Establishment of Rat Model of Acute Antibody-Mediated Rejection in Kidney Transplantation

Kidney transplantation (KT) has become a critical therapeutic approach for end-stage renal disease^1,2. However, transplant rejection remains a major factor affecting the long-term survival of renal allografts^3,4. The primary types of rejection observed clinically are acute rejection (AR) and chronic rejection (CR). Acute rejection is further classified into acute T cell-mediated rejection (TCMR) and acute antibody-mediated rejection (AAMR)^5,6,7. In recent years, with the development and application of immunosuppressive agents, the incidence of TCMR has been effectively controlled. However, AAMR remains one of the leading causes of allograft dysfunction^8,9.

The pathophysiology of AAMR is widely believed to be triggered by donor-specific antibodies (DSA) produced either before or after transplantation¹⁰. When DSA binds to antigens on the vascular endothelial cells of the allografts, the classical complement cascade is activated, leading to the formation of membrane attack complexes and subsequent allograft damage¹¹. During complement activation, the cleavage fragment C4d covalently binds to allograft capillary endothelial cell surfaces. Additionally, chemoattractants such as complement cleavage products C3a and C5a recruit inflammatory cells (e.g., macrophages) to infiltrate the allograft capillaries, resulting in capillaritis and further graft injury^12,13. Therefore, the clinical diagnosis of AAMR primarily relies on evidence of elevated serum DSA levels, capillaritis, tubular necrosis, and C4d deposition in graft capillaries^14,15,16. Current therapeutic strategies for AAMR primarily involve suppressing B-cell or plasma-cell activity, removing circulating DSA, and inhibiting complement activation. These include plasmapheresis, immunoadsorption, CD20 monoclonal antibody (rituximab), proteasome inhibitors (bortezomib), complement inhibitors (eculizumab), and an IgG-degrading enzyme (IdeS)^{17,18,19,20,21,22,23,24,25}. However, the overall treatment efficacy remains suboptimal, as no standardized therapy has been proven robustly effective in randomized trials. Furthermore, current immunosuppressive strategies are often associated with a significant burden of adverse effects, particularly increased susceptibility to severe infectious complications and malignancy^21,25,26,27. Thus, further research is needed to elucidate the pathophysiology of AAMR and develop more effective therapeutic approaches.

Investigating disease mechanisms and exploring novel prevention strategies rely on small-animal models, making the establishment of a reliable AAMR model in KT essential. Our research group has extensive experience in constructing rat KT AAMR models²⁸. This study performed KT using the Brown Norway (BN) rat to the Lewis rat. BN (RT1ⁿ) and Lewis (RT1^l) rats represent a fully MHC-mismatched pair, providing a strong genetic basis for allogeneic rejection²⁹. Compared to alternative modeling approaches, the skin-graft presensitization strategy utilized here offers distinct advantages in robustness and physiological relevance. Unlike passive antibody transfer models that rely on transient injection and lack host immune engagement, this active sensitization approach establishes long-term immunological memory, closely mimicking the clinical scenario of highly sensitized patients³⁰. Furthermore, unlike other sensitization designs, such as donor-specific blood transfusion, which may yield variable antibody titers or even induce tolerance depending on the protocol, skin grafting provides a singular, potent, and highly immunogenic stimulus²⁹. This guarantees the reproducible generation of high-titer DSA and the consistent manifestation of AAMR phenotypes (e.g., capillaritis, C4d deposition) within a defined 5-to-7-day window, making it a highly feasible platform for evaluating therapeutic efficacy. We established the AAMR model by performing KT in these allogeneic rats following two weeks of skin graft pre-sensitization and evaluated the model through serological, histopathological, and immunological analyses.

Regarding the model's applicability, users of this protocol can expect a reproducible onset of AAMR phenotypes between days 5 and 7 post-transplantation. The primary readouts essential for validating this model include the kinetics of serum DSA (IgG and IgM) production, histological evidence of microvascular inflammation (glomerulitis and peritubular capillaritis), and diffuse C4d deposition in peritubular capillaries. However, readers should be aware of a fundamental limitation: the skin presensitization method elicits a broad alloimmune response involving both humoral and cellular arms. Consequently, this model typically manifests as a mixed rejection pathology -- characterized by dominant AAMR features with concomitant TCMR, rather than an isolated antibody-mediated process.

All animal procedures were conducted in strict compliance with institutional guidelines and approved by the Institutional Animal Ethics Committee of Shandong First Medical University & Shandong Provincial Qianfoshan Hospital. All procedures adhered to the 3R principle and the Declaration of Helsinki (Approval No. JENNIO-IACUC-2024-A066). The reagents and equipment used in this study are detailed in the Table of Materials.

