This protocol details the surgical steps of a mouse model of vascularized heterotopic spleen transplantation, a technically challenging model that can serve as a powerful tool in studying the fate and longevity of spleen cells, the mechanisms of distinct spleen cell populations in disease progression, and transplant immunity.
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Wang, J. J., Qiu, L., Fernandez, R., Yeap, X. Y., Lin, C. X., Zhang, Z. J. A Mouse Model of Vascularized Heterotopic Spleen Transplantation for Studying Spleen Cell Biology and Transplant Immunity. J. Vis. Exp. (148), e59616, doi:10.3791/59616 (2019).
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The spleen is a unique lymphoid organ that plays a critical role in the homeostasis of the immune and hematopoietic systems. Patients that have undergone splenectomy regardless of precipitating causes are prone to develop an overwhelming post-splenectomy infection and experience increased risks of deep venous thrombosis and malignancies. Recently, epidemiological studies indicated that splenectomy might be associated with the occurrence of cardiovascular diseases, suggesting that physiological functions of the spleen have not yet been fully recognized. Here, we introduce a mouse model of vascularized heterotopic spleen transplantation, which not only can be utilized to study the function and behavioral activity of splenic immune cell subsets in different biologic processes, but also can be a powerful tool to test the therapeutic potential of spleen transplantation in certain diseases. The main surgical steps of this model include donor spleen harvest, the removal of recipient native spleen, and spleen graft revascularization. Using congenic mouse strains (e.g., mice with CD45.1/CD45.2 backgrounds), we observed that after syngeneic transplantation, both donor-derived splenic lymphocytes and myeloid cells migrated out of the graft as early as post-operative day 1, concomitant with the influx of multiple types of recipient cells, thus generating a unique chimera. Despite relatively challenging techniques, this procedure can be performed with >90% success rate. This model allows tracking the fate, longevity, and function of splenocytes during steady state and in a disease setting following a spleen transplantation, thereby offering a great opportunity to discover the distinct role for spleen-derived immune cells in different disease processes.
The spleen is the largest secondary lymphoid organ in the body and is critical in the immune and hematopoietic systems. Its functions are primarily carried out by two morphologically distinct compartments, the red pulp and the white pulp1. The red pulp is a three-dimensional meshwork of venous sinuses and splenic cords that consist of reticular fibers, reticular cells, and associated macrophages. This unique structure allows the red pulp to act as an effective blood filter that removes foreign materials and old or damaged erythrocytes. The white pulp includes follicles, marginal zone, and the periarteriolar lymphoid sheaths (PALS) and is an important site for antigen trapping and processing, lymphocyte homing, transformation, proliferation, and maturation2. Nevertheless, the spleen has commonly been considered as a dispensable organ because other lymphatic organs, such as lymph nodes, can also carry out some of its functions and the loss of spleen does not usually lead to death. Splenectomy has therefore been widely performed as a therapeutic method for patients with splenic injury or benign hematologic diseases3. However, patients with splenectomy face a number of long-term complications. Bacterial infections are the best-recognized complications of splenectomy4,5. Recently, the overwhelming post-splenectomy sepsis has been recognized as an intensive complication of splenectomy associated with a high mortality6. Moreover, recent epidemiological studies indicate that splenectomy may be associated with the occurrence of cardiovascular diseases, suggesting that further physiological functions of the spleen remain to be explored7,8.
Both spleen autotransplantation and spleen allotransplantation have been utilized in the clinic. Currently, spleen autotransplantation by implanting sections of splenic tissue into pouches created in the greater omentum is considered as the only possibility for preserving splenic function after traumatic splenectomy9,10. However, the efficacy of this surgery is debatable as post-surgery complications like aseptic necrosis of the splenic tissue and small bowel obstruction due to postoperative adhesions could occur11. Spleen allotransplantation is involved in multivisceral transplantation12. Clinical evidence from multivisceral transplantation suggests that spleen allotransplantation may play a protective role in small bowel allograft rejection without causing graft-versus-host disease (GVHD)12. Yet literature regarding the beneficial effect of spleen allotransplantation as a component of multivisceral transplantation is still limited and the underlying mechanisms remain to be defined. In 2006, Yair Reisner et al. reported that transplanting pig embryonic spleen tissue that has no T cells to mice could cure hemophilia A, a genetic disease without causing GVHD13, supporting that spleen transplantation holds therapeutic promise in certain diseases. Therefore, there is a need for further investigations on the therapeutic potential of spleen transplantation.
