;C and change sample to fresh clearing buffer every other day.
	<ol>
		<li>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.</li>
		<li>Monitor tissue clearing and stop when complete.&#160;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).</li>
	</ol>
	</li>
</ol>
7. Multiple Immunofluorescence
<ol>
	<li>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.</li>
	<li>Prepare PACT staining buffer. To 500 mL 0.01 M PBS, add 50 mg sodium azide and 0.5 mL TritonX-100. Mix well.</li>
	<li>Incubate the sample with primary antibodies.
	<ol>
		<li>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 1 mL total volume is recommended per sample/tube).</li>
		<li>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).</li>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
	<li>Incubate samples with secondary antibodies.
	<ol>
		<li>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).</li>
		<li>Add secondary antibodies at a concentration of 1:200 (5 &#181;l in 1 mL buffer). 
		NOTE: Small format antibodies are preferred, as well as highly cross-adsorbed antibodies if using more than one primary antibody.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Incubate at RT on a shaker at 60 rpm for 2 days and protect the sample from light.</li>
	<li>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.</li>
</ol>
8. Mounting Samples for Imaging
<ol>
	<li>Prepare the refractive index matched solution (RIMS) buffer.
	<ol>
		<li>Weigh out 11 g of non-ionic density gradient medium (e.g., Histodenz) and carefully transfer to a 50-mL conical tube.</li>
		<li>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.</li>
		<li>Bring the volume to 10 mL using more PB, mix with a spatula and scrape the excess off the spatula into the tube.</li>
		<li>Incubate RIMS at 37 &#176;C until dissolved, invert and gently mix as needed.</li>
	</ol>
	</li>
	<li>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.
	<ol>
		<li>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.</li>
	</ol>
	</li>
	<li>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.</li>
</ol>
9. Imaging
<ol>
	<li>Confocal microscopy and imaging software
	<ol>
		<li>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 two fluorophores have similar excitation spectra).</li>
		<li>Set up the image acquisition
		<ol>
			<li>Choose maximum acquisition speed, 16 bit, 4 or more averages, and 1024 x 1024 resolution or better.</li>
			<li>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.</li>
		</ol>
		</li>
		<li>Setup the z-stack.
		<ol>
			<li>Select the optimal sectioning for the objective being used. If deconvolution is desired later, use a z-step smaller than optimal such as 1 &#181;m.</li>
			<li>Use the z-stack correction.
			<ol>
				<li>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!</li>
				<li>Acquire a test stack with one average,&#160;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 z-stack.</li>
			</ol>
			</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Lightsheet Imaging
	<ol>
		<li>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.</li>
		<li>Mount the sample for imaging
		<ol>
			<li>Select the smallest (black) capillary to mount the sample</li>
			<li>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.</li>
			<li>Insert the sample for imaging.</li>
		</ol>
		</li>
		<li>Image using 5X&#160;or 25X&#160;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.</li>
		<li>View and edit the image stacks using the image analysis software.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Availability of new optical clearing methods has permitted unprecedented large-scale examinations of the central and peripheral nervous systems in animal models. The overall innervation patterns of the human pancreas are largely unexplored due to tissue density and difficulties in acquiring high quality biospecimens. This protocol offers an optimized tissue clearing protocol for human pancreas tissue from either fixed or paraffin-embedded archival samples.
Clearing time depends on the sample size, fixative type, duration, storage duration and may require 1 - 8 weeks, so the following considerations are critical. It is best to clear tissues soon after the 4% PFA fixation if possible because long-term storage increases clearing time. To minimize clearing time, the amount of PFA in the hydrogel monomer was decreased from 4% as used in the fixation step to 1% for the human pancreas. For other tissues, especially mouse tissues, the amount of PFA necessary to provide sufficient hydrogel rigidity to preserve antigens must be determined empirically. Adequate clearing is also essential since antibody penetration is reduced in poorly cleared tissues and surface staining by secondary antibodies is increased. Changing the clearing solution every other day (or, even daily) is best to keep the pH constant and detergent fresh. Conversely, over-clearing negatively impacts cellular morphology and increases tissue friability. Since some pancreas samples clear unevenly due to inherent pathologies such as regions of fibrosis from chronic pancreatitis or other pathologies, it is important to frequently monitor this process. Use of vibratome thick sections can also be used to make uniform thicknesses yet are not required. Monitoring the clearing process, so samples are removed from detergent as soon as light passes easily through the sample, ensures that the sample is sufficiently cleared.
Antibody penetration and surface staining are important issues to consider with PACT samples. To ensure good antibody penetration, it is critical to incubate samples with the primary antibodies for at least 2 days. Various antibodies have different diffusion rates and should be optimized individually, but for most antibodies, 4 days was sufficient duration for full penetration in a 1 mm3 sample. If surface staining is a problem, especially for Lightsheet imaging, the sample may be cut in half, or the surface dissected away after staining. Small format antibodies when available would be expected to assist with tissue penetration8. Preconjugated antibodies do not appear to work as well as the same unconjugated clone and higher background from nonspecific binding and poorer tissue penetration can be observed if they work at all. For improved success with preconjugated antibodies, increase serum and TritonX-100 concentrations in the staining buffer and use longer incubations (&#62;4 days). After staining, incubation of the sample in RIMS for several days is critical. Cleared tissues swell in PBS and RIMS incubation causes them to shrink back to original size. Especially with lightsheet imaging, the sample should be fully equilibrated before imaging.
When imaging samples on the confocal microscope, the correction must be set each time a new position is chosen. Various acquisition depths and variation in staining intensity throughout the tissue necessitates adjustments for each area of the tissue to be imaged optimally. Likewise, the secondary antibodies used must be carefully considered. Spectral unmixing is challenging in PACT samples, so fluorophores must be appropriately chosen for a low excitation or emission overlap to ensure a bright signal when filtering emissions on the confocal microscope. For each application, a balance must be struck between resolution, imaging speed, and noise reduction so that the best quality image can be acquired without photo-bleaching. Resolution greater than 1024 x 1024 is not detectable by the human eye but may be necessary for certain applications if quantification of a stain is needed.
There are many things to consider when choosing an optical clearing technique and when choosing optical clearing over other more traditional methods. Low abundance antigens may not be detectable using the PACT method thus there are limitations on antigen detection. Certain antigens such as immunological antigens (CD3, CD4, CD8, etc.) are destroyed by the PACT method and are undetectable despite high quality antibodies available to detect them. Another limitation of this method is the inherent variability between donor pancreases which are difficult to discern until after clearing staining. Each donor tissue tends to clear and stain similarly between procedures, but different donor tissues have widely varying clearing times and tissue morphology quality after clearing. The ability to use archival tissue may mitigate this limitation if one can have adequate access to a large number of patient pancreas samples though this has not been thoroughly investigated. The PACT protocol reported herein was found to be inexpensive and readily implemented with standard laboratory equipment. Cleared samples were suitable for imaging via traditional confocal microscopy, as well as 2-photon and Lightsheet technologies and provided high resolution 3D images of nerves and islets in large sections of the human pancreas. Future applications of this procedure include studies on the development of the fetal pancreas for both exocrine and endocrine compartments and islet studies in type 1 and type 2 diabetes.