1. Animal preparation

1. House adult male BN (RT1ⁿ) and Lewis (RT1^l) rats (200-250 g) in specific pathogen-free (SPF) facilities. As inbred strains, individuals within the same strain are genetically identical, ensuring donor uniformity.
2. Divide the rats into four experimental groups as described in Figure 1: (1) Syngeneic KT, (2) Syngeneic KT after skin transplantation (ST), (3) Allogeneic KT, and (4) Allogeneic KT after ST.
   NOTE: Perform all surgical procedures in an SPF operating room under a stereomicroscope. Maintain animal body temperature at 37 °C using a thermostatic surgical table throughout all procedures.
3. Induce anesthesia with 4%-5% isoflurane carried by 100% oxygen at a flow rate of 1.0 L/min in an induction chamber (following institutionally approved protocols). Following induction, maintain the anesthesia with 1.5%-2% isoflurane via a nose cone. The depth of anesthesia was monitored by verifying the loss of the pedal withdrawal reflex throughout the procedure.

2. Pre-sensitization via ST (for Groups 2 and 4)

1. Harvest a 2 cm x 2 cm full-thickness skin graft from the tail of a donor rat (Lewis for Group 2; BN for Group 4).
   NOTE: The donor rat used for skin grafting is a different individual from the subsequent kidney donor, although they are from the same inbred strain.
2. Prepare a graft bed of the same size on the back of the recipient rat.
3. Place the donor skin graft onto the prepared bed and secure it using interrupted sutures as previously described³¹.
4. Allow 14 days for sensitization before proceeding with kidney transplantation.

3. Donor operation for KT

1. Anesthetize the donor rat (Lewis for Groups 1 & 2; BN for Groups 3 and 4).
2. Place the rat in a supine position and perform a midline laparotomy extending from the xiphoid process to the pubic symphysis (approximately 3-4 cm in length) to fully expose the abdominal organs³².
3. Gently retract the intestines to visualize the left kidney and its associated vasculature.
4. Isolate the left renal artery at its origin from the abdominal aorta and the left renal vein at its junction with the inferior vena cava.
   NOTE: We elect to isolate and transect the vessels directly at their ostia without harvesting an aortic or vena caval patch (Carrel patch). While this technique demands precise microsurgical skills for anastomosis, we prefer it to minimize the extent of arteriotomy on the recipient aorta and to reduce the risk of turbulence-induced thrombosis associated with patch reconstruction.
5. Perfuse the left kidney with approximately 2 mL of cold (4 °C) heparinized saline (125 U/mL) via the abdominal aorta until the kidney becomes pale³¹.
6. Transect the left renal vein, followed by the left renal artery.
7. Transect the ureter, ensuring a small patch of the bladder remains attached.
8. Harvest the kidney and immediately place it in 0.5-1.0 mL of cold hypertonic citrate adenine (HCA) kidney preservation solution on ice.

4. Recipient operation for KT

1. Anesthetize the recipient Lewis rat.
2. Perform a midline laparotomy and expose the abdominal aorta and inferior vena cava below the level of the native renal vessels.
3. Isolate a segment of the abdominal aorta and inferior vena cava.
4. Clamp the isolated vascular segment using vascular bulldog clamps.
5. Make a small longitudinal incision (approximately 1.0-1.5 mm in length, matching the diameter of the donor renal artery) in the abdominal aorta and perform an end-to-side anastomosis with the donor renal artery using 10-0 nylon sutures in a continuous fashion.
6. Make a corresponding incision in the inferior vena cava and perform an end-to-side anastomosis with the donor renal vein using 8-0 nylon sutures.
   NOTE: Systemic heparinization is not performed on the recipient to prevent excessive bleeding.
7. First, remove the vascular clamp on the inferior vena cava to establish venous outflow. Then, slowly remove the arterial clamp on the abdominal aorta to restore blood flow to the graft.
8. Check for hemostasis and assess graft perfusion. Compress any minor bleeding points with cotton swabs.
   NOTE: A successfully perfused kidney should turn pink and firm promptly.
9. Perform urinary tract reconstruction by anastomosing the donor bladder patch to the dome of the recipient's bladder using 10-0 nylon sutures.
10. Excise both native kidneys of the recipient rat to ensure that survival depends solely on the function of the graft.
    NOTE: Throughout the procedure, the total cold ischemia time (from donor harvest to reperfusion) was maintained below 60 min, and the warm ischemia time (during anastomosis) was kept under 30 min to minimize ischemic injury.
11. Close the abdominal incision in layers using 5-0 silk sutures.
12. Immediately after wound closure, administer Buprenorphine (0.05 mg/kg) subcutaneously for analgesia and Penicillin (40,000 U/kg) intramuscularly to prevent infection. Administer 1-2 mL of warm sterile saline subcutaneously to prevent dehydration.

