In this protocol, we discuss the implementation of a model of successful orthotopic liver transplantation (OLT) in mice. Additionally, adjuvants to further analyze allograft patency after successful OLT in a mouse are discussed as well, specifically utilizing microcomputed tomography (microCT) scans.
Microcomputed tomography (microCT) angiography is an invaluable resource to researchers. New advances in this technology have allowed for high-quality images to be obtained of micro-vasculature and are high-fidelity tools in the field of organ transplantation. In this model of orthotopic liver transplantation (OLT) in mice, microCT affords the opportunity to evaluate allograft anastomosis in real time and has the added benefit of not having to sacrifice study animals. The choice of contrast, as well as image acquisition settings, create a high-definition image, which gives researchers invaluable information. This allows for evaluation of the technical aspects of the procedure as well as potentially evaluating different therapeutics over an extended duration of time. In this protocol, we detail an OLT model in mice in a stepwise fashion and finally describe a microCT protocol that can give high-quality images, which aid researchers in in-depth analysis of solid organ transplantation. We provide a step-by-step guide for liver transplantation in a mouse, as well as briefly discuss a protocol for evaluating the patency of the graft through microCT angiography.
Transplantation is the only effective therapy for end-stage liver disease. Undeniably, the benefit of liver transplantation is excellent, with a median survival of 11.6 years versus 3.1 years on the waiting list1. However, there are significant constraints, which limit the broad application of liver transplantation, and include most importantly, a lack of suitable, high-quality donor organs. Expanding the donor organ pool will, thus, require innovative strategies that allow for the use of allografts currently considered unsuitable today, increasing the margin of safety for transplantation. Therefore, to improve access to liver transplantation, it is imperative to conduct preclinical studies in small animals.
Particularly important to transplantation research are in vivo models of transplantation. Mouse orthotopic liver transplantation (OLT) has been around for almost 30 years2 and is vital to studying many aspects of transplantation, including the characterization of immune responses, ischemia-reperfusion injury, acute rejection, therapeutic effects of novel agents, and long-term survival3,4,5,6,7. The use of mice to study transplantation is vital as it allows for the use of transgenic mouse lines to study the impact of specific molecular pathways on the outcomes of transplantation. Established protocols of mouse liver transplantation have been well described previously8,9.
Multiple methods of anastomoses exist for the supra and infra hepatic inferior vena cava (IVC), portal vein (PV), and common bile duct (CBD). They typically rely on either hand anastomosis or a modified vascular cuff technique similar to murine lung transplantation10,11,12. An important step in the long-term study and survival of the recipient mice, as well as the development of a sustained mouse liver transplant program, is the ability to evaluate these critical anastomoses. Imaging modalities to evaluate liver allograft patency often rely on ultrasound and computer tomography (CT) in the clinical setting13,14. CT has a distinct advantage over ultrasound as it can offer views of the entire abdomen to include every anastomosis, although obtaining these views with ultrasound can be particularly difficult in small animals. Significant research and resources have been devoted to developing accurate microCT for the purpose of enhancing animal studies and the information we can gather from these models of injury and disease15,16. Here we describe a protocol for orthotopic mouse liver transplantation (Figure 1) and briefly describe a protocol for microCT to evaluate allograft patency and durability of anastomoses.
Male C57BL/6J mice (30 g body weight) were housed under pathogen-free conditions at the Nationwide Children's Hospital Animal Facility. All procedures were humanely performed according to the NIH and the National Research Council's Guide for the Humane Care and Use of Laboratory Animals and with the approval of the Nationwide Children's Hospital Institutional Animal Care and Use Committee (IACUC Protocol AR17-00045). See the Table of Materials for details related to all materials, instruments, and equipment used in this protocol.
1. Initial setup for transplantation surgery
2. Donor mouse procurement
3. Back table preparation of liver allograft
4. Recipient operation
NOTE: As this is a sterile operation, use gloves and proper personal protective equipment and administer antibiotics. Administer 0.1 mg/kg Buprenorphine subcutaneously as preoperative analgesia at the time of surgery.
