The authors would like to thank Wei Zhang from Guanhao Biotech for providing the decellularized scaffolds. We thank Xiao-hong Peng for the helpful discussions. This research was financially supported by the National Natural Science Foundation of China (Project No.31322021).

All experiments were approved by Peking University Institutional Animal Care and Use Committee (IACUC, IACUC no. COE-LuoY-1).
1. Islet Isolation
<ol>
	<li>Preparation of reagents and equipment.
	<ol>
		<li>Reconstitute collagenase P powder (2 U/mg) in HBSS to make a 5 mg/mL solution and filter it through a 0.22 &#181;m filter to remove the bacteria. Prepare 0.6 mL-aliquot solutions of collagenase P in 15-mL conical tubes and store at -20 &#176;C. 
		NOTE: During use, each aliquot is diluted with HBSS to give 6-mL working solutions with final concentrations of 0.5 mg/mL, or 1 U/mL (enough for treating 3 mice). The working solution is kept on ice for immediate use within 1 h. The working solution should not be restored or re-frozen for additional usage.</li>
		<li>Prepare neutralization solution by adding FBS (2.5%) and P/S (1%) into HBSS; keep the solution on ice. Prepare 60 mL of neutralization solution to treat 6 mice.</li>
		<li>For the islet culture medium, add D-glucose (7 mM), FBS (10%), and P/S (1%) to RPMI 1640 medium.</li>
		<li>Autoclave the surgery instruments at 115 &#176;C for 30 min and 15 psi of pressure.</li>
	</ol>
	</li>
	<li>Inflation of the pancreas.
	<ol>
		<li>Prepare 12 mL of collagenase working solution, as specified in step 1.1.1, for 6 mice (12 weeks old). Fill the 10 mL syringe with 9 mL of working solution and connect the syringe to the 27 G intravenous needle. Store the syringe on ice and use the solution within 1 h.</li>
		<li>Euthanize the mouse by cervical dislocation. Place the mouse in supine position on a paper towel, with the tail pointing towards the operator. Spray the whole body with 70% ethanol, making it completely wet.</li>
		<li>Make a V-incision, starting from the genital area and extending to the diaphragm, using dissection scissors and forceps. Fold the skin over the chest to completely reveal the abdominal cavity.</li>
		<li>Move the bowel to the right side of the mouse and expose the pancreas and common bile duct. Grab the duodenum carefully with forceps and pull it until the bile duct is taut (Figure 1A-1 and A-2).</li>
		<li>Find the portal vein and bile duct leading into the liver. Clamp the portal vein and bile duct with a microscopic hemostatic clamp. 
		NOTE: The clamping prevents excessive bleeding and the entry of the collagenase into the liver.</li>
		<li>While still holding the intestine with the forceps, find the location of the ampulla that connects the bile duct and the duodenum. Insert the 27G intravenous needle into the common bile duct through the ampulla (Figure 1B-1 and B-2).</li>
		<li>Dispense about 200 &#181;L of collagenase working solution to check if the cannulation is all the way through bile duct. If the collagenase solution begins to fill the duct ( Figure 1C-1 and C2), the needle is in the lumen of the duct; clamp the duct segment containing the needle using an artery hemostatic clamp and dispense the rest of the 2 mL of solution at a slow and constant rate within 1 min (Figure 2A-1, A-2, and B-1). 
		NOTE: If the tissue surrounding the duct begins to inflate (Figure 2B-2), the needle has poked through the wall of the bile duct, making it necessary to reposition the needle and try cannulating the bile duct again. Similarly, if the duodenum begins to inflate (Figure 2C-1 and C-2), the artery hemostatic clamp needs to be adjusted and the bile duct re-cannulated. As the collagenase solution fills up the pancreas, the tissue near the duodenum inflates first, followed by the region near the pancreatic tail. The perfusion of the pancreatic tail (the splenic lobe) is important to maximize the islet yield.</li>
		<li>After the complete inflation of the pancreas (Figure 2C-1), push the bowel to the left side of the mouse and remove the pancreas by starting at the descending colon. Use the forceps to lift the bowel and separate it from the pancreas with another pair of forceps. Continue to remove the pancreas until it is unattached from the top of the stomach. Finally, lift the pancreas out of the abdomen and cut it free from the remaining spleen. 
		NOTE: The whole separation should be performed quickly, as the digestion continues during the removal process.</li>