Until recently, 3-dimensional (3D) reconstruction of large tissues or whole organs was performed through laborious serial sectioning, staining, and image reconstruction of multiple sections. These methods had several disadvantages including reliance on a large number of serial sections, poor tissue penetration with antibodies, and light scattering preventing deep imaging within tissues. To allow for greater light and antibody penetration, researchers developed chemical methods to preserve protein antigens while removing the majority of light-scattering lipids. Several related methods are Clear Lipid-Exchanged Acrylamide-hybridized Rigid Imaging Tissue hYdrogel (CLARITY), Passive Clarity Technique (PACT), and Perfusion-assisted Agent Release in situ (PARS) (reviewed in 1,2,3,4,5,6). The CLARITY method is based on stabilization of proteins using a formaldehyde, acrylamide, and bis hydrogel followed by lipid removal using electrophoresis in a SDS solution2. This approach was later modified for passive clearing and advanced imaging7. Further optimization led to the elimination of bis and varying percentages of paraformaldehyde in the hydrogel solution (1 - 4%) to obtain optimal antigen stabilization with the shortest clearing time for a technique called PACT 8. Early attempts to develop these optical clearing techniques involved organic or other compounds that were difficult to work with and quenched endogenous fluorescent proteins in transgenic mouse models. These additional methods included ScaleS, Unobstructed Brain/Body Cocktails and Computational analysis (CUBIC), SWITCH and Dimensional Imaging of Solvent-Cleared Organs (DISCO) methods, all with various advantages and disadvantages9,10,11,12. The original CLARITY method was developed in lipid-rich mouse brain tissue and optical clearing methods had limited testing in human tissues7,8,11,13,14.
Optical clearing methods are ideal for tracing nerves over long distances as in the intact mouse central nervous system. The pancreas is well innervated by both autonomic and sensory nervous systems. The pancreatic endocrine compartment, islets of Langerhans, comprise a very small portion of the entire organ (1 - 2%) and islets have known heterogeneity in sizes (50 - 250 &#181;m), endocrine cell proportions, and density particularly in diabetes (reviewed in 15,16). In developing a protocol, several optical clearing methods were tested for human pancreas and a PACT procedure was found to provide the best balance of time to clear (~2 weeks) with excellent morphological preservation of nerves and islets. The final optimized procedure is described here in the delineation of the neuro-insular network for large-scale (millimeter distances) and high-resolution 3D imaging of multiple intact pancreatic islets. The technique is suitable for human pancreas immediately following the fixation or after storage as well as samples fixed in neutral buffered formalin and embedded in paraffin wax. The samples are suitable for imaging by confocal or lightsheet microscopy.