5. Mouse microCT angiography imaging
For those researchers who are not surgeons, unfamiliar with anatomy, or uncomfortable interpreting radiologic results, proper image analysis should be done by personnel with proper training. The success of an OLT in a mouse is demonstrated in the above protocol. Furthermore, to enhance study metrics and provide real-time feedback for the success of a transplant, as well as eliminate the need for necropsy, a microCT angiography scan can be used to provide accurate and clear images. Representative images are included in this manuscript (Figure 11). Representative images of in vivo failed anastomosis can be seen in Figure 12.
Those familiar with hepatic anatomy and vasculature can see patent venous anastomoses of the IVC. In some circumstances, the portal vein can be visualized as well, which is made easily in this model due to the portal vein cuff. The viewing of open anastomoses indicates the technical success of the operation. Additionally, 3D reconstruction of these images can provide additional information to researchers and a more detailed image of the vascular anatomy. Utilizing this above model, mortality in the OLT mice cohort is ~40-45%.
Figure 1: Overview of orthotopic liver transplant. (A) Graphical drawing depicting the four different anastomoses: i) suprahepatic IVC anastomosis, ii) infrahepatic IVC anastomosis, iii) portal vein anastomosis, iv) common bile duct anastomosis. Each arrow indicates a relative location for where the vessel or duct should be cut-supra-hepatic IVC (protocol step 2.13), infra-hepatic IVC (protocol step 2.11), portal vein (protocol step 2.10), and common bile duct (protocol step 2.7). (B) In vivo diagram of anastomoses. Scale bar = 2 mm. Abbreviation: IVC = inferior vena cava. Please click here to view a larger version of this figure.
Figure 2: Surgical tools used in surgery. (A) 45° fine forceps, (B–E) fine forceps, (F) curved needle holder/forceps, (G) straight forceps, (H) vascular clamp applier, (I) hemostat, (J) needle holder, (K) electro-cautery device, (L) #11 blade, (M) abdominal retractor, (N,O) micro-scissors, (P) fine scissors, (Q) surgical scissors, (R,S) Yasargil clamps, (T) bulldog vein clamp, (U) micro-vascular clamp. Please click here to view a larger version of this figure.
Figure 3: Portal vein cuff and bile duct stent. Ex vivo image of stents and cuffs prior to use. Scale bar = 3.5 mm. Please click here to view a larger version of this figure.
Figure 4: Common bile duct stenting during donor operation. (A) Bile duct stent being inserted into the common bile duct. (B) Bile duct stent secured within the bile duct. Scale bar = 2 mm. Please click here to view a larger version of this figure.
Figure 5: Placing portal vein cuff during back table preparation of the liver allograft. (A) Threading portal vein through the venous cuff. (B) Everted vein over the cuff. Scale bar = 2 mm. Please click here to view a larger version of this figure.
Figure 6: Portal vein anastomosis during recipient operation. (A) Inserting vein cuff into recipient portal vein. (B) Portal vein anastomosis secured with suture. Scale bar = 2 mm. Please click here to view a larger version of this figure.
Figure 7: Suprahepatic IVC anastomosis during recipient operation. (A) Posterior wall of the anastomosis is complete. (B) Completed SHIVC anastomosis. Scale bar = 2 mm. Abbreviations: IVC = inferior vena cava; SHIVC = suprahepatic IVC. Please click here to view a larger version of this figure.
Figure 8: Infrahepatic IVC anastomosis during recipient operation. (A) Posterior wall of the anastomosis is complete. (B) Completed IHIVC anastomosis. Scale bar = 2 mm. Abbreviations: IVC = inferior vena cava; IHIVC = infrahepatic IVC. Please click here to view a larger version of this figure.