		<li>Place the pancreas in an empty 15-mL conical tube and leave it on ice. Repeat the above procedure for the remaining mice. All pancreas should be further processed within 1 h by promptly following step 1.3 to prevent over-digestion by collagenase P.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/54995/54995fig1.jpg" /> 
Figure 1: Photographs showing the cannulation of the bile duct and the perfusion of the pancreas with collagenase solutions. (A1) Pulling the duodenum until the bile duct is taut. (ampulla: the triangular, milky area on the surface of the duodenum; bile duct: the cord-like milky structure on the surface). (B1) Inserting the needle into the bile duct from the ampulla. (C1) Inflating the pancreas with the injection of enzyme. (A2, B2, and C2) Cartoon images of the procedures shown in A1, B1, and C1, respectively. <a href="//cloudfront.jove.com/files/ftp_upload/54995/54995fig1large.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/54995/54995fig2.jpg" /> 
Figure 2: Troubleshooting for the cannulation. (A1) The needle tip inserted in the lumen of the bile duct. (A2) The duct filled with enzyme solutions. (B1) The needle inserted in the lumen of the bile duct, and the duct filled with a blue dye. (B2) Due to inappropriate cannulation, the needle is beneath the bile duct, and only an inflated capsule is observed after dispensing the blue dye. (C1) A successful cannulation is evidenced by the distension of pancreas. (C2) Due to inappropriate clamping, blue dye enters the duodenum and causes distention. <a href="//cloudfront.jove.com/files/ftp_upload/54995/54995fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<ol start="3">
	<li>Digestion and purification of islets.
	<ol>
		<li>Incubate the conical tubes containing the perfused pancreas at 37 &#176;C for 17 min. 
		NOTE: The incubation time may vary with the age and species of the animal.</li>
		<li>Terminate the digestion by adding 7 mL of neutralization solution and putting the tubes on ice.</li>
		<li>Dissociate the tissue by shaking the tubes vigorously (e.g., 20 times in 10 s) until fine tissue particles are obtained. 
		NOTE: The islet yield will be low if the pancreas fails to dissociate completely.</li>
		<li>Filter the digested tissue samples through 0.5-mm wire mesh to remove any non-digested tissue chunks. Collect the islet suspensions in a new 50-mL conical tube.</li>
		<li>Centrifuge the tubes for 3 min at 230 x g and 4 &#176;C and pour off the supernatant carefully, without disturbing the tissue pellets.</li>
		<li>Resuspend the pellets from 3 mice in 4 mL of polysucrose density gradient by vortexing gently or pipetting the suspension up and down a few times. Slowly pipette 4 mL of HBSS down the side of the tube to the top of the polysucrose solutions. 
		NOTE: The two solutions should be well-separated layers with a sharp interface. 
		NOTE: The addition of HBSS should be performed with care, without disturbing the polysucrose density gradient at the bottom.</li>
		<li>Centrifuge the suspension for 20 min at 900 x g and 4 &#176;C by selecting a very slow acceleration rate. End the centrifugation with no brake. 
		NOTE: This is a step to purify the islets from the exocrine cells, with most of the islets migrating to the interface between the layers of polysucrose and HBSS and most exocrine cells settling at the bottom.</li>
		<li>Remove the entirety of the 8-mL supernatant solutions using a large-bore 15-mL pipette. Pass the solutions through an inverted 70-&#181;m strainer. 
		NOTE: The islets will be further purified and retained by the strainer, while the exocrine cells will pass through the filter. 
		Caution: The polysucorse density gradient is cell-toxic; the filtration will help islets get rid of the polysucrose.</li>
		<li>Pipette 2 mL of cold neutralization solution into a 35-mm Petri dish. Invert the cell strainer right-side-up, dip the surface retaining the islets in the solution, and gently shake to release the islets.</li>
		<li>Hand-pick the islets under a microscope using a 20-&#181;L pipette with a white tip. 
		NOTE: The islets are yellowish, compact cell aggregates (Figure 3A), while the contaminating exocrine tissue or cells have a blackish and loose structure under the dissecting microscope.</li>
		<li>Place about 200 islets in a 35-mm dish with 2 mL of culture medium and incubate at 37 &#176;C in an incubator supplied with 5% CO2 for 12 h. 