5. Post-operative care and monitoring

1. Monitor recipient rats daily for survival, behavior (alertness, mobility), and physiological status (food intake, excretion).
2. Inspect the surgical incision daily for signs of infection or dehiscence.
3. Monitor rats hourly for the first 24 h post-surgery, at least three times daily for days 2-3, and once or twice daily thereafter.
4. Exclude recipients that die within 24 h post-transplantation due to technical complications, such as vascular thrombosis, urine leakage, or hemorrhage, from the final analysis. In this study, the overall surgical success rate was approximately 90%.

6. Post-transplant assessments

1. Detection of DSA levels
   1. Collect blood samples from the tail vein on days 0, 3, 7, and 14 post-ST and on days 1, 3, and 5 post-KT.
   2. Centrifuge the samples at ~1500 x g for 10 min to separate serum and store at -20 °C until analysis.
   3. Harvest the spleen from a donor rat and mechanically dissociate it through a 70-µm cell strainer into cold Phosphate Buffered Saline (PBS). Lyse red blood cells using ACK lysing buffer for 5 min, then wash the cells twice with PBS (centrifuge at 300 × g for 5 min). Count the cells and adjust the final concentration to 1 × 10⁷ cells/mL.
   4. Dilute the recipient serum 1:25 with PBS.
   5. Incubate 50 µL of the diluted serum with 5 × 10⁵ donor rat splenocytes for 30 min at 37 °C.
   6. Wash the cells twice with 2 mL of PBS by centrifuging at 400 × g for 5 min at 4 °C
   7. Incubate the cells with FITC-conjugated anti-rat IgG and PE-conjugated anti-rat IgM antibodies (refer to the Table of Materials for specific clones) for 1 h at 4 °C.
   8. Wash the cells and resuspend in PBS to a concentration of 5 × 10⁶ cells/mL.
   9. Analyze the samples using a flow cytometer to quantify the mean fluorescence intensity (MFI) of DSA-IgG and IgM.
2. Pathological examination of renal grafts
   1. Harvest renal grafts at specified time points (harvest on day 5 post-KT for syngeneic KT after ST group; 6 h, 12 h, 1 d, 2 d, 3 d, 4 d, and 5 d post-KT for allogeneic KT after ST group).
   2. Fix the tissue samples in 4% paraformaldehyde for 24 h at room temperature. After dehydration and clearing, embed the tissues in paraffin blocks and cut into 4 µm sections.
   3. Perform hematoxylin-eosin (HE) and periodic acid-schiff (PAS) staining on sections.
   4. For immunohistochemistry, perform antigen retrieval by immersing the sections in EDTA buffer (pH 9.0) and heating in a pressure cooker. Once full pressure is reached, maintain for 5 min, followed by natural cooling to room temperature.
   5. Treat sections with 3% hydrogen peroxide for 10 min.
   6. Incubate sections overnight at 4 °C with primary antibodies against CD31 and C4d.
   7. Apply DAB+ substrate-chromogen for 30 s at room temperature for visualization.
   8. Scan the stained sections using a digital slide scanner.

7. Statistical analysis

1. Quantitative data are presented as mean ± standard deviation (SD). Statistical analyses were performed using appropriate statistical software.
2. Evaluate recipient survival rates using Kaplan-Meier analysis.
3. Compare quantitative data between groups using Student's t-test.
   NOTE: A P-value < 0.05 is considered statistically significant.

Renal allograft survival time
Recipients in both the syngeneic KT and the syngeneic KT after ST groups survived long-term without rejection throughout the one-month observation period. Conversely, the survival time of allografts in the allogeneic KT group and the allogeneic KT after ST group was (10.0 ± 0.7) days and (6.2 ± 1.1) days, respectively, with a statistically significant difference between the two groups (Figure 2A).