<table>
	<tbody>
		<tr>
			<td>10x phosphate buffered saline (PBS)</td>
			<td>Fisher</td>
			<td>BP399-1</td>
			<td>Buffers</td>
		</tr>
		<tr>
			<td>Sodium phosphate dibasic anhydrous</td>
			<td>Fisher</td>
			<td>S375-500</td>
			<td>PB buffer (RIMS)</td>
		</tr>
		<tr>
			<td>Sodium phosphate monobasic monohydrate</td>
			<td>Sigma</td>
			<td>71507-250</td>
			<td>PB buffer (RIMS)</td>
		</tr>
		<tr>
			<td>16% paraformaldehyde (PFA)</td>
			<td>Electron Microscopy Sciences</td>
			<td>15714-5</td>
			<td>Immersion fixation, hydrogel, storage solution</td>
		</tr>
		<tr>
			<td>40% acrylamide</td>
			<td>Bio-Rad</td>
			<td>161-0140</td>
			<td>Hydrogel</td>
		</tr>
		<tr>
			<td>2% bis-acrylamide</td>
			<td>Bio-Rad</td>
			<td>161-0142</td>
			<td>Hydrogel</td>
		</tr>
		<tr>
			<td>VA-044 initiator</td>
			<td>Wako Pure Chemical Industries, Ltd.</td>
			<td>VA044</td>
			<td>Hydrogel</td>
		</tr>
		<tr>
			<td>Sodium dodecyl sulfate (SDS)</td>
			<td>Fisher</td>
			<td>BP166-5</td>
			<td>Clearing buffer</td>
		</tr>
		<tr>
			<td>Sodium azide</td>
			<td>Sigma</td>
			<td>S8032</td>
			<td>Sample storage buffer</td>
		</tr>
		<tr>
			<td>18 gauge needles</td>
			<td>Fisher</td>
			<td>14-840-91</td>
			<td>Degassing hydrogel solution</td>
		</tr>
		<tr>
			<td>N2 tank</td>
			<td>AirGas</td>
			<td>various</td>
			<td>Degassing hydrogel solution</td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma-Aldrich</td>
			<td>100 ml</td>
			<td>Buffers</td>
		</tr>
		<tr>
			<td>Goat, normal serum</td>
			<td>Vector</td>
			<td>S-1000</td>
			<td>Use as 2% in blocking buffer</td>
		</tr>
		<tr>
			<td>Histodenz</td>
			<td>Sigma</td>
			<td>D2158-100G</td>
			<td>RIMS</td>
		</tr>
		<tr>
			<td>8-well chamber slides</td>
			<td>Ibidi</td>
			<td>80827</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>Laser scanning confocal microscope</td>
			<td>Zeiss</td>
			<td>710</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>LightSheet microscope</td>
			<td>Zeiss</td>
			<td>Z1</td>
			<td>Imaging</td>
		</tr>
		<tr>
			<td>Name</td>
			<td>Company</td>
			<td>Catalog Number</td>
			<td>Comments</td>
		</tr>
		<tr>
			<td>Primary Antibody</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CD45</td>
			<td>Bioss</td>
			<td>bs-4820R-A488</td>
			<td>Host: Rabbit 
			Dilution: 1:100 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>CD45</td>
			<td>DAKO</td>
			<td>M0754</td>
			<td>Host: Mouse 
			Dilution: 1:200 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>GFAP</td>
			<td>DAKO</td>
			<td>Z0334</td>
			<td>Host: Rabbit 
			Dilution: 1:50 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Glucagon</td>
			<td>BD Biosciences</td>
			<td>565891</td>
			<td>Host: Mouse 
			Dilution: 1:50 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Glucagon</td>
			<td>Cell Signaling</td>
			<td>2760S</td>
			<td>Host: Rabbit 
			Dilution: 1:200 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>Glucagon</td>
			<td>Abcam</td>
			<td>ab10988</td>