		NOTE: Usually 150-300 islets can be harvested from one 12-week-old C57BL/6J mouse. Islets are prone to forming aggregates during culture, and the big islets (&#62;300 &#181;m) are susceptible to central necrosis (Figure 1B, inset). Shake the suspension well to evenly distribute the islets in the dish and to reduce clumping.</li>
	</ol>
	</li>
</ol>
2. Islet Culture on the Scaffold
NOTE: DCS has a porosity of about 79%, a thickness of about 0.6 mm, and a pore size ranging from 12 to 300 &#181;m.
<ol>
	<li>Cut the scaffolds into 7-mm disks, soak them in 70% ethanol, and wash them with HBSS. Place the scaffolds in the 24-well tissue culture inserts. 
	NOTE: When fresh islets are recovered after overnight culture, the islets appear bright and tight, with smooth borders (Figure 3B).</li>
	<li>Swirl the dish and collect the islets from the center of the dish using a 20 &#181;L pipette with a white tip. Transfer the islets to the scaffolds (250 islets/scaffold) using a pipette and add 2 mL of culture medium to the well. Culture the islets for 12 h before transplantation.</li>
</ol>
3. Islets Transplantation at the EFP Site
<ol>
	<li>Diabetes induction in recipient mice.
	<ol>
		<li>Place C57BL/6J mice (more than 10 weeks old) in a fresh cage with a water supply but no food. Fast the mice for 10 h before the induction of diabetes.</li>
		<li>Prepare buffer solutions with pH values of 4.2-4.5 by mixing 0.1 M citric acid with 0.1 M sodium citrate. Dissolve STZ (10 mg/mL) in the freshly prepared buffer and sterilize the solution by passaging it through a 0.22 &#181;m filter. 
		NOTE: STZ is light-sensitive and loses activity within 10 min; always prepare the fresh STZ solution right before injection.</li>
		<li>Inject each mouse intraperitoneally with STZ at a dose of 140-150 mg/kg for each C57BL/6J mouse. 
		NOTE: The dose varies depending upon the age and species of the animal. It is recommended to perform a small-scale dose optimization test for the given species before starting the formal experiment.</li>
		<li>Collect the tail vein blood and monitor the blood glucose with a glucose meter on days 2, 3, and 4 post-STZ injection. 
		NOTE: When the animals are hyperglycemic (non-fasting blood glucose &#62; 16.7 mM) on two consecutive days, they are ready for islet transplantation.</li>
	</ol>
	</li>
	<li>Transplantation of islets to the EFP site.
	<ol>
		<li>Anesthetize the mice with pentobarbital delivered intraperitoneally (50 mg/kg). Place the mouse in supine position on a paper towel, with the tail pointing towards the operator. Shave the abdomen extensively to remove the fur from around the incision site. Tape down the four limbs and swab the skin completely with alternating alcohol and iodophor wipes moved in a circular fashion to sterilize the incision site. Drape the mouse with a sterile drape, only allowing access to the incision area. Make a 7 mm incision through the peritoneal wall in the midline, close to the genital area. 
		NOTE: All instruments used during the surgery, including gloves, should be sterile.</li>
		<li>Gently grab and remove the EFP from the abdominal cavity using forceps. Spread the EFP on wetted sterile gauze. Place the scaffold containing islets on the EFP and fold the EFP to wrap the transplant. Secure the direct contact between the islets and EFP by suturing the EFP with absorbable 6-0 sutures. Wet the surface of the EFP with sterile saline using a soaked cotton swab to prevent the EFP from drying out.</li>
		<li>Gently place the EFP back in the abdominal cavity. Close the incision by suturing the peritoneal wall and clamping the dermal layer with wound clips.</li>
		<li>Inject buprenorphine (0.1 mg/kg) subcutaneously as an analgesic.</li>
		<li>Inject 1 mL of saline subcutaneously to prevent dehydration.</li>
		<li>Place the mouse in a cage on a heating pad until the mouse recovers from anesthesia.</li>
		<li>Check the non-fasting glucose levels two days later by collecting blood from the tail vein. If retrieving the graft, repeat the above transplantion procedures, gently pull out the EFP, use 3-0 suture to ligate the end of the fat pad adjacent to the epididymis, occlude the blood vessels, explant the EFP graft, and preserve it for histology. 
		NOTE: Explanting the islets should render the recipient mice hyperglycemic again within 3 days, confirming the function of the graft.</li>
	</ol>
	</li>
</ol>