Animal models of spleen transplantation are valuable to explore the unappreciated function of the spleen-derived immune cells in disease progression as well as to test the potential therapeutic effect of spleen transplantation. Experimental whole spleen transplant models have been documented since early 1900s, as reviewed by Cohen14. In 1969, Coburn Richard J. and Lee et al. detailed the technique of spleen transplantation in rats15,16. More recently, Swirski FK et al. described a mouse model of spleen transplantation17. Compared to rat models, mouse models of spleen transplantation are more attractive due to its several inherent advantages. For example, by utilizing a mouse model, we can access an expansive variety of reagents unavailable to that of rat models. Moreover, by using congenic mice (e.g., mice with CD45.1/CD45.2 background), a syngeneic spleen transplantation makes it possible to track the fate, longevity, and function of splenocytes18. Based on the work by Swirski FK et al.17, we further established this simplified and enhanced protocol of spleen transplantation in mice. The protocol described below combines both reliability and feasibility in a standardized manner and can be utilized as a tool to study spleen biology and transplant immunity.
All procedures and animal use in this study were performed according to protocols approved by the Northwestern University Internal Animal Care and Use Committee (IACUC). In this study, 8 to 10 week old male CD45.2 and CD45.1 mice (both on BALB/c background, from Jackson laboratory) were used as spleen donors and recipients, respectively, to create syngeneic spleen transplantation models. All animals were housed in the sterile environment in the animal facilities of Northwestern University. The eye lubricant was applied to all mice post-anesthetization to prevent dryness.
1. Surgical Preparation, Anesthetization, and Analgesia Regimen
- Place a sterile disposable drape (45.7 cm x 66 cm) on the surgical platform. Gently grab the mouse, inject ketamine (50 mg/kg) and xylazine (10 mg/kg) intraperitoneally (i.p) for anesthesia, and inject 0.05 mg/kg buprenorphine subcutaneously for analgesia.
- Ensure the depth of anesthesia by toe-pinch, shave the hair in the whole abdomen area with a razor and place the mouse on the sterile surgical platform under the operating microscope at 6-10x magnification.
2. Donor Spleen Harvest
- Sterilize the abdomen with an alcohol prep pad, secure the limbs with surgical tape, and make a 3-4 cm midline vertical skin incision from the pubis to the xiphoid process with scissors.
- Retract the abdominal wall with sterile retractors made from paperclips. Move the intestines to the right flank of the abdomen (surgeon’s left) side with a sterile cotton swab to expose the spleen. Cauterize the short gastric vein attached to the spleen with a sterile low temperature cautery (Figure 1A). Place a piece of sterile gauze soaked with 37 °C saline over the spleen to keep it moist (Figure 1B).
- Separate and mobilize the portal vein from the pancreatic tissue (Figure 1C) by ligating the portal vein branches (superior pancreaticoduodenal vein and right gastric vein); place a suture around the portal vein distal from the splenic vein (Figure 1D).
- Flip the spleen to the right side to expose the aorta and celiac trunk with the splenic artery (Figure 1E). Dissect and mobilize aortic-celiac-splenic artery by ligating the hepatic artery and gastric artery; place a suture around the aorta proximal to the celiac artery (Figure 1F-G).
- Inject 100 international units (IU) heparin into the inferior vena cava (IVC) to heparinize the whole body and wait 3 min to ensure the heparin take effects. Ligate the aorta proximal to celiac artery, transect the portal vein, and then perfuse the whole body using 10 mL of heparinized cold (4 °C) saline (10 mL/20 s) from the abdominal aorta distal to celiac trunk (Figure 1H).
- Collect the spleen graft en bloc with the associated aortic-celiac-splenic segment and the portal vein along with a segment of splenic vein and a small portion of pancreatic tissue. Preserve the graft in 5 mL of 4 °C saline before transplant. Euthanize the mouse by cervical dislocation.