			<td>Host: Mouse 
			Dilution: 1:200 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Insulin</td>
			<td>DAKO</td>
			<td>A0564</td>
			<td>Host: Guinea Pig 
			Dilution: 1:200 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>NCAM (CD56)</td>
			<td>DAKO</td>
			<td>M730429-2</td>
			<td>Host: Mouse 
			Dilution: 1:50 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>NCAM (CD56)-FITC conjugate</td>
			<td>DAKO</td>
			<td>M730429-2</td>
			<td>Host: Mouse 
			Dilution: 1:50 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>Peripherin</td>
			<td>EnCor</td>
			<td>RPCA-Peri</td>
			<td>Host: Rabbit 
			Dilution: 1:100 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>PGP9.5</td>
			<td>DAKO</td>
			<td>Z5116</td>
			<td>Host: Rabbit 
			Dilution: 1:50 
			Comments: Did not work</td>
		</tr>
		<tr>
			<td>Secretogranin 3</td>
			<td>Sigma</td>
			<td>HPA006880</td>
			<td>Host: Rabbit 
			Dilution: 1:200 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Smooth muscle actin</td>
			<td>Sigma</td>
			<td>A5228; C6198 (Cy5)</td>
			<td>Host: Mouse 
			Dilution: 1:200; 1:200 
			Comments: Worked; Conjugated worked better than unconjugated</td>
		</tr>
		<tr>
			<td>Substance P</td>
			<td>BioRad</td>
			<td>8450-0505</td>
			<td>Host: Rat 
			Dilution: 1:200 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Tyrosine Hydroxylase</td>
			<td>Millipore</td>
			<td>AB152</td>
			<td>Host: Rabbit 
			Dilution: 1:200 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Tyrosine Hydroxylase</td>
			<td>Abcam</td>
			<td>Ab76442</td>
			<td>Host: Chicken 
			Dilution: 1:100 
			Comments: Worked, but weak staining</td>
		</tr>
		<tr>
			<td>Vasoactive Intestinal Peptide (VIP)</td>
			<td>Immunostar</td>
			<td>20077</td>
			<td>Host: Rabbit 
			Dilution: 1:100 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Vesicular Acetylcholine Transporter (VAChT)</td>
			<td>Synaptic Systems</td>
			<td>139103</td>
			<td>Host: Rabbit 
			Dilution: 1:50 
			Comments: Worked</td>
		</tr>
		<tr>
			<td>Secondary Antibody</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Guinea pig IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>Various</td>
			<td>Host: Goat 
			Dilution: 1:200 
			Comments: AlexaFluor conjugates</td>
		</tr>
		<tr>
			<td>Mouse IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>Various</td>
			<td>Host: Goat 
			Dilution: 1:200 
			Comments: AlexaFluor conjugates</td>
		</tr>
		<tr>
			<td>Rabbit IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>Various</td>
			<td>Host: Goat 
			Dilution: 1:200 
			Comments: AlexaFluor conjugates</td>
		</tr>
		<tr>
			<td>Rat IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>Various</td>
			<td>Host: Goat, Donkey 
			Dilution: 1:200 
			Comments: AlexaFluor conjugates</td>
		</tr>
		<tr>
			<td>Chicken IgG</td>
			<td>ThermoFisher Scientific</td>
			<td>Various</td>
			<td>Host: Goat 
			Dilution: 1:200 
			Comments: AlexaFluor conjugates</td>
		</tr>
	</tbody>
</table>