<ol>
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The authors have nothing to disclose.

Pancreas perfusion and digestion time are two key parameters that affect islet yield and quality. Moskalewski first reported the use of a crude collagenase mixture to digest minced guinea pig pancreas11. Lacy et al. reported the injection of enzymes into the duct system to perfuse the pancreas, which greatly increased the islet yield12. The ductal perfusion of enzyme allows for the maximal exposure of pancreatic surface area to the enzyme, resulting in a more homog...

Type 1 diabetes mellitus (T1D) is an autoimmune endocrine disorder in which islet cells are ablated by the immune system, rendering patients dependent upon the injection of exogenous insulin for their whole lives. The Edmonton protocol represents a milestone in clinical studies of islet transplantation; islets were infused through the portal vein and transplanted at the intrahepatic site1. However, two main obstacles-inadequate sources of donor islets and poor islet engraftment-prevent the wide success of the islet transplantation2. Usually, islets need to be collected from three cadaveric donors to reverse the hyperglycemic condition of one patient; this is due to the low yield of islet isolation procedures and the islet loss after transplantation. In particular, although the post-transplantation islets were bathed in oxygen-rich blood, the direct contact with blood often evoked the instant blood-mediated inflammatory reaction (IBMIR), which could cause the acute loss of the islets. In the long term, it is thought that the gradual loss of islets in patients accounted for the drop of diabetes reversal rates in the clinical groups, which could reach 90% in the first year and declined to 30% and 10% by 2 and 5 years post-transplantation, respectively3.
Islet transplantation at extrahepatic sites has been an attractive strategy to reduce the direct contact of islets with blood while confining the transplants to more definable locations compared to intrahepatic infusion. Studies have been carried out in the kidney capsule, eye, muscle, fat pads, and subcutaneous spaces over the past years, showing that islets at these sites are able to survive and function to restore normoglycemia4. In addition, the islets at these sites are retrievable, making it possible for biopsy or even for further replacement procedures. Extrahepatic sites therefore demonstrate great potential for clinical transplantation5.
Biomaterial-based scaffolds have been intensively investigated for cell transplantation and tissue engineering. Three-dimensional (3D) scaffolds usually contain porous structures and can serve as cellular templates to generate spatial structure/organization of cells or as reservoirs to provide the controlled release of bioactive cues. Scaffolds have also been fabricated from polymeric materials, such as poly(glycolide-L-lactide)6, poly(dimethylsiloxane)7, and thermoplastic poly(urethane)8, to transplant islets in the EFP. Compared to the direct transplantation of islets, the use of scaffolds was found to reduce islet loss by preventing the leakage of islets into the intraperitoneal cavity9,10, providing mechanical protection and modulating the local inflammatory reaction. The scaffolds thus may be developed to promote islet engraftment at the transplantation sites7.
In this study, we intend to demonstrate a paradigm of islet transplantation in the EFP, carried out in mice models using a DCS. Scaffolds derived from extracellular matrices have attracted great interest in recent years due to the superior biocompatibility and more natural porous structures compared to synthetic products. Here, we describe a robust isolation protocol to obtain pancreatic islets at high yields from C57BL/6J mice. DCSs processed from the bovine pericardium were then seeded with islets, and the grafts were transplanted to the EFP in syngeneic diabetic models. Normoglycemia in mice was achieved within 10 days and was maintained for up to 100 days, until the removal of the grafts.

<table><tbody><tr><td>Dissecting scissor</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Forceps</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>0.5 mm diameter wire mesh</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>70 &amp;mu;m cell strainer</td><td>Falcon</td><td>352350</td><td></td></tr><tr><td>Artery hemostatic clamp</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Microscopic hemostatic clamp</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Hemostatic forceps</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Absorbable 6-0 PGLA sutures&amp;nbsp;</td><td>JINHUAN</td><td></td><td>With needle</td></tr><tr><td>Wound clip</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Cotton swab</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Gauze</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>Sterile drapes</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>10mL syringe</td><td>JINGHUAN</td><td></td><td></td></tr><tr><td>1 mL syringe</td><td>JINGHUAN</td><td></td><td></td></tr><tr><td>27G intravenous needle</td><td>JINGHUAN</td><td>0.45x15 RWSB</td><td></td></tr><tr><td>1.5 mL Eppendorf tube</td><td>Axygen</td><td></td><td></td></tr><tr><td>15mL conical tube</td><td>Corning</td><td>430791</td><td></td></tr><tr><td>50mL conical tube</td><td>Corning</td><td>430829</td><td></td></tr><tr><td>35mm Non-treated&amp;nbsp; Peri-dishes</td><td>Corning</td><td>430588</td><td></td></tr><tr><td>Transwell</td><td>Corning</td><td>3422</td><td></td></tr><tr><td>0.22 &amp;mu;m filter</td><td>Pall</td><td>PN4612</td><td></td></tr><tr><td>10 mL serological pipet</td><td>Corning</td><td>4488</td><td></td></tr><tr><td>Pipet filler S1</td><td>Thermo Scientific</td><td>9501</td><td></td></tr><tr><td>Pipette (2-20&amp;mu;L)</td><td>Axygen</td><td>AP-20</td><td>AXYPET&lt;sup&gt;TM&lt;/sup&gt;</td></tr><tr><td>Dissecting microscope</td><td>Olympus</td><td>SZ61</td><td></td></tr><tr><td>Centrifuge</td><td>Eppendorf</td><td>5810R</td><td></td></tr><tr><td>Hank&amp;rsquo;s balanced salt solution&amp;nbsp;</td><td>Gibco</td><td>C14175500CP</td><td></td></tr><tr><td>Collagenase P</td><td>Roche</td><td>COLLP-RO</td><td></td></tr><tr><td>Histopaque 1077</td><td>Sigma</td><td>10771</td><td></td></tr><tr><td>RPMI 1640</td><td>Gibco</td><td>11879-20</td><td></td></tr><tr><td>FBS</td><td>Gibco</td><td>16000-044</td><td></td></tr><tr><td>D-glucose</td><td>Gibco</td><td>A24940-01</td><td></td></tr><tr><td>Glucose meter</td><td>Roche</td><td></td><td>ACCU-CHEK</td></tr><tr><td>Penicillin-streptomycin</td><td>Gibco</td><td>15140-122</td><td></td></tr><tr><td>Streptozotocin</td><td>Sigma</td><td>V900890</td><td>Vetec&lt;sup&gt;TM&lt;/sup&gt;</td></tr><tr><td>Chloral hydrate</td><td>J&amp;amp;K</td><td>C0073</td><td></td></tr><tr><td>Sodium citrate</td><td>Sigma</td><td>71497</td><td></td></tr><tr><td>Citric acid</td><td>Sigma</td><td>C2404</td><td></td></tr><tr><td>Iodophors</td><td>Ningbo Medical</td><td></td><td></td></tr><tr><td>C57BL/6J, 10-12 weeks old</td><td>VitalRiver</td><td></td><td>Beijing, China</td></tr><tr><td>Decellularized scaffold</td><td>Guanhao Biotec</td><td>131102</td><td>Guangzhou, China</td></tr></tbody></table>