3. Recipient Splenectomy and Spleen Graft Implantation
- Place a heating pad on the surgical platform and adjust the temperature to 37 °C. Place a sterile drape (45.7 cm x 66 cm) on top of the heating pad to create a sterile surgical platform. Repeat steps 2.1 and 2.2 for surgical preparation and anesthetization. Make a 3-4 cm midline incision and retract the abdominal wall as described in step 2.1 and step 2.2.
- Carefully move the intestine to right side of the mouse with a sterile cotton swab to expose the recipient’s spleen. Ligate the splenic vein and artery and remove the spleen.
- Carefully move the intestine to left side of the mouse and cover the intestines with wet gauze (soaked with sterile 37 °C saline). Dissect and ligate the lumbar branches of the infrarenal aorta and IVC; cross-clamp the infrarenal aorta and IVC by using two 4 mm microvascular clamps.
- Place an 11-0 nylon suture through the infrarenal aorta (a full thickness) and retract to create an elliptical aortotomy by a single cut with microscissors (the length should match the diameter of the donor aorta, Figure 2A). Pierce the IVC using a 30 G needle to create an elliptical venotomy and extend the opening to donor portal vein-matched length using microscissors (Figure 2A).
- Clear the intraluminal blood or blood clot (in the aorta and IVC) with 500 μL of heparinized saline (10 units/mL).
- Place the spleen graft in the right flank of the recipient mouse abdomen; carefully identify the donor’s aortic cuff and the donor’s portal vein. After making sure that the vessels are not twisted, cover the spleen graft with gauze soaked with cold (4 °C) saline.
- Connect the donor’s aortic cuff to the proximal and distal apex of the recipient’s aortaotomy with two stay sutures (11-0 nylon suture, same as below) (Figure 2B,C). Make an anastomosis with 2-3 bites of continuous 11-0 nylon sutures between the donor’s aortic cuff and the recipient’s aortaotomy (anterior wall) (Figure 2D). Turn the spleen graft over to the left side of the recipient; make the anastomosis between the donor’s aortic cuff and the recipient’s aortotomy (posterior wall) (Figure 2E).
- Perform an anastomosis to connect the donor’s portal vein to the posterior wall of the recipient’s IVC, using 4 to 5 bites of continuous sutures on the inside of the IVC and then close the suture on the outside of the IVC (Figure 2F,G).
- Release the vessel clamps and use sterile cotton swab to tamponade bleeding until the spleen color is recovered (Figure 2H).
- Close the abdomen with a 5-0 synthetic absorbable vicryl suture in a continuous pattern. Close the skin layer with a 5-0 nylon suture in an interrupted pattern.
NOTE: For the steps 4.7-4.8, alternatively, make an anastomosis between the donor’s portal vein and the recipient’s IVC first (step 4.8); then make an anastomosis between the donor’s aortic cuff and the posterior wall of the recipient’s aortotomy, using 2 to 3 continuous sutures in the inside of the aorta and close the suture on the outside of the aorta.
4. Animal Recovery
- Inject 1 mL of warm saline subcutaneously via 4 separate locations (0.25 mL/location) after closing the abdomen.
- Keep the mouse in a temperature-controlled incubator (30 °C) for the first few hours post-operation, monitor the mouse until it has regained sufficient consciousness, and then transfer the mouse to a new clean cage with regular food and water, with a heating pad (30 °C) underneath the cage. Keep the mouse post-surgery in a separate cage.
5. Post-surgical Pain Management
- Inject 0.05 mg/kg buprenorphine subcutaneously 24 h and 48 h post-surgery to maintain analgesia regimen.
The entire procedure of mouse spleen transplant can be completed within 90 min by experienced microsurgeons. Our laboratory has performed over 100 spleen transplants in mice. The success rate is over 90%, as defined by the survival of both recipient mouse and the spleen graft to post-operative day (POD) 1 or POD 7 (our study endpoint). The survival of the spleen graft was confirmed by the macroscopic appearance and flow cytometry analysis of the splenocytes. Based on our experience, the flow cytometry analysis (LIVE/DEAD Cell Viability Assays) is very sensitive to determine whether a spleen graft is survived, as the majority of the spleen cells would be dead if the spleen grafts were necrotic. The technical challenges of this procedure, common complications, and their troubleshooting are summarized in Table 1.