Levels of DSA (IgG and IgM)
Serum DSA-IgG levels in allogeneic KT after ST group recipients showed significant elevation from day 7 post-ST through day 5 post-KT compared with syngeneic KT after ST group recipients. No statistically significant differences in DSA-IgG levels emerged between groups at day 0 and day 3 post-ST. Specifically, at day 7 post-ST, the DSA-IgG level for the allogeneic KT after ST group was significantly higher than control levels (135.0 ± 11.8 vs 76.0 ± 5.6, P < 0.001). This disparity intensified by day 14 post-ST, when DSA-IgG levels were 325.8 ± 34.8 and 75.5 ± 8.7, respectively. Significantly elevated DSA-IgG levels in the allogeneic KT after ST group were sustained from day 1 to day 5 post-KT. The allogeneic KT after ST group exhibited higher DSA-IgM levels than the syngeneic KT after ST group at day 7 (55.5 ± 8.8 vs 34.0 ± 7.7, P < 0.01) and day 14 (57.8 ± 10.4 vs 31.3 ± 7.7, P < 0.01) post-ST. There were no significant elevations at other time points (Figure 2B,C).

Pathological characteristics of renal grafts
Histopathological analysis of renal grafts from the syngeneic KT after ST group at day 5 revealed normal renal architecture, with intact glomeruli and tubules. Peritubular capillaries exhibited continuous and strong linear CD31 expression without intraluminal lymphocyte infiltration, and C4d staining was negative. In contrast, the allogeneic KT after ST group demonstrated a time-dependent progression of AAMR. At 6 h and 12 h after KT, mild glomerulitis, peritubular capillaritis, and intravascular C4d deposition were observed in the allografts. These manifestations gradually worsened over time, and by day 1 and day 2 after KT, the allografts showed moderate glomerulitis, peritubular capillaritis, and intravascular C4d deposition. Furthermore, on the 3rd to 5th day after KT, in addition to severe glomerulitis, peritubular capillaritis, and C4d deposition, the allografts also showed tissue damage, tubular necrosis, and extensive infiltration of inflammatory cells. Additionally, immunohistochemical staining for CD3 revealed moderate infiltration of T lymphocytes, predominantly localized in the renal interstitium, but without the severe tubulitis characteristic of typical acute cellular rejection (Figure 3). These pathological manifestations were consistent with the Banff criteria of AAMR.

Figure 1: Experimental grouping design. The study employed four experimental groups: (1) Syngeneic KT group: Lewis rats served as both donors and recipients; (2) Syngeneic KT after ST: Lewis recipients received a skin graft from a Lewis donor two weeks prior to KT; (3) Allogeneic KT group: BN rats served as donors and Lewis rats as recipients; (4) Allogeneic KT after ST group: Lewis recipients received a skin graft from a BN donor two weeks before KT. The timeline illustrates the experimental schedule, with skin grafting preceding renal transplantation by two weeks. KT, kidney transplantation; ST, skin transplantation; BN, Brown Norway. Please click here to view a larger version of this figure.

Figure 2: Renal allograft survival and DSA dynamics. (A) Renal allograft survival in different transplant groups, including the syngeneic group, syngeneic KT after ST group, allogeneic KT group, and allogeneic KT after ST group (n = 6/group). (B) DSA-IgG levels in syngeneic KT after ST group (n = 4/group) and allogeneic KT after ST group at different time points (n = 4/group). (C) DSA-IgM levels in syngeneic KT after ST group and allogeneic KT after ST group at different time points (n = 4/group). KT, kidney transplantation; ST, skin transplantation; DSA, donor-specific antibody; MFI, mean fluorescence intensity. NS = no significance; **P < 0.01; ***P < 0.001. Please click here to view a larger version of this figure.

Figure 3: Pathological characteristics of renal grafts in the syngeneic KT after ST group and the allogeneic KT after ST group. Renal grafts were obtained at 5 days after KT in the syngeneic KT after ST group and at 6 h, 12 h, 1 d, 2 d, 3 d, 4 d, and 5 d after KT in the allogeneic KT after ST group. These grafts were then stained with HE, PAS, CD31, C4d, and CD3, and representative images are used for display. The arrows refer to the typical pathological change of peritubular capillaritis in AAMR. Scale bar = 100 µm. HE, hematoxylin and eosin; PAS, periodic acid-Schiff; KT, kidney transplantation; ST, skin transplantation. Please click here to view a larger version of this figure.