scaffold-supported transplantation of islets in the epididymal fat pad of diabetic mice

Islet transplantation has been clinically proven to be effective at treating type 1 diabetes. However, the current intrahepatic transplantation strategy may incur acute whole blood reactions and result in poor islet engraftment. Here, we report a robust protocol for the transplantation of islets at the extrahepatic transplantation site-the epididymal fat pad (EFP)-in a diabetic mouse model. A protocol to isolate and purify islets at high yields from C57BL/6J mice is described, as well as a transplantation method performed by seeding islets onto a decellularized scaffold (DCS) and implanting them at the EFP site in syngeneic C57BL/6J mice rendered diabetic by streptozotocin. The DCS graft containing 500 islets reversed the hyperglycemic condition within 10 days, while the free islets without DCS required at least 30 days. The normoglycemia was maintained for up to 3 months until the graft was explanted. In conclusion, DCS enhanced the engraftment of islets into the extrahepatic site of the EFP, which could easily be retrieved and might provide a reproducible and useful platform for investigating the scaffold materials, as well as other transplantation parameters required for a successful islet engraftment.

Our clamping method, performed using a microscopic hemostatic clamp, is straightforward and time-saving compared with the suture ligation technique. It took roughly 4 h to isolate and purify about 1,200 islets from 6 mice. The freshly isolated islets typically had a rough periphery under an optical microscope (Figure 3A). Once the islets recovered from the isolation process, they looked bright and tight and acquired a smooth surface. However, the stressful is...

Watch this Scientific Journal Video about Scaffold-supported Transplantation of Islets in the Epididymal Fat Pad of Diabetic Mice at JoVE.com

Scaffold-supported Transplantation of Islets in the Epididymal Fat Pad of Diabetic Mice

This protocol demonstrates murine islet isolation and seeding onto a decellularized scaffold. Scaffold-supported islets were transplanted into the epididymal fat pad of streptozotocin (STZ)-induced diabetic mice. Islets survived at the transplantation site and reversed the hyperglycemic condition.

Islet transplantation has been clinically proven to be effective at treating type 1 diabetes. However, the current intrahepatic transplantation strategy ...

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Research

JoVE Journal

Bioengineering

10.0K Views.  Peking University. This protocol demonstrates murine islet isolation and seeding onto a decellularized scaffold. Scaffold-supported islets were transplanted into the epididymal fat pad of streptozotocin (STZ)-induced diabetic mice. Islets survived at the transplantation site and reversed the hyperglycemic condition.

Video: Scaffold-supported Transplantation of Islets in the Epididymal Fat Pad of Diabetic Mice

A Method for Murine Islet Isolation and Subcapsular Kidney Transplantation

Since the early pioneering work of Ballinger and Reckard demonstrating that transplantation of islets of Langerhans into diabetic rodents could normalize their blood glucose levels, islet transplantation has been proposed to be a potential treatment for type 1 diabetes 1,2. More recently, advances in human islet transplantation have further strengthened this view 1,3. However, two major limitations prevent islet transplantation from being a widespread clinical reality: (a) the requirement for large numbers of islets per patient, which severely reduces the number of potential recipients, and (b) the need for heavy immunosuppression, which significantly affects the pediatric population of patients due to their vulnerability to long-term immunosuppression. Strategies that can overcome these limitations have the potential to enhance the therapeutic utility of islet transplantation.
Islet transplantation under the mouse kidney capsule is a widely accepted model to investigate various strategies to improve islet transplantation. This experiment requires the isolation of high quality islets and implantation of islets to the diabetic recipients. Both procedures require surgical steps that can be better demonstrated by video than by text. Here, we document the detailed steps for these procedures by both video and written protocol. We also briefly discuss different transplantation models: syngeneic, allogeneic, syngeneic autoimmune, and allogeneic autoimmune.

Transplantation of isolated islets has been proposed to be a potential treatment for type 1 diabetes. Here we describe a method to isolate islets from mouse pancreata and transplant them to the subcapsular space of the kidney.

Since the early pioneering work of Ballinger and Reckard demonstrating that transplantation of islets of Langerhans into diabetic rodents could normalize ...

Medicine

Transplantation of Pancreatic Islets Into the Kidney Capsule of Diabetic Mice

Our protocol was developed to cleanly and easily deliver islets or cells under the kidney capsule of diabetic or normal mice. We found that it was easier to concentrate the islets or cells into pellets in the final delivery tubing (PE50) used to transplant the cells under the kidney capsule. This technique provides both speed and ease while reducing any undue stress to the cells or to the mouse.

Loading: Settled, hand picked, islets or pelleted cells are carefully aspirated off the bottom of a 1.5 mL microcentrifuge tube using a p200 pipetteman and a straight, thin-wall pipette tip. A length of PE50 tubing is attached to the pipette tip using a small silicone adapter tubing. Cells are allowed to settle, in the tip, and then are transferred to the PE50 tubing by slowly dialing the pipetteman. Once the cells are near the end of the PE50 tubing, a kink is made and the silicone adaptor tubing is placed over the kink. The PE50 tubing is transferred to a 15 mL conical containing a cut 5 mL pipet, and the PE50 tubing is taped over the side of the 5 mL pipet to prevent curling during centrifuging. Cells are allowed to reach 1,000 rpm and stopped.
Transplantation: Recipient mice are anesthetized, shaved, and cleaned. A small incision is made on the left flank of the mouse and the kidney is exposed. The kidney, fat, and tissue are kept moist with normal saline swab. The distal end of the PE50 is attached to a Hamilton screw drive syringe, containing a pipette tip, using the silicone adaptor tubing. A small nick is made on the right flank side of the kidney, not too large nor too deep. The beveled end of the PE50 tubing, nearest the cells, is carefully placed under the capsule, the tubing is moved around gently to make space while swabbing normal saline; a dry capsule can tear easily. A small air bubble is delivered under the capsule by slowly dialing the syringe screw drive. Islets are then slowly delivered behind the air bubble. Once the islets have been delivered kidney homeostasis is maintained and the knick is cauterized with low heat. The kidney is placed back into the cavity and the peritoneum and skin are sutured and stapled. Mice are immediately treated with Flunixin and Buprenorphine s.q. and placed in a cage on a heating pad.

Our protocol was developed to cleanly and easily deliver islets or cells under the kidney capsule of mice. Cells are concentrated into pellets in the final tubing used for transplanting the cells under the kidney capsule. The ease of this technique reduces stress to the cells and the mouse.

Our protocol was developed to cleanly and easily deliver islets or cells under the kidney capsule of diabetic or normal mice.  We found that it was easier ...

Biology

Quantification of Global Diastolic Function by Kinematic Modeling-based Analysis of Transmitral Flow via the Parametrized Diastolic Filling Formalism

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised the only means available, the development of noninvasive imaging modalities (echocardiography, MRI, CT) having high temporal and spatial resolution provide a new window for quantitative diastolic function assessment. Echocardiography is the agreed upon standard for diastolic function assessment, but indexes in current clinical use merely utilize selected features of chamber dimension (M-mode) or blood/tissue motion (Doppler) waveforms without incorporating the physiologic causal determinants of the motion itself.&#160;The recognition that all left ventricles (LV) initiate filling by serving as mechanical suction pumps allows global diastolic function to be assessed based on laws of motion that apply to all chambers. What differentiates one heart from another are the parameters of the equation of motion that governs filling. Accordingly, development of the Parametrized Diastolic Filling (PDF) formalism&#160;has shown that the entire range of clinically observed early transmitral flow (Doppler E-wave) patterns are extremely well fit by the laws of damped oscillatory motion.&#160;This permits analysis of individual E-waves in accordance with a causal mechanism (recoil-initiated suction) that yields three (numerically) unique lumped parameters whose physiologic analogues are chamber stiffness (k), viscoelasticity/relaxation (c), and load (xo).&#160;The recording of transmitral flow (Doppler E-waves) is standard practice in clinical cardiology and, therefore, the echocardiographic recording method is only briefly reviewed. Our focus is on determination of the PDF parameters from routinely recorded E-wave data.&#160;As the highlighted results indicate, once the PDF parameters have been obtained from a suitable number of load varying E-waves, the investigator is free to use the parameters or construct indexes from the parameters (such as stored energy 1/2kxo2, maximum A-V pressure gradient kxo, load independent index of diastolic function, etc.) and select the aspect of physiology or pathophysiology to be quantified.

Accurate, causality-based quantification of global diastolic function has been achieved by kinematic modeling-based analysis of transmitral flow via the Parametrized Diastolic Filling (PDF) formalism. PDF generates unique stiffness, relaxation,&#160;and load parameters and elucidates &#39;new&#39; physiology while providing sensitive and specific indexes of dysfunction.

Quantitative cardiac function assessment remains a challenge for physiologists and clinicians.&#160;Although historically invasive methods have comprised ...

Intra-Omental Islet Transplantation Using h-Omental Matrix Islet filliNG (hOMING)

Regenerative medicine based on cell therapy represents a new hope for curing disease. Current obstacles include proper in vivo validation of the efficiency of the therapy. For transfer to the recipient body, cells often need to be combined with biomaterials, especially hydrogels. However, validation of the efficacy of such a graft requires the right environment, the right hydrogel, and the right recipient site. The omentum might be such a site. Based on the example of islet transplantation, we developed the hOMING (h-Omental Matrix Islet filliNG) technique, which consists of the injection of the graft inside the tissue, in between the omental layers, to improve islet implantation and survival. To achieve this, islets have to be embedded in a hydrogel with a viscosity that enables its injection using an atraumatic needle. Syringes are loaded with a combination of hydrogel and islets. Several injections are performed inside the omental tissue at different entry points, and the deposition of the islet/hydrogel mixture is made along a line. We tested the feasibility of this innovative approach using dextran beads. The beads were well spread throughout the omental tissue, in close proximity to blood vessels. To test the efficacy of the graft, we transplanted islets into diabetic rats and perform a metabolic follow-up over two months. The transplanted islets exhibited a high rate of re-vascularization around and inside islets, and reversed diabetes. The hOMING technique could be applicable for other types of hydrogel or cell therapy, for cells with high metabolic activity.

Intra-Omental Islet Transplantation Using h-Omental Matrix Islet filliNG hOMING

Here, we present a protocol for in vivo validation of hydrogel-based cell therapy, illustrated by the example of islet transplantation. h-Omental Matrix Islet filliNG (hOMING) implantation allows implantation of a cell-hydrogel mixture between the omental layers, near to blood vessels, to maximize engraftment in a proper metabolic environment.

Regenerative medicine based on cell therapy represents a new hope for curing disease. Current obstacles include proper in vivo validation of the ...

Intraportal Transplantation of Pancreatic Islets in Mouse Model

Pancreatic islet transplantation to reduce hyperglycemia is highly successful in rodents with chemically-induced diabetes. The most common transplantation site in experimental islet transplantation is the kidney capsule. However, as less is known about the interaction of pancreatic islets with blood constituents, it also makes sense to utilize the portal vein approach in experimental islet transplantation.
This protocol demonstrates an intraportal islet transplantation technique in NMRI nude mice. Streptozotocin (180 mg/kg) is injected intraperitoneally to induce hyperglycemia in recipient mice. They are considered as diabetic at a non-fasting blood glucose level greater than 20 mmol/L. One day prior to transplantation, mouse pancreatic islets are isolated from the donor pancreas by collagenase digestion; a minimum of 350 islets are utilized per diabetic recipient. Depending upon the islet isolation yield, two or more donor mice are utilized per recipient. After overnight culture at 37 &#176;C, islets are administered into the recipient liver via the portal vein. After surgery, the mice are protected in red Makrolon houses and observed until are awake. This protocol maintains glycemic control for 120 days in syngeneic mice and 15 days in allogeneic mice.

Pancreatic islet transplantation is a way to achieve normoglycemia in type 1 diabetes. This article focuses on the transplantation technique through the intraportal route to maintain normal glucose levels in diabetic mice.

Pancreatic islet transplantation to reduce hyperglycemia is highly successful in rodents with chemically-induced diabetes. The most common transplantation ...

Inguinal Subcutaneous White Adipose Tissue (ISWAT) Transplantation Model of Murine Islets

Pancreatic islet transplantation is a well-established therapeutic treatment for type 1 diabetes. The kidney capsule is the most commonly used site for islet transplantation in rodent models. However, the tight kidney capsule limits the transplantation of sufficient islets in large animals and humans. The inguinal subcutaneous white adipose tissue (ISWAT), a new subcutaneous space, was found to be a potentially valuable site for islet transplantation. This site has better blood supply than other subcutaneous spaces. Moreover, the ISWAT accommodates a larger islet mass than the kidney capsule, and transplantation into it is simple. This manuscript describes the procedure of mouse islet isolation and transplantation in the ISWAT site of syngeneic diabetic mouse recipients. Using this protocol, murine pancreatic islets were isolated by standard collagenase digestion and a basement membrane matrix hydrogel was used for fixing the purified islets in the ISWAT site. The blood glucose levels of the recipient mice were monitored for more than 100 days. Islet grafts were retrieved at day 100 after transplantation for histological analysis. The protocol for islet transplantation in the ISWAT site described in this manuscript is simple and effective.

Inguinal Subcutaneous White Adipose Tissue ISWAT Transplantation Model of Murine Islets

In this protocol, a method of murine islet isolation and transplantation into the inguinal subcutaneous white adipose tissue is described. Isolated syngeneic murine islets are transplanted into a murine recipient using a basement membrane hydrogel. The blood glucose level of the recipients is monitored, and histology analysis of the islet grafts is performed.

Pancreatic islet transplantation is a well-established therapeutic treatment for type 1 diabetes. The kidney capsule is the most commonly used site for ...