To test the graft morphology post-transplantation, Haemotoxylin and Eosin (H&E) staining was performed in spleen isografts of BALB/c mice at POD 1 and POD 7. The representative pictures are shown in Figure 3. The architecture of the spleen isografts remained intact during the first postoperative week. The red pulp, white pulp, and the marginal zone were still clear and distinguishable. To investigate the cell migration after spleen transplantation, flow cytometry was performed at POD 1 and POD 7 to examine the phenotypes of leukocytes in the spleen isografts, lymph nodes, blood, and bone marrow. As shown in Figure 4A, at POD 1, 51 ± 7% (mean ± SD, same as below) of the spleen cells were donor-derived and 46 ± 3% were recipient-derived. At POD 7, donor-derived leukocytes accounted for 32 ± 10 % of total spleen cells, and recipient-derived cells were up to 56 ± 13%. We also observed that spleen leukocytes migrated into the lymph nodes, blood, and bone marrow as early at day 1 and maintained at day 7 (Figure 4B), generating a unique chimera valuable for splenocyte trafficking research.
Table 1: Troubleshooting methods.
Figure 1: Donor spleen harvest. (A) Cauterize the short gastric vein attached to the spleen. (B) Place a small piece of sterile warm wet gauze over the spleen to keep it moist. (C) Dissect and isolate the portal vein behind the pancreas. (D) Ligate the side branches of the portal vein and place a suture round the portal vein distal to the splenic vein. The dashed lines represent the location transecting later for the anastomosis with the recipient IVC. (E) Flip the spleen over to the right side of the abdomen (surgeon’s left) to expose the aorta and celiac trunk with its branches including splenic artery. (F) Dissect and mobilize aortic -celiac -splenic artery by ligating the two other branches. (G) Place a suture around the aorta proximal to the celiac artery. The dashed lines represent the location transecting later used for the anastomosis with recipient abdominal aorta. (H) After ligating the aorta and transecting the portal vein, perfuse the spleen graft with 10 mL of heparinized saline through the aorta. Please click here to view a larger version of this figure.
Figure 2: Spleen graft implantation. (A) After isolation and cross clamping the aorta and IVC, make a longitudinal aortotomy and venotomy in the aorta and IVC, respectively. (B-D) Place the spleen graft on the right side of the abdomen (surgeon’s left side). Make the end-to-side anastomosis between donor aortic cuff and the anterior wall of the recipient’s aortotomy, using 2-3 bites of continuing suture. (E) Turn the spleen graft over to the left flank of the recipient; repeat the previous procedure between the donor aortic cuff and the posterior wall of the recipient’s aortotomy and close the suture on the outside of aorta. (F-G) Make an end-to-side anastomosis between the donor portal vein and the posterior wall of the recipient’s IVC, using 4 to 5 bites of continuous 11-0 nylon suture in the inside of the IVC and then close the suture on the outside of the IVC. (H) After completing the anastomosis, release the vessel clamps and place some cotton buds to help stop the bleeding. Please click here to view a larger version of this figure.
Figure 3: Representative histology of spleen isografts on day 1 and day 7 post-operation. Syngeneic spleen transplantations were performed using BALB/c mice. Spleen isografts were harvested on day 1 and day 7 post-transplantation and fixed in 10% formalin for 48 h. H&E staining was performed using paraffin-embedded tissue section. The representative histology shows that the spleen isografts remain intact at 1 week post-transplantation. Scale bar = 250 μm. POD, post-operative day. Please click here to view a larger version of this figure.
Figure 4: The chimera created by syngeneic spleen transplantation. Three syngeneic spleen transplantations were performed using BALB/c CD45.2 mice as donors and BALB/c CD45.1 mice as recipients. The spleen grafts, blood, lymph node (LN), and bone marrow (BM) in recipient mice were harvested on day 1 and day 7 post-transplantation. Single cell isolation and flow cytometry analysis were performed to analyze the phenotype of the cells. (A) Representative dot plots (gating from live singlets) showing the percentages of donor or recipient — derived leukocytes in the indicated samples. (B) Percentages (mean ± SEM) of the indicated populations in donor- derived cells in the indicated samples at day 1 and day 7. POD = post-operative day. Please click here to view a larger version of this figure.