AAMR remains a significant challenge in KT due to its rapid progression and unfavorable clinical outcomes^33,34,35,36. Even with effective current anti-rejection treatments reversing acute episodes, more than 40% of patients advance to chronic AMR. After a chronic AMR diagnosis, the five-year graft survival rate often drops below 50%. The 4-year graft survival rate in C4d-positive patients is only 50%, whereas in C4d-negative patients it remains about 50% even at 8 years³⁷. The occurrence of AAMR primarily stems from inadequate immunosuppression in recipients, which leads to B-cell activation and production of DSA^{24,29,38,39,40,41,42}. These antibodies, in conjunction with the complement system and other immune cells, collectively damage the graft. However, the specific mechanisms underlying AAMR have not been fully elucidated, and clinical prevention and treatment outcomes remain largely suboptimal⁴³. Thus, there is an urgent need to establish a KT model of AAMR. Such a model would facilitate the exploration of more effective therapeutic strategies and further clarify its pathophysiology²⁹.

This study established an AAMR model in rats by pre-sensitizing ST for 2 weeks, followed by KT. When BN rats were used as donors and Lewis rats as recipients for KT, the recipients had a survival time of 10.0 ± 0.7 days, with acute TCMR as the primary manifestation²⁸. The purpose of pre-sensitization for ST was to increase the level of DSA in the recipient's serum prior to KT, thereby inducing a rapid onset of AAMR following KT. Indeed, on the 7th and 14th days after ST, serum levels of DSA-IgG and IgM increased, with a particularly pronounced rise in IgG. It is worth noting that both DSA-IgG and IgM levels appeared to rise simultaneously at Day 7. Classically, IgM production precedes IgG class switching. However, since the current serological sampling interval spanned from Day 3 to Day 7, it is highly probable that the initial IgM-only elevation phase occurred between these time points (e.g., Days 4-5) and was not captured. Thus, the simultaneous detection at Day 7 likely reflects a stage in which isotype switching to IgG had already initiated, while IgM levels remained elevated. Furthermore, from a technical perspective, we acknowledge that the 1:25 serum dilution used in this study may have been near the detection limit. This concentration could introduce a prozone-like effect, in which high antibody titers interfere with optimal antigen-antibody binding detection, potentially masking an earlier or more pronounced rise in antibody levels. Future studies might benefit from testing serial dilutions to fully characterize the kinetic profile. Regarding the experimental controls, the syngeneic sensitized group was utilized as the primary control for DSA analysis to rule out non-specific inflammatory responses associated with the skin grafting procedure, although comparisons with non-sensitized allogeneic recipients would also demonstrate the efficacy of the sensitization protocol. This elevation in DSA levels precipitated pathological manifestations of mild AAMR (glomerulitis, peritubular capillaritis, and intravascular C4d deposition), which began as early as 6 h after KT. It should be noted that ST not only activates B cells to produce DSA, but also promotes T cell activation. Therefore, more precisely, this model represents a mixed rejection response, predominantly characterized by AAMR. Notably, while CD3 staining confirmed moderate T-cell infiltration in the renal interstitium, the histological hallmarks of severe acute cellular rejection (ACR), particularly extensive tubulitis, were not prominent. This specific pathological presentation supports the hypothesis that the rapid onset of microvascular injury, driven by high levels of preformed DSA and early complement activation, likely induced graft necrosis before the full spectrum of typical T-cell-mediated lesions (e.g., severe tubulitis) could develop. Furthermore, the severe vascular compromise may have restricted further recruitment and organized infiltration of lymphocytes into the graft tissue. Thus, the absence of dominant ACR histology reflects the overwhelming intensity and kinetics of the humoral injury rather than a complete absence of T-cell activation.

This protocol offers distinct advantages over existing methods. Unlike models relying on the passive transfer of anti-MHC serum³⁰, which is costly and produces only transient injury, our active skin pre-sensitization generates a robust, endogenous, and persistent immune response that better mimics the clinical scenario of highly sensitized patients. Furthermore, regarding the surgical technique, standard protocols often employ the Carrel patch method. In contrast, the current approach utilizes a direct end-to-side anastomosis without an aortic patch. While this technique demands greater microsurgical proficiency, it significantly reduces the extent of the arteriotomy and trauma to the recipient's abdominal aorta, thereby minimizing the risk of post-operative lower limb ischemia and improving overall animal survival rates. The stability and reliability of this model are underpinned by strict adherence to protocol-critical parameters. First, the timing of pre-sensitization is paramount. This study established that a 14-day interval between ST and KT is optimal for inducing peak DSA titers at the time of grafting. Deviating from this window may result in variable sensitization levels. Second, minimizing warm ischemia time is crucial. Since severe ischemia-reperfusion injury (IRI) can mimic or mask histological signs of rejection (e.g., tubular necrosis), maintaining warm ischemia time below 30 min is essential for a clear pathological readout. Mastery of these technical variables is the strongest determinant of model consistency.

Several common technical challenges and corrective actions have been identified to ensure reproducