Observing Islet Function and Islet-Immune Cell Interactions in Live Pancreatic Tissue Slices

NOTES: All experimental protocols using mice were approved by the University of Florida Animal Care and Use Committee (201808642). Human pancreatic sections from tissue donors of both sexes were obtained via the Network for Pancreatic Organ Donors with Diabetes (nPOD) tissue bank, University of Florida. Human pancreata were harvested from cadaveric organ donors by certified organ procurement organizations partnering with nPOD in accordance with organ donation laws and regulations and classified as "Non-Human Subjects" by the University of Florida Institutional Review Board (IRB) (IRB no. 392-2008), waiving the need for consent. nPOD tissues specifically used for this project were approved as nonhuman by the University of Florida IRB (IRB20140093). The objectives of sections 1-3 of this protocol are to explain how to successfully dissect a mouse, prepare and process the pancreas, and generate live pancreatic tissue slices. Solutions should be prepared ahead of time, and the recipes can be found in Supplemental Table 1. Time is the most critical factor during these protocol steps. Once the mouse has been sacrificed, tissue viability will begin to decline. All three parts of this protocol need to be completed as quickly as possible until all the necessary slices are generated.

1. Preparation for generation of mouse pancreas slices

1. Clamp the blade onto the vibratome blade holder, but do not attach it to the vibratome yet.
2. Melt 100 mL of 1.25% w/v low melting-point agarose in a microwave. Operate the microwave in 1 min intervals, and stop the microwave for 10 s if the agarose solution begins to boil. Repeat this process until the agarose is melted, and a homogeneous solution is produced. Place the bottle in a 37 °C water bath.
   NOTE: The low agarose concentration is to account for the lower density of the mouse pancreas.
3. Fill a 10 mL Luer lock syringe with 3 mL of warm agarose. Fit a 27 G 25 mm needle onto the syringe. Keep the syringe with the capped attached needle in a 37 °C water bath until the solution is to be injected.
   NOTE: A 27 G needle is preferred as it fits securely into the common bile duct of mice between 10-25 g in body weight and allows for the flow of the highly viscous agarose solution. While larger bore needles may be selected for use, smaller (larger gauge) needles are not recommended as these may be more easily clogged with the agarose solution.
4. Add 20 mL of chilled extracellular solution (ECS) to a 10 cm Petri dish.
   NOTE: The ECS does not need to be bubbled at any point.
5. Fill two 10 cm untreated Petri dishes with 15 mL of Krebs-Ringer bicarbonate buffer (KRBH) containing 3 mM D-glucose and soybean trypsin inhibitor at a concentration of 0.1 mg/mL per dish.
   ​NOTE: Throughout this protocol, it is essential that all solutions used for maintaining slices contain soybean trypsin inhibitor to prevent tissue damage caused by pancreatic digestive proteases.

2. Mouse pancreas excision and tissue processing

NOTE: The protocol for excising the pancreas, processing the tissue, and generating slices is modified from Marciniak et al⁵. To ensure tissue viability, minimize the amount of time between pancreas removal and slice generation. All necessary equipment should be prepared in advance and oriented in a manner to allow for rapid progression through the steps below. Bile duct canulation and injection as well as pancreas excision are best performed under a stereoscope.