Supplementary Figure 1: Representative plots (n = 3) showing the percentages of donor-derived versus recipient-derived lymphocytes in the transplanted spleen, lymph node, blood, and bone marrow. Please click here to download this file.
Supplementary Figure 2: Representative plots (n = 3) showing the percentages of donor-derived versus recipient-derived CD11b+Ly6C+monocytes in the transplanted spleen, lymph node, blood, and bone marrow. Please click here to download this file.
Compelling evidence suggests that spleen-derived monocytes play an important role in sterile inflammatory processes such as atherosclerosis19, acute ischemic brain20 or lung injury18, as well as myocardial I/R injury and remodeling21,22,23. These reports highlight the under-recognition role of the spleen in many chronic diseases, of which cardiovascular disease is an important one (especially given it is the number one killer globally). The mouse model of spleen transplantation offers a great opportunity to discover the role of spleen-derived immune cells in various diseases as well as how they are primed in the spleen. For example, by using the mouse models of spleen transplantation, Swirski et al. found that in response to ischemic myocardial injury, spleen-derived monocytes increase their motility, migrate out of the spleen, adhere to injured tissue, and contribute to the wound healing17. Furthermore, this model is useful to address the longevities of mature immune cells in the spleen and underlying mechanisms and to explore therapeutic potentials of spleen transplantation.
Several aspects should be taken into consideration to improve the success of this protocol. First, it is critical to choose the proper experimental animals during the design process since the mouse weight, strain, and health condition could affect the difficulty of the surgical steps and experimental results. Our laboratory recommends using 8 to 12-week old mice with over 25 g weight to decrease the mortality that might be caused by bleeding. Second, during the spleen harvest procedure, too much manipulation of the spleen vessels could easily lead to breeding or vasospasm and potentially result in micro thrombosis in the spleen graft. Getting familiar with the anatomy of the mouse abdomen before surgery would be helpful to accelerate the learning process. Third, during the recipient surgery, the spleen graft should be always maintained moist and cool. Appropriate protection of spleen grafts could reduce the transplant ischemia/reperfusion (I/R) injury and prevent graft failure after transplantation. In addition, considering that the donor vessels for anastomosis are relatively long, positioning the spleen grafts properly before the anastomosis is critical to prevent twisting of the vessels. Moreover, the diameters of the recipient aortotomy and venotomy of IVC should be always comparable to those of the donor aorta and portal vein to ensure the proper blood flow and prevent thrombosis.
The average storage time of spleen grafts in 4 °C saline is around 10 min. The total ischemic time should be limited to less than 50 min to ensure minimum transplant failure. We recommend using the University of Wisconsin (UW) cold storage solution to preserve the spleen graft, if over 1 h cold, ischemic time is needed in studies. It should be noted that a small portion of the pancreas intimately attach with the spleen, and attempt to remove the whole from the spleen grafts would easily lead to graft or vascular complications and markedly increase the operation time. We observed that the pancreas tissue attached to the spleen graft would undergo atrophy at POD 7 though some remained viable with the spleen grafts. Whether or not to remove the pancreas tissue attached with spleen grafts depends on the research purposes.
Availability of congenic mice has made it possible to track the origin of the spleen cells, donor versus recipient after transplantation. In this study, we used BALB/c CD45.2 and BALB/c CD45.1 congenic mice as spleen donors and recipients to create a syngeneic spleen transplantation model. Organ or tissue transplantation between these congenic mouse strains has been widely used to track the origin and the development of immune system. Despite the recent report regarding a point mutation associated with CD45.1 that influences NK cell response24, no transplant rejection was reported between these strain combinations. We observed a substantial influx of recipient cells into spleen grafts that occurred as soon as at POD 1. It is likely that recipient cells responded to the transplant I/R injury as the majority of the recipient cells were granulocytes. Our results showed that a relatively high percentage of splenic lymphocytes remained of donor origin. More interestingly, these lymphocytes migrated to (repopulated) in other lymphoid compartments (lymph node, bone marrow, and circulation). These findings prompt us to speculate that lymphocytes that originated from spleens are very important in the adaptive immunity. However, more investigations are required to delineate the distinct roles of splenic lymphocytes in adaptive versus innate immunity (Figure 4, Supplementary Figure 1 and Supplementary Figure 2).