Figure 9: Common bile duct anastomosis during recipient operation. (A) Placing bile duct stent within the recipient common bile duct. (B) Securing bile duct anastomosis. Scale bar = 1 mm. Please click here to view a larger version of this figure.
Figure 10: Mouse microCT angiography animal preparation. (A) Mouse tail vein injection to administer contrast. (B) Mouse being passed through microCT machine. Abbreviation: microCT = microcomputed tomography. Please click here to view a larger version of this figure.
Figure 11: Representative images showing microCT angiography of allograft patency. (A,B) Contrast can be seen throughout the IVC, demonstrating patency of the suprahepatic and infrahepatic anastomoses. (C) Contrast in the portal-vein, again demonstrating patency. (D) 3D reconstruction of the vasculature. Abbreviations: microCT = microcomputed tomography; IVC = inferior vena cava; PV = portal vein. Please click here to view a larger version of this figure.
Figure 12: Representative images showing failed in vivo anastomoses. (A) Failed portal vein anastomosis due to distortion of the vein resulting in lack of blood flow. (B) Failed suprahepatic IVC anastomosis due to excessive bleeding. Scale bar = 2 mm. Please click here to view a larger version of this figure.
OLT in rodents has been well described in the literature2,8. To perform this technically demanding procedure, often several years of micro-surgery (or surgery in general) is required as this involves a robust understanding of anatomy and technical ability. In developing this model, we faced several technical issues all revolving around the anastomoses. Particularly with the PV anastomosis, it is often difficult to stabilize the vein for anastomosis. We have found that placing one or two sutures (surgeon preference) helps with facilitating cuff placements. It is to be noted that placing more stay sutures increases surgical time.
Additionally, the SHIVC is deep within the abdominal cavity and is difficult to place a clamp on to give adequate exposure. We have found that if the mouse is relaxed as possible in its restraint, that will add to the flexibility of the vein. Ultimately, it will be up to the surgeon to determine the proper placement with practice. Furthermore, with the CBD anastomosis, the duct is again very delicate. It can be difficult to place stay sutures to stabilize the duct, and possibly, placing it on a small piece of gauze will aid in its stabilization. Finally, as all small mammals are uniquely delicate with respect to anesthesia time, it is important to perform the surgery as quickly as possible. Ideal surgical times are as follows: 1) donor operation, 45-60 min; 2) back table preparation, 15 min; 3) recipient operation, 60-80 min. Practice will help with decreasing wasted movement.
As animal models advance, the ability to evaluate the success of study interventions has also advanced. MicroCT was first used to conduct studies on vasculature in rats in the late 1990s17. There are many challenges to performing accurate and clear microCT angiography studies in rodents. However, most of the challenges arise from the short cardiac and respiratory cycles of these mammals. This is overcome by using short exposures to limit motion artifacts as well as higher photon fluence rates18. In general, we found that the use of cardiac gating, as well as the adjustment of isoflurane concentrations to decrease the respiratory rate, produced the clearest images. We have also found that utilizing rodent-specific contrast timing for specific phases: hepatic arterial phase, portal-venous phase, and delayed phase have also improved visualization19. The use of ExiTron nano 12000 contrast has several advantages and can improve overall image quality. It offers the strongest contrast enhancement in the liver20 and the blood21. Another advantage is that the contrast is present in the liver for up to 120 h after the initial injection, which could cut down on associated liver toxicity as less contrast is needed if repeated scans are required20.
Furthermore, because scans are performed with the mouse sedated with isoflurane, contrast enhancement is not altered with this change in physiology20. By employing these imaging techniques and ExiTron contrast, a clear evaluation of successful anastomoses in OLT is possible. MicroCT allows for noninvasive evaluation of in-vivo allografts over an extended period. This protocol decreases the number of animals that must be sacrificed to evaluate vascular anastomoses and afford the opportunity to study therapeutics over several weeks and their effect on the vasculature.
Limitations
It is to be noted that while multiple revisions of the OLT model have occurred to perfect its technique, the visualization of the anastomoses utilizing microCT is still an ongoing process. Furthermore, mouse OLT offers a unique insight into transplantation medicine. However, it is not an encompassing model as it is difficult to keep these mice alive past 1 week. Additional transplant models should be used as well to further substantiate preclinical experiments.
Conclusions
Advances in microCT have rapidly progressed over the past decade, providing researchers with invaluable new tools in the field of animal models and transplantation. In the future, more detailed 3D imaging will offer further insights into research and discovery.
The authors have nothing to disclose.
SMB is supported by the National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK) grant R01DK1234750. BAW is supported through the National Institutes of Health National Heart Lung and Blood Institute grant R01HL143000.
#11 Blade | Fisher Scientific | 3120030 | |
4-0 silk suture | Surgical Specialties Corp. | SP116 | |
6-0 nylon suture | AD Surgical | S-N618R13 | |
7-0 nylon suture | AD Surgical | S-N718SP13 | |
8-0 nylon suture | AD Surgical | XXS-N807T6 | |
10-0 nylon suture | AD Surgical | M-N510R19-B | |
20 G Angiocath | Boundtree | 602032D | |
30 G Needle | Med Needles | BD-305106 | |
Baytril (enrofloxacin) Antibacterial Tablets | Elanco | NA | |
Bovie Chang-A-Tip High Temp Cauterizer | USA Medical and Surgical Supplies | BM-DEL1 | |
Bulldog Vein Clamp 1 1/8 | Ambler Surgical USA | 18-181 | |
C57BL/6J mice | Jackson Labs | ||
Castroviejo Micro Dissecting Spring Scissors | Roboz Surgical Store | RS-5668 | |
Dumont #5 – Fine Forceps | Fine Science tools | 11254-20 | |
Dumont #5 Forceps | Fine Science tools | 11252-50 | |
Dumont Medical #5/45 Forceps – Angled 45° | Fine Science tools | 11253-25 | |
ExiTron nano 12000 | Miltenyi Biotec | 130 - 095 - 698 | CT contrast agent |
Forceps | Fine Science tools | 11027-12 | |
Halsted-Mosquito Hemostat | Roboz Surgical | RS-7112 | |
heparin | Fresnius Lab, Lake Zurich, IL | C504701 | |
histidine-trypotophan-ketoglutarate | University Pharmacy | NA | |
Insulated Container | YETI | ROADIE 24 HARD COOLER | https://www.yeti.com/coolers/hard-coolers/roadie/10022350000.html |
Isoflurane | Piramal Critical Care | NDC 66794-017-25 | |
ketamine | Hikma Pharmaceuticals PLC | NDC 0413-9505-10 | |
Mirco Serrefines | Fine Science tools | 18055-05 | |
Mouse Rectal Temperature Probe | WPI Inc | NA | |
NEEDLE HOLDER/FORCEPS straight | Micrins | MI1540 | |
PE10 Tubing | Fisher Scientific | BD 427400 | |
perfadex | XVIVO Perfusion AB | REF99450 | |
PhysioSuite | Kent Scientific | PS-MSTAT-RT | |
Puralube Ophthalmic Ointment | Dechra | NA | |
saline | PP Pharmaceuticals LLC | NDC 63323-186-10 | |
Scissors | Fine Science tools | 14090-11 | |
Small Mouse Restraint – 1” inner diameter | Pro Lab Corp | MH-100 | |
SomnoSuite Small Animal Anesthesia System | Kent scientific | SS-MVG-Module | |
Surgical microscope | Leica | M500-N w/ OHS | |
U-CTHR | MI Labs | NA | CT Scanner software |
Vannas-Tubingen Spring Scissors | Fine Science Tools | 15008-08 | |
xylazine | Korn Pharmaceuticals Corp | NDC 59399-110-20 | |
Yasagil clamp | Aesculap | FT351T | |
Yasagil clamp | Aesculap | FT261T | |
Yasagil clamp applicator | Aesculap | FT484T |