1. The mouse is deeply anesthetized with isoflurane and sacrificed by cervical dislocation.
   NOTE: Isoflurane is the preferred anesthetization method. A concentration of 5% isoflurane should be used. For example, 0.26 mL should be used with a 1 L chamber¹⁶. A decrease in pancreatic tissue viability was observed when CO₂ was used.
2. Spray the mouse with 70% v/v ethanol liberally to reduce tissue contamination with fur during the dissection and excision. Place the mouse in a dorsal down, ventral up orientation with the anterior side to the left.
3. Using scissors, open the peritoneum and remove the rib cage, taking care not to puncture the heart or adjacent vessels. Use forceps to flip the liver into the chest cavity, and to move the intestines out of the body cavity to expose the common bile duct. Use a Johns Hopkins bulldog clamp to occlude the ampulla of Vater.
4. Retrieve a 10 mL Luer lock syringe preloaded with 3 mL of warm agarose solution from the 37 °C water bath.
   NOTE: Once the syringe with the agarose is removed from the water bath, the pancreas injection needs to be performed quickly before the agarose cools and sets in the syringe.
5. Holding the forceps in the left hand, use them to gently support and stabilize the bile duct for the injection.
6. Hold the syringe in the right hand, insert the needle bevel-side up into the bile duct. Slowly and steadily inject the pancreas. Once the injection starts, the flow cannot be stopped without the agarose hardening in the syringe and in the pancreas.
   NOTE: The volume used will depend on the weight of the mouse. Based on experience, it is recommended that 1 mL of agarose solution be used per 10 g of mouse body weight with a maximum volume of 2 mL. The pancreas should look slightly inflated with a more definitive structure, but not overextended. Over-injection results in islets that become separated from the exocrine tissue and that have a "blown-out" appearance in the slices.
7. Excise the agarose-filled pancreas from the mouse. Using forceps and scissors, cut the pancreas away, starting at the stomach, moving to the intestines, and ending at the spleen. Once cut away, gently remove the injected pancreas with forceps, and place in a 10 cm Petri dish filled with chilled ECS.
8. Use scissors to remove the adipose, connective tissue, fibrotic tissue, and parts of the pancreas that are not injected with agarose.
   NOTE: Parts of the tissue that should be removed will not have strongly established structures and will appear somewhat gelatinous.
9. After trimming the tissue, use scissors to cut it into smaller sections that are approximately 5 mm in diameter while leaving it submerged under ECS. Cut the tissue carefully, taking care to not push the agarose out of the tissue.
10. Remove the pieces of tissue from the ECS, and place them on a two-ply wiper (see the Table of Materials). Gently roll them on the wiper using forceps to remove excess liquid.
11. Using forceps, carefully place the pieces of tissue in a 35 mm Petri dish with no more than 4 pieces per plate. Place the flattest side of the tissue block facing downward. Gently press down on the tissue using forceps.
    NOTE: Make sure there is space, at least a few millimeters, between the pieces of tissue, and that they are not touching the edge of the plate.
12. Slowly pour the 37 °C agarose into the dish, taking care not to pour it directly on to the tissue. Pour enough so that the tissue pieces are completely covered. Make sure there is a layer of agarose above the tissue pieces as this part will be glued to the specimen holder.
13. Carefully transfer the dish with the pieces of tissue to a refrigerator to allow the agarose to set. Ensure the tissue pieces do not shift or start floating. If they do, quickly readjust them using forceps.
    NOTE: Setting of the agarose should only take a few minutes.
14. Once the agarose has set, use a scalpel to cut around the tissue in straight lines to make agarose blocks as if making a grid between the pieces of tissue. Be sure to leave a few millimeters of agarose surrounding all sides of the tissue.
    NOTE: There should not be any tissue protruding from the agarose. Each block should be a cube of approximately 5 mm x 5 mm x 5 mm volume.
15. Use the scalpel to flip out the empty sections of agarose that were cut around the edges of the plate. Remove the blocks with the tissue from the dish by lifting them carefully with forceps.

3. Mouse pancreatic slice generation

1. Using forceps, arrange the blocks on the specimen holder; place them sideways, keeping in mind that they will be flipped onto the super glue. Arrange the blocks so that they do not extend further than the blade width. As the vibratome moves slowly, arrange the blocks so that the blade has to move forward the least possible distance.
   ​NOTE: Two rows of three or four blocks each with a few millimeters between the rows works well. The blocks within the same row can touch each other, but when both rows touch, it can be difficult to retrieve slices as they come off the blocks.
2. Apply a line of super glue on the specimen holder, and use the end of the glue dispenser to spread the glue out into a thin layer. Flip the tissue blocks onto the glue so that the side closest to the tissue faces upward. Gently push down on the blocks, and let the glue dry for three minutes.
3. Attach the blade holder and plate to the vibratome, typically with either a screw or magnet, depending on the vibratome model. Adjust the blade height and distance travelled so that the blade moves over the length of the blocks and just barely above them.
   NOTE: Forceps can be helpful while adjusting the blade height. They can be placed on top of the tissue block to help place the blade as close to the top of the block as possible without touching the block.
4. Ensure that the glue has dried by gently nudging the blocks with forceps, and fill the vibratome tray with chilled ECS until the blade is covered. Set the vibratome to make 120 µm thickness slices at a speed of 0.175 mm/s, a frequency of 70 Hz, and an amplitude of 1 mm.
   NOTE: The vibratome speed can be adjusted depending on the ease of cutting the tissue.
5. Start the vibratome, and watch for when slices start coming off of the tissue blocks. Use 10 cm Graefe forceps or a small No. 4 paintbrush to carefully remove the slices once they float off the block, and place them in 10 cm plates with KRBH containing 3 mM D-glucose and trypsin inhibitor. Pick up the slices by placing a paintbrush or forceps below them and gently lifting the slices. Do not incubate more than 15 slices together in a single plate.
   NOTE: It is normal for the vibratome to have a few passes over the blocks where no slices are made, but these should be minimized for time. Have scissors ready in case the slices do not fully separate from the tissue blocks. If this happens, a corner or edge of the slice will be stuck to the block after the vibratome blade passes. Do not pull off the slice or block when removing the stuck slice.
6. Place the plates with the slices on a rocker at room temperature and at 25 rpm. Let the slices rest at room temperature for an hour. If they are going to be left for longer, place the slices in 15 mL of slice culture medium (see Supplemental Table 1) in an incubator. Incubate slices prepared for same-day studies at 37 °C, and culture slices that are cultured overnight at 24 °C, transferring them to 37 °C at least 1 h before experiments.
   NOTE: Over the long term, the slices have better viability when cultured at 24 °C, although 37 °C is closer to their native physiological environment, probably due to the lower activity of the secreted protease enzymes at lower temperature. Mouse and human pancreatic tissue slices are both cultured at the same temperature and with a maximum of 15 slices per dish. However, the media recipes differ for human and mouse slices. Both formulations are listed in Supplemental Table 1. Additionally, the procedure is the same for generating mouse and human slices with the exception of the mouse pancreas requiring injection with agarose for stabilization. Human slices are acquired through the nPOD Pancreas Slice Program. Both mouse and human slices are 120 µm thick. A variety of experiments can be performed on the slices; choose staining panels that work best for planned experiments.

4. Slice preparation for staining procedures

1. Culture the slices at 37 °C for at least 1 h ahead of the planned experiments. Warm KRBH containing 3 mM D-glucose in a 37 °C water bath. Transfer 2 mL of KRBH containing 3mM D-glucose in a 35 mm dish, and use a paintbrush to gently place the slice in the dish.
2. If the slice is being transferred from medium, wash it twice with KRBH containing 3 mM D-glucose. Carefully aspirate the KRBH with 3 mM D-glucose using a transfer pipette or Pasteur pipette, taking care not to disturb the slice. Keep the slice in the plate with KRBH containing 3 mM D-glucose while the staining panels are prepared.

5. Dithizone staining

NOTE: Although dithizone can be used to stain the islets red, it will kill the slice as it has been found to be cytotoxic to islets¹⁷.

1. Measure 12.5 mg of dithizone, add it to 1.25 mL of dimethylsulfoxide, and take this mixture up in a 50 mL syringe. Fill the syringe to a volume of 25 mL using Hanks Balanced Salt Solution, and attach a filter to the end of the syringe. Aliquot 2 mL of KRBH with 3 mM D-glucose, and add 2 drops of the filtered dithizone solution from the 50 mL syringe into a 35 mm dish.
2. Using a paintbrush, carefully place a slice in a 35 mm petri dish. Image the slice with the islets indicated by red dithizone staining using a stereomicroscope.

6. Viability staining

NOTE: This section of the protocol describes how to assess slice viability using calcein-AM and blue-fluorescent SYTOX Blue (see the Table of Materials). Calcein-AM should be used at a concentration of 4 µM and SYTOX Blue at 1 µM.