The major limitation of this protocol is that it requires extensive microsurgical training for individuals with limited microsurgical experience to master this technique. Based on our learning experience in overall mouse solid organ transplant models (e.g., mouse heart, lung, or kidney transplant), it may take 6-10 months for a newly trained person (without any experimental microsurgical technique) to skillfully master this technique. Compared to mouse models of heart, or kidney transplantation, this model could be more challenging since it involves additional tissue dissection steps to isolate the spleen graft during the donor procedures. Moreover, the diameter of the donor portal vein (around 0.6 mm) is smaller than the IVC, which makes it more difficult for the anastomosis. Another limitation is that it is not clear if the grafted spleen would be massively invaded by recipient’ cells at later transplant phase (e.g., POD 60). However, this model is also very attractive for its several inherent advantages. Firstly, mice and humans share a substantial range of similar genomes, thereby allowing for a relatively accurate representation of a realistic application. Moreover, comparing to the mouse model of non-vascularized spleen autotransplantation using the sections of splenic tissue, this vascularized spleen transplant model is less likely to develop complications such as aseptic necrosis of the splenic tissue and small bowel obstruction due to postoperative adhesions.
The mouse model of spleen transplantation has been previously reported by Swirski FK et al. However, the detailed information was not described. Our study provides a comprehensive step-by-step protocol of mouse spleen transplantation for interested researchers to follow and to mater this technique. Moreover, this protocol eliminates several unnecessary steps described in the report by Swirski FK et al. (e.g., the bile duct ligation) and introduces the 11-0 suture for anastomosis, which would help shorten the surgical time and prevent the bleeding.
In conclusion, this model could be a powerful tool to explore the mechanisms of the splenic cell population in responses to pathogens, injury, inflammation, or transplant rejection and is valuable for the test of the therapeutic potential of the spleen transplantation. With proper training and practice, this procedure can be performed with >90% success.
The authors have nothing to disclose.
Authors thank Northwestern University Comprehensive Transplant Center and the Feinberg School of Medicine Research Cores program for resource and funding support. Specifically, flow Cytometry and histology services were provided by the Northwestern University Flow Cytometry Core Facility and Mouse Histology and Phenotyping Laboratory, respectively, both of which are supported by NCI P30-CA060553 awarded to the Robert H Lurie Comprehensive Cancer Center. We thank Mr. Nate Esparza for proofreading this manuscript.
|Heparin solution||Abraxis Pharmaceutical Products||504031|
|Injection grade normal saline||Hospira Inc.||NDC 0409-4888-20|
|70% Ethanol||Pharmco Products Inc.||111000140|
|ThermoCare Small Animal ICU System||Thermocare, Inc.|
|Adson Forceps||Roboz Surgical Instruments||RS-5230|
|Derf Needle Holder||Roboz Surgical Instruments||RS-7822|
|Extra Fine Micro Dissecting Scissors||Roboz Surgical Instruments||RS-5881|
|Micro-clip||Roboz Surgical Instruments||RS-5420|
|7-0 silk||Braintree Scientific||SUT-S 103|
|11-0 nylon on 4 mm (3/8) needle||Sharpoint DR4||AK-2119|
|Ms CD45.2 antibody||BD Bioscience||553772|
|Ms CD45.1 antibody||BD Bioscience||553776|
|Ms CD11b antibody||BD Bioscience||557657|
|Ms B220 antibody||BD Bioscience||553089|
|Ms Ly6C antibody||eBioscience||48-5932-80|
|Ms Ly6G antibody||BD Bioscience||561236|
|Ms F4/80 antibody||BD Bioscience||565614|
|Ms CD11c antibody||BD Bioscience||558079|
|Ms CD3 antibody||eBioscience||48-0032-82|
|Ms CD4 antibody||BD Bioscience||552051|
|Ms CD8 antibody||BD Bioscience||563786|
|LIVE/DEAD™ Fixable Violet Dead Cell Stain Kit||Thermo Fisher||L34955|
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