Fat-Covered Islet Transplantation Using Epididymal White Adipose Tissue

Islet transplantation is a cellular replacement therapy for severe diabetes mellitus. The intraperitoneal cavity is typically the transplant site for this procedure. However, intraperitoneal islet transplantation has some limitations, including poor transplant efficacy, difficult graft detection ability, and a lack of graftectomy capability for post-transplant analysis. In this paper, "fat-covered islet transplantation", an intraperitoneal islet transplantation method that utilizes epididymal white adipose tissue, is used to assess the therapeutic effects of bioengineered islets. The simplicity of the method lies in the seeding of islets onto epididymal white adipose tissue and using the tissue to cover the islets. While this method can be categorized as an intraperitoneal islet transplantation technique, it shares characteristics with intra-adipose tissue islet transplantation. The fat-covered islet transplantation method demonstrates more robust therapeutic effects than intra-adipose tissue islet transplantation, however, including the improvement of blood glucose and plasma insulin levels and the potential for graft removal. We recommend the adoption of this method for assessing the mechanisms of islet engraftment into white adipose tissue and the therapeutic effects of bioengineered islets.

This fat-covered islet transplantation method is suitable for the detection of engrafted islets in the intraperitoneal cavity. Notably, it does not require the use of biobinding agents or suturing.

Islet transplantation is a cellular replacement therapy for severe diabetes mellitus. The intraperitoneal cavity is typically the transplant site for this ...

Minimizing Post-Infusion Portal Vein Bleeding during Intrahepatic Islet Transplantation in Mice

Although the liver is currently accepted as the primary transplantation site for human islets in clinical settings, islets are transplanted under the kidney capsule in most rodent preclinical islet transplantation studies. This model is commonly used because murine intrahepatic islet transplantation is technically challenging, and a high percentage of mice could die from surgical complications, especially bleeding from the injection site post-transplantation. In this study, two procedures that can minimize the incidence of post-infusion portal vein bleeding are demonstrated. The first method applies an absorbable hemostatic gelatin sponge to the injection site, and the second method involves penetrating the islet injection needle through the fat tissue first and then into the portal vein by using the fat tissue as a physical barrier to stop bleeding. Both methods could effectively prevent bleeding-induced mouse death. The whole liver section showing islet distribution and evidence of islet thrombosis post-transplantation, a typical feature for intrahepatic islet transplantation, were presented. These improved protocols refine the intrahepatic islet transplantation procedures and may help laboratories set up the procedure to study islet survival and function in pre-clinical settings.

Here we present refined surgical procedures on successfully performing intraportal islet transplantation, a clinically relevant but technically challenging surgical procedure, in mice.

Although the liver is currently accepted as the primary transplantation site for human islets in clinical settings, islets are transplanted under the ...

A Method for Islet Transplantation to the Omentum in Mouse

Islet transplantation has been proposed to be a potential treatment for type 1 diabetes. Recent compelling evidence indicates that intravascular islet infusion is far from ideal and therefore, the omentum is re-emerging as a potentially valuable site for islet transplantation. This experiment requires the isolation of high quality islets and the implantation of the islets to the diabetic recipients. Transplantation to the omentum requires surgical steps that can be better demonstrated visually. Here, the detailed steps for this procedure are presented. Two methods of mixing the isolated islets with hydrogel before placing the mixture into the omental pouch of diabetic mice are described here. Different hydrogels are used for the different conditions. Blood glucose levels of diabetic mouse recipients of syngeneic islets in the omentum were monitored for up to 35 days. Some animals were sacrificed after 14 days to perform immuno-histochemical analysis. This pre-clinical transplantation approach can be used as preliminary data leading up to translation to clinical transplantation.

A method for the omental transplantation of islets in a mouse is introduced. The isolated islets are mixed with hydrogel and the mixture is placed into the omental pouch of a diabetic mouse. Then, the blood glucose is monitored, and immuno-histochemical analysis is performed.

Islet transplantation has been proposed to be a potential treatment for type 1 diabetes. Recent compelling evidence indicates that intravascular islet ...

Murine Model of Islet Transplantation: A Surgical Method to Infuse Pancreatic Islets in Portal Vein and Prevention of Postoperative Bleeding

Source: Gou, W. et al. <a href="https://www.jove.com/v/62530">Minimizing Post-Infusion Portal Vein Bleeding during Intrahepatic Islet Transplantation in Mice.</a> J. Vis. Exp. (2021)
In this video, we describe a procedure to infuse pancreatic islet cells into the&#160;hepatic portal vein, followed by application of gel foam to successfully prevent bleeding. Postoperative bleeding is a common cause of high mortality during islet transplantation.

Source: Gou, W. et al. Minimizing Post-Infusion Portal Vein Bleeding during Intrahepatic Islet Transplantation in Mice. J. Vis. Exp. (2021)
In this video, ...