1. Aliquot 2 mL of KRBH containing 3 mM D-glucose, and add 2 µL of calcein-AM dye and SYTOX Blue to separate aliquots. Vortex the mixtures for 5 s.
2. Add 200 µL of KRBH containing 3 mM D-glucose and calcein-AM dye to each well of an 8-well chambered coverglass.
   NOTE: Alternative plates and/or imaging chambers other than an 8-well chambered coverglass can be used.
3. Using a paintbrush, carefully place a slice in each well of the plate, and transfer the plate with the slices to a 37 °C incubator for 20 min. Wash the slices twice with KRBH containing 3 mM D-glucose. Carefully aspirate the KRBH using a transfer or Pasteur pipette, taking care not to disturb the slice.
4. Place the slice in a 35 mm coverglass-bottom Petri dish containing 2 mL of KRBH with 3 mM D-glucose and 2 µL of the SYTOX Blue at a concentration of 1 µL per 1 mL of solution. Cover the slice with a slice anchor, ensuring that the side with the harp faces downward. Take images of the slice.
   NOTE: If the slice anchor keeps floating, wet it on both sides with KRBH containing 3 mM D-glucose to submerge it in the solution. It is critical to always maintain the slices in solutions containing protease inhibitor, even during dye loading. Viability stains used can be adapted for the specific experiment or microscope setup.

7. Slice Ca²⁺ indicator staining

NOTE: This section of the protocol describes how to stain slices for Ca²⁺ recordings using Oregon Green 488 BAPTA-1, AM and SYTOX Blue in mouse slices (see the Table of Materials). The Oregon Green 488 BAPTA-1, AM should be used at a concentration of 5.6 µM and the SYTOX Blue at 1 µM. In human slices, Fluo-4-AM should be used at a concentration of 6.4 µM.

1. Aliquot 2 mL of KRBH containing 3 mM D-glucose, add 7 µL of the Oregon Green 488 BAPTA-1, AM and vortex the mixture for 5 s.
   NOTE: For human tissue slices, use Fluo-4-AM instead of the Oregon Green 488 BAPTA-1, AM. The Fluo-4-AM is preferable because it is brighter when the intracellular Ca²⁺ concentration increases; however, it does not load well in mouse pancreatic tissue. The protocol is the same as described above for Fluo-4-AM with the exception that Fluo-4-AM only needs to be incubated for 30 min.
2. Add 200 µL of KRBH containing 3mM D-glucose and the dye to each well of an 8-well chambered coverglass. Using a paintbrush, carefully place a slice in each well of the chambered coverglass. Transfer the chambered coverglass with the slices to a 37 °C incubator for 45 min.
3. Wash the slices twice with KRBH containing 3 mM D-glucose. Carefully aspirate the KRBH with 3 mM D-glucose using a transfer or Pasteur pipette, taking care to not disturb the slice.
4. Place a slice in an imaging plate or chamber with KRBH containing 3 mM D-glucose and SYTOX Blue at a concentration of 1 µL per 1 mL, and cover with a slice anchor, ensuring that the harp faces downward. Take images of the slice.
   NOTE: In this protocol, a 35 mm dish filled with 2 mL of KRBH containing 3 mM D-glucose and 2 µL of SYTOX Blue was used. If the slice anchor keeps floating, wet it on both sides with KRBH containing 3 mM D-glucose to submerge it in solution.

8. Mouse slice Ca²⁺ recordings

NOTES: The following section describes how to perform Ca²⁺ recordings on mouse pancreatic tissue slices using the Oregon Green 488 BAPTA-1, AM and SYTOX Blue. Imaging was performed on a confocal laser-scanning microscope (see the Table of Materials for details). The lasers used were 405 nm for the SYTOX Blue, 488 nm for the Oregon Green 488 BAPTA-1, AM, and 638 nm for reflectance. A HyD detector was used for the Oregon Green 488 BAPTA-1, AM. Photomultiplier tube (PMT) detectors were used for reflectance and the SYTOX Blue. The Ca²⁺ imaging protocol is the same for human pancreatic tissue slices except that Fluo-4-AM was used as the indicator. Laser power levels, gain, and pinhole size should be adjusted based on the sample and particular islet imaged. Typically, a pinhole of 1.5 airy units and a laser power of 1% are good starting points.

1. At least 1 h before recording, switch on the microscope and equilibrate the stage-top or cage-style incubator to 37 °C. Place the 35 mm coverglass-bottom Petri dish containing the slice on the stage after removing the lid. Focus by setting the microscope to the 10x objective and brightfield mode. Locate islets using brightfield by looking for orange-brown ovals within the slice.
2. Once a probable islet is located, switch the microscope to the confocal imaging mode. To confirm islets by reflectance, switch on the 638 nm laser, set the laser power between 1% and 2%, and switch off the 638 nm notch filter that would normally remove backscattered light. Set the detection limits on the PMT detector to a bandwidth of approximately 20 nm centered around 638 nm.
   NOTE: Exercise caution as operating the microscope in reflectance mode could damage the detector. Due to the high granularity of the endocrine tissue, reflected light can now be used to locate islets. The islets will appear as groups of brightly backscattering granular cells on this reflectance channel.
3. To view the SYTOX Blue, switch on the 405 nm laser and PMT detector, and set the laser power between 1% and 2%. Center the islet of interest in the field of view using the X and Y knobs of the stage controller. Once an islet of interest is located and confirmed by backscatter, switch to the 20x objective, and zoom in so the islet takes up most of the frame.
4. Take a z-stack of the islet with a z-step size of 1.5 µm. Find the best optical section of the islet where most of the cells are alive (negative for the SYTOX Blue) and well-loaded with the Oregon Green 488 BAPTA-1, AM or Fluo-4-AM.
   NOTE: It is not unusual to see cells that are overloaded with a large amount of dye and that are very bright. These may be dying pancreatic cells in which Ca²⁺ storage in the endoplasmic reticulum may be released, resulting in high levels of loading; these are not ideal cells to record. Look for cells that have clearly loaded the dye, but are not oversaturating the detector so that an increase in brightness that occurs when cytosolic Ca²⁺ levels fluctuate is visible. Dye loading of islets within slices is variable; however, the dye typically loads well through ~10-15 µm of the islet. However, dye loading can be difficult to visualize if the cells are deep in the tissue.
5. To prevent dye fading during the recording, ensure that the laser power on the 488 nm channel does not exceed 2%. Increase the pinhole to 2 airy units to collect more signal with lower excitation power.
6. Set the microscope to record in XYZT mode. Optimize the settings to reduce the frame rate to 2 s or less per frame.
   NOTE: Settings adjustments that can be made to help decrease the frame rate include turning on bidirectional scanning, decreasing or turning off the line averaging, and increasing scanning speed.
7. Once the settings have been optimized, record several minutes of basal activity.
   NOTE: Another good indicator of tissue viability is if the cells appear active and are visibly flashing during this recording of basal activity.
8. Add 100 µL of 20x concentrated glucose and KCl in KRBH to the plate using a 200 µL micropipette at the given time points to achieve a final concentration of 16.7 mM glucose or 30 mM KCl.
   NOTES: Add the solutions carefully, taking care not to disturb the slice during the recording. Be sure not to bump the plate with the micropipette. It is typical to see the tissue contract in response to these stimulations. The Ca²⁺ flux recordings were processed and quantified in ImageJ¹⁸. Using ImageJ software, the staining intensity of the Oregon Green 488 BAPTA-1, AM was measured in the cells by manually selecting regions of interest (ROIs). The fluorescence intensity from these ROIs was calculated by dividing the fluorescence values at later timepoints by the initial fluorescence values of the cells (F/F₀). A perfusion system can be used along with a specialized imaging chamber to administer the solutions to slices dynamically as opposed to adding them manually. Perfusion system and imaging chamber recommendations can be found in the Table of Materials.

9. Staining of mouse T cells in live pancreatic slices

NOTE: This section of the protocol describes how to stain immune cells within mouse slices. The mouse strain used is the NOD.Rag1^-/-.AI4^α/β as this model consistently develops disease with significant insulitis. The CD8⁺ T cells in this mouse all target an epitope of insulin, allowing the use of a phycoerythrin (PE)-labelled insulin-D^b tetramer¹⁵. The CD8 antibody should be used at a concentration of 1:20 and the insulin tetramer at 1:50.