high resolution 3d imaging of the human pancreas neuro-insular network

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 &#60;20 &#181;m as formalin fixed paraffin section on glass slides or &#60;500 &#181;m for free-floating fixed sections. Therefore, extensive efforts are required for serial sectioning and large-scale image reconstruction methods to recreate 3D for samples &#62;500 &#181;m using traditional methods. In addition, light scatters from macromolecules within tissues, particularly lipids, prevents imaging to a depth &#62;150 &#181;m 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.

These staining procedures were developed to provide a large-scale examination of the human pancreas to examine islets and associated autonomic and sensory networks. Several procedures were tested including CLARITY7, iDISCO17, and PACT8 with the PACT method found best suited for human pancreas and confocal imaging.
Primary antibodies were initially tested on fixed human pancreas samples with suitable positive and negative control samples (mouse brain, other). The primary antibodies and suggested dilutions are listed in Table of Materials, though each laboratory can expect to optimize antibody dilutions depending on lot number and tissue source. Primary antibody lot-to-lot variation can impact the staining intensity and the specificity, and require validation to achieve the same degree of stain intensity as with former reagents.
In the human pancreas, islets were delineated by insulin, glucagon, and secretogranin 3 (Figure 2). Schwann cells were delineated using glial fibrillary acid protein (GFAP, Figure 2 and Figure 3A). Nerves stained with antibodies against the parasympathetic marker vasoactive intestinal peptide (VIP) were imaged on the Lightsheet (Figure 3B). Sympathetic nerves can be seen with staining for tyrosine hydroxylase (not shown).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56859/56859fig1.jpg" /> 
Figure 1. Human pancreas optical clearing. (A) A razor blade is used to cut and loosen a section of paraffin-embedded tissue (top panels) before removing the tissue section and scraping away excess paraffin from the tissue (bottom panels). Representative tissue samples are shown before (B left, C top) and after (B right, C middle) optical clearing of paraffin-embedded tissue, as described in the protocol section. The cleared, fixed tissue is not completely transparent after clearing (B right, C top) and often tissue swelling can be appreciated (C, middle panel). The cleared tissue becomes fully transparent after equilibration in RIMS (C, lower panel). Scale bars: 500 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56859/56859fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56859/56859fig2.jpg" /> 
Figure 2. Human neuro-insular network. Human pancreas samples were cleared as described from nPOD organ donors or from archival paraffin-embedded tissues. (A-D)&#160;Schwann cells (GFAP+, white) and endocrine cells are shown stained by insulin (red) and glucagon (green). (A)&#160;Confocal microscopy and maximum intensity projection (MIP) shows tracing of Schwann cells (GFAP+) on nerves coursing next to blood vessels at the islet periphery and extending into the islets. The inset demonstrates high resolution obtained and shows contact between GFAP+ Schwann cells and endocrine cells. (B)&#160;A 3D image of panel (scale: x = 50 &#181;m, y = 20 &#181;m, z = 25 &#181;m) A. A paraffin-PACT sample is shown imaged via confocal microscopy and presented as a max intensity projection (C) and in 3D (D), scale: x, y = 50 &#181;m, z = 25 &#181;m). Scale bars A, C: 50 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56859/56859fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56859/56859fig3.jpg" /> 
Figure 3. 3D visualization and stitching. Three stitched image stacks were acquired of Schwann cells stained using GFAP and confocal microscopy and traced using a neurite tracer plugin in the image analysis software. (A) The 3D fill of the trace is shown (Scale bars: x = 150 &#181;m, y = 200 &#181;m (0.2 mm), z = 120 &#181;m) (B)&#160;A PACT sample was stained with VIP antibody and imaged using a lightsheet microscope. The stack is &#62;1 mm in depth and nerve fibers are clearly seen in high resolution. Fibers wrapping a duct (D, foreground) and a ganglion (G, background) can be seen (Large grid: x, y, z = 200 &#181;m; tick marks: x, y, z = 40 &#181;m). <a href="https://cloudflare.jove.com/files/ftp_upload/56859/56859fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High Resolution 3D Imaging of the Human Pancreas Neuro-insular Network at JoVE.com

High Resolution 3D Imaging of the Human Pancreas Neuro-insular Network

Here, we present a protocol to image human pancreas sections in three dimensions (3D) using optimized passive clearing methods. This manuscript demonstrates these procedures for passive optical clearing followed by multiple immunofluorescence staining to identify key elements of the autonomic and sensory neural networks innervating human islets.

Using traditional histological methods, researchers are hampered in their ability to image whole tissues or organs in large-scale 3D. Histological ...

high-resolution-3d-imaging-of-the-human-pancreas-neuro-insular-network

Research

JoVE Journal

Bioengineering

10.1K Views.  University of Florida. Here, we present a protocol to image human pancreas sections in three dimensions (3D) using optimized passive clearing methods. This manuscript demonstrates these procedures for passive optical clearing followed by multiple immunofluorescence staining to identify key elements of the autonomic and sensory neural networks innervating human islets. 

Video: High Resolution 3D Imaging of the Human Pancreas Neuro-insular Network

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution. Traditionally, multiple experiments are run in parallel, with individual samples removed from the study at sequential time points for evaluation by light microscopy. Several intravital techniques have been developed, with confocal, multiphoton, and second harmonic microscopy all demonstrating their ability to be used for imaging in situ 1. With these systems, however, the required infrastructure is complex and expensive, involving scanning laser systems and complex light sources. Here we present a protocol for the design and assembly of a high-resolution microendoscope which can be built in a day using off-the-shelf components for under US$5,000. The platform offers flexibility in terms of image resolution, field-of-view, and operating wavelength, and we describe how these parameters can be easily modified to meet the specific needs of the end user.
We and others have explored the use of the high-resolution microendoscope (HRME) in in vitro cell culture 2-5, in excised 6 and living animal tissues 2,5, and in human tissues in vivo 2,7. Users have reported the use of several different fluorescent contrast agents, including proflavine 2-4, benzoporphyrin-derivative monoacid ring A (BPD-MA) 5, and fluoroscein 6,7, all of which have received full, or investigational approval from the FDA for use in human subjects. High-resolution microendoscopy, in the form described here, may appeal to a wide range of researchers working in the basic and clinical sciences. The technique offers an effective and economical approach which complements traditional benchtop microscopy, by enabling the user to perform high-resolution, longitudinal imaging in situ.

High-resolution Fiber-optic Microendoscopy for in situ Cellular Imaging

In many biological and clinical situations it is advantageous to study cellular processes as they evolve in their native microenvironment. Here we describe the assembly and use of a low-cost fiber-optic microscope which can provide real time imaging in cell culture, animal studies, and clinical patient studies.

Many biological and clinical studies require the longitudinal study and analysis of morphology and function with cellular level resolution.  ...

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography (&mu;CT).
High resolution &mu;CT is a very versatile yet accurate (1-2 microns of resolution) technique for 3D examination of ex-vivo biological samples1, 2. As opposed to electron tomography, the &mu;CT allows the examination of up to 4 cm thick samples. This technique requires only few hours of measurement as compared to weeks in histology. In addition, &mu;CT does not rely on 2D stereologic models, thus it may complement and in some cases can even replace histological methods3, 4, which are both time consuming and destructive. Sample conditioning and positioning in &mu;CT is straightforward and does not require high vacuum or low temperatures, which may adversely affect the structure. The sample is positioned and rotated 180&deg; or 360&deg;between a microfocused x-ray source and a detector, which includes a scintillator and an accurate CCD camera, For each angle a 2D image is taken, and then the entire volume is reconstructed using one of the different available algorithms5-7. The 3D resolution increases with the decrease of the rotation step. The present video protocol shows the main steps in preparation, immobilization and positioning of the sample followed by imaging at high resolution.

High Resolution 3D Imaging of Ex-Vivo Biological Samples by Micro CT

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized tomography ( CT).

Non-destructive volume visualization can be achieved only by tomographic techniques, of which the most efficient is the x-ray micro computerized ...

Three-dimensional Optical-resolution Photoacoustic Microscopy

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As the mainstays of in vivo three-dimensional (3-D) optical microscopy, single-/multi-photon fluorescence microscopy and optical coherence tomography (OCT) have demonstrated their extraordinary sensitivities to fluorescence and optical scattering contrasts, respectively. However, the optical absorption contrast of biological tissues, which encodes essential physiological/pathological information, has not yet been assessable. 
The emergence of biomedical photoacoustics has led to a new branch of optical microscopy optical-resolution photoacoustic microscopy (OR-PAM)1, where the optical irradiation is focused to the diffraction limit to achieve cellular1 or even subcellular2 level lateral resolution. As a valuable complement to existing optical microscopy technologies, OR-PAM brings in at least two novelties. First and most importantly, OR-PAM detects optical absorption contrasts with extraordinary sensitivity (i.e., 100%). Combining OR-PAM with fluorescence microscopy3 or with optical-scattering-based OCT4 (or with both) provides comprehensive optical properties of biological tissues. Second, OR-PAM encodes optical absorption into acoustic waves, in contrast to the pure optical processes in fluorescence microscopy and OCT, and provides background-free detection. The acoustic detection in OR-PAM mitigates the impacts of optical scattering on signal degradation and naturally eliminates possible interferences (i.e., crosstalks) between excitation and detection, which is a common problem in fluorescence microscopy due to the overlap between the excitation and fluorescence spectra. 
Unique for optical absorption imaging, OR-PAM has demonstrated broad biomedical applications since its invention, including, but not limited to, neurology5, 6, ophthalmology7, 8, vascular biology9, and dermatology10. In this video, we teach the system configuration and alignment of OR-PAM as well as the experimental procedures for in vivo functional microvascular imaging.

Optical-resolution photoacoustic microscopy (OR-PAM) is an emerging technology capable of imaging optical absorption contrasts in vivo with cellular resolution and sensitivity. Here, we provide a visualized instruction on the experimental protocols of OR-PAM, including system configuration, system alignment, typical in vivo experimental procedures, and functional imaging schemes.

Optical microscopy, providing valuable insights at the cellular and organelle levels, has been widely recognized as an enabling biomedical technology. As ...