1. Aliquot 100 µL of KRBH containing 3 mM D-glucose, add 2 µL of PE insulin tetramer and 5 µL of allophycocyanin (APC) CD8 antibody, and vortex the mixture for 5 s.
2. Add 100 µL of KRBH containing 3 mM D-glucose and the tetramer and antibody to a well of an 8-well chambered coverglass. Using a paintbrush, carefully place a slice in the well of the chambered coverglass. Transfer the chambered coverglass with the slice to a 37 °C incubator for 30 min.
3. Wash the slice twice with 2 mL KRBH containing 3 mM D-glucose. Carefully aspirate the KRBH with 3 mM D-glucose using a transfer or Pasteur pipette, taking care not to disturb the slice.
4. Place the slice in a 35 mm coverglass-bottom Petri dish containing KRBH with 3 mM D-glucose and the SYTOX Blue at a concentration of 1 µL per 1 mL, and cover with a slice anchor, placing the side with the harp facing downward. Take images of the slice.
   ​NOTE: If the slice anchor keeps floating, wet it on both sides with KRBH containing 3 mM D-glucose to submerge it in the solution. The diluted antibody and tetramer can be reused once. After staining two slices, a fresh antibody mixture should be made.

10. Recording of mouse immune cells

NOTE: The following section describes how to perform immune cell recordings on mouse pancreatic tissue slices using CD8 antibody, PE insulin tetramer, and SYTOX Blue. The imaging setup is as described in section 8. Recordings were made at 800 × 800 pixel resolution. The lasers used were 405 nm for the SYTOX Blue, 488 nm for the insulin tetramer, and 638 nm for CD8 antibody and reflectance. HyD detectors were used for CD8 antibody and PE insulin tetramer. PMT detectors were used for reflectance and the SYTOX Blue. The immune cell imaging protocol is the same for human pancreatic tissue slices except for the use of different antibodies and antigen-complexed HLA-multimers for human tissue. For both insulin tetramer staining in mouse tissue and HLA-multimer staining in human tissue, an immune cell co-stain should be used to verify the presence of the specific antigen-reactive T cells. Here, an anti-CD8 antibody was used. Antibodies, such as anti-CD3 or anti-CD4, can also be used depending on the target cell population.

1. At least 1 h before recording, switch on the microscope, and equilibrate the stage-top incubator to 37 °C. Secure the 35 mm coverglass-bottom Petri dish containing the slice on the stage. Focus the microscope by setting the 10x objective in the brightfield mode. Locate the islets using the brightfield mode by looking for orange-brown colored ovals within the slice.
2. Switch the microscope to confocal imaging by pressing the CS button on the microscope's touch screen controller. To view islets by reflectance, turn on the 638 laser and PMT detector, set the laser power between 1% and 2%, and turn off the notch filters.
   NOTE: Due to the increased granularity of the endocrine tissue, reflected light can now be used to locate islets. The islets will appear as groups of bright granular cells on this channel.
3. To view the SYTOX Blue, CD8 antibody, and insulin tetramer, check that the laser power is between 1% and 2%. Use the following settings to view each of the three: for the SYTOX Blue, turn on the 405 nm laser and PMT detector; for the CD8 antibody, turn on the HyD detector; and to view the insulin tetramer, turn on the 488 nm laser and HyD detector.
4. Center the islet of interest in the field of view using the X and Y knobs of the stage controller. Once an islet of interest is located, switch to the 20x objective, and zoom in so the islet takes up most of the frame. Take a z-stack of the islet with a z-step size of 1.5 µm. Find the best optical sections (a series of between 5 to 10 sections) of the islet where most of the cells are alive (negative for the SYTOX Blue) and any surrounding immune cells are in focus.
   NOTE: Try to find frames where there are multiple CD8-positive and insulin tetramer-positive cells surrounding or infiltrating the islet.
5. Set the microscope to record in XYZT mode. Optimize the settings to record a Z-stack of the selected steps every 20 min over a period of several hours.
   NOTE: If possible, it is best to do these recordings in an imaging chamber where temperature and CO₂ levels can be controlled, particularly when recording for over four hours. In the case of overnight recording, excess antibody can be added to the media to compensate for T cell receptor cycling and dye fading. Additionally, different fluorophores can be used for the T cell antibodies. Based on experience, antibodies in the far-red range work best for T cells.