Continuous High-resolution Microscopic Observation of Replicative Aging in Budding Yeast

We demonstrate the use of a simple microfluidic setup, in which single budding yeast cells can be tracked throughout their entire lifespan. The microfluidic chip exploits the size difference between mother and daughter cells using an array of micropads. Upon loading, cells are trapped underneath these micropads, because the distance between the micropad and cover glass is similar to the diameter of a yeast cell (3-4 &mu;m). After the loading procedure, culture medium is continuously flushed through the chip, which not only creates a constant and defined environment throughout the entire experiment, but also flushes out the emerging daughter cells, which are not retained underneath the pads due to their smaller size. The setup retains mother cells so efficiently that in a single experiment up to 50 individual cells can be monitored in a fully automated manner for 5 days or, if necessary, longer. In addition, the excellent optical properties of the chip allow high-resolution imaging of cells during the entire aging process.

We describe here the operation of a microfluidic device that allows continuous and high-resolution microscopic imaging of single budding yeast cells during their complete replicative and/or chronological lifespan.

We demonstrate the use of a simple microfluidic setup, in which single budding yeast cells can be tracked throughout their entire lifespan. The ...

Characteristics of Precipitation-formed Polyethylene Glycol Microgels Are Controlled by Molecular Weight of Reactants

This work describes the formation of poly(ethylene glycol) (PEG) microgels via a photopolymerized precipitation reaction. Precipitation reactions offer several advantages over traditional microsphere fabrication techniques. Contrary to emulsion, suspension, and dispersion techniques, microgels formed by precipitation are of uniform shape and size, i.e.&#160;low polydispersity index, without the use of organic solvents or stabilizers. The mild conditions of the precipitation reaction, customizable properties of the microgels, and low viscosity for injections make them applicable for in vivo purposes. Unlike other fabrication techniques, microgel characteristics can be modified by changing the starting polymer molecular weight. Increasing the starting PEG molecular weight increased microgel diameter and swelling ratio. Further modifications are suggested such as encapsulating molecules during microgel crosslinking. Simple adaptations to the PEG microgel building blocks are explored for future applications of microgels as drug delivery vehicles and tissue engineering scaffolds.

This work describes the formation of poly(ethylene glycol) (PEG) microgels via a photopolymerized precipitation reaction. Increasing the PEG molecular weight increased microgel diameter and swelling ratio. Simple adaptations to the PEG microgel precipitation reaction are explored for future applications of microgels as drug delivery vehicles and tissue engineering scaffolds.

This work describes the formation of poly(ethylene glycol) (PEG) microgels via a photopolymerized precipitation reaction. Precipitation reactions offer ...

High-resolution Spatiotemporal Analysis of Receptor Dynamics by Single-molecule Fluorescence Microscopy

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., kinetics, coexistence of different states and populations, transient interactions), which are typically hidden in ensemble measurements, such as those obtained with standard biochemical or microscopy methods. Thus, dynamic events, such as receptor-receptor interactions, can be followed in real time in a living cell with high spatiotemporal resolution. This protocol describes a method based on labeling with small and bright organic fluorophores and total internal reflection fluorescence (TIRF) microscopy to directly visualize single receptors on the surface of living cells. This approach allows one to precisely localize receptors, measure the size of receptor complexes, and capture dynamic events such as transient receptor-receptor interactions. The protocol provides a detailed description of how to perform a single-molecule experiment, including sample preparation, image acquisition and image analysis. As an example, the application of this method to analyze two G-protein-coupled receptors, i.e., &#946;2-adrenergic and &#947;-aminobutyric acid type B (GABAB) receptor, is reported. The protocol can be adapted to other membrane proteins and different cell models, transfection methods and labeling strategies.

This protocol describes how to use total internal reflection fluorescence microscopy to visualize and track single receptors on the surface of living cells and thereby analyze receptor lateral mobility, size of receptor complexes as well as to visualize transient receptor-receptor interactions. This protocol can be extended to other membrane proteins.

Single-molecule microscopy is emerging as a powerful approach to analyze the behavior of signaling molecules, in particular concerning those aspect (e.g., ...

Preparation of Mica Supported Lipid Bilayers for High Resolution Optical Microscopy Imaging

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction with other compounds, such as drugs or peptides. However SLB characteristics differ depending on the support used. 
Commonly used techniques for SLB imaging and measurements are single molecule fluorescence microscopy, FCS and atomic force microscopy (AFM). Because most optical imaging studies are carried out on a glass support, while AFM requires an extremely flat surface (generally mica), results from these techniques cannot be compared directly, since the charge and smoothness properties of these materials strongly influence diffusion. Unfortunately, the high level of manual dexterity required for the cutting and gluing thin slices of mica to the glass slide presents a hurdle to routine use of mica for SLB preparation. Although this would be the method of choice, such prepared mica surfaces often end up being uneven (wavy) and difficult to image, especially with small working distance, high numerical aperture lenses. Here we present a simple and reproducible method for preparing thin, flat mica surfaces for lipid vesicle deposition and SLB preparation. Additionally, our custom made chamber requires only very small volumes of vesicles for SLB formation. The overall procedure results in the efficient, simple and inexpensive production of high quality lipid bilayer surfaces that are directly comparable to those used in AFM studies.

We present a method of preparing mica supported lipid bilayers for high resolution microscopy. Mica is transparent and flat on an atomic scale, but rarely used in imaging because of handling difficulties; our preparation results in even deposition of the mica sheet, and reduces the material used in bilayer preparation.

Supported lipid bilayers (SLBs) are widely used as a model for studying membrane properties (phase separation, clustering, dynamics) and its interaction ...

A Hormone-responsive 3D Culture Model of the Human Mammary Gland Epithelium

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is limited to cell proliferation and gene expression endpoints. However, in the organism, mammary morphogenesis occurs in a 3D environment. 3D culture systems help bridge the gap between monolayer cell culture (2D) and the complexity of the organism. Herein, we describe a 3D culture model of the human breast epithelium that is suitable to study hormone action. It uses the commercially available hormone-responsive human breast epithelial cell line, T47D, and rat tail collagen type 1 as a matrix. This 3D culture model responds to the main mammotropic hormones: estradiol, progestins and prolactin. The influence of these hormones on epithelial morphogenesis can be observed after 1- or 2-week treatment according to the endpoint. The 3D cultures can be harvested for analysis of epithelial morphogenesis, cell proliferation and gene expression.

We describe a 3D culture model of the human breast epithelium that is suitable to study hormone action.

The process of mammary epithelial morphogenesis is influenced by hormones. The study of hormone action on the breast epithelium using 2D cultures is ...

Selective Cell Elimination from Mixed 3D Culture Using a Near Infrared Photoimmunotherapy Technique

Recent developments in tissue engineering offer innovative solutions for many diseases. For example, tissue engineering using induced pluripotent stem cell (iPS) emerged as a new method in regenerative medicine. Although this tissue regeneration is promising, contamination with unwanted cells during tissue cultures is a major concern. Moreover, there is a safety concern regarding tumorigenicity after transplantation. Therefore, there is an urgent need for eliminating specific cells without damaging other cells that need to be protected, especially in established tissue. Here, we present a method for specific cell elimination from a mixed 3D cell culture in vitro with near infrared photoimmunotherapy (NIR-PIT) without damaging non-targeted cells. This technique enables the elimination of specific cells from mixed cell cultures or tissues.

Eliminating specific cells without damaging other cells is extremely difficult, especially in established tissue, yet there is an urgent need for a cell elimination method in the tissue engineering field. Here, we present a method for specific cell elimination from a mixed 3D cell culture using near infrared photoimmunotherapy (NIR-PIT).

Recent developments in tissue engineering offer innovative solutions for many diseases. For example, tissue engineering using induced pluripotent stem ...

Isolation and Decellularization of a Whole Porcine Pancreas

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an allogeneic or xenogeneic source with human cells. A demonstration of methods for the efficient dissection, decellularization, and recellularization of porcine pancreas might benefit the field. Akin to human pancreases, porcine pancreases have a special anatomical arrangement with three lobes (splenic, duodenal, and connection) rounded by the duodenum and small intestine. The duodenal lobe of the pancreas connects to the duodenum by several small blood vessels. Tissue engineering of the pancreas is complicated because of its exocrine and endocrine nature. In this paper, we show a detailed protocol to dissect the whole porcine pancreas and decellularize it with detergents while saving its structure and some extracellular matrix components. To achieve complete perfusion, the aorta is chosen as inlet and the portal vein as outlet. The other blood vessels (hepatic artery, splenic vein, splenic artery,&#160;mesenteric artery and vein tree) and bile duct are ligated. To prevent the formation of thrombus, the pig is heparinized and, immediately after dissection, the organ is flushed with cold heparin. To inhibit the action of exocrine enzymes, the pancreas decellularization is set at 4 &#176;C. The decellularization is performed by perfusion of Triton X-100, sodium deoxycholate, and deoxyribonuclease, with an intermittent and final extensive washing. With a successful decellularization, the pancreas appears white, and a histological evaluation with hematoxylin and eosin shows an absence of nuclei with a preserved extracellular matrix structure. Thus, the proposed method can be used to successfully dissect and decellularize whole porcine pancreas.

Tissue engineering of the whole pancreas is a challenge because of its exocrine and endocrine functions. We show a method for the dissection of an intact porcine pancreas and the process of successful decellularization by perfusion of detergents Triton X-100, sodium deoxycholate, and deoxyribonuclease.

Tissue engineering of the whole pancreas can improve current treatments for diabetes mellitus. The ultimate goal is to tissue engineer pancreas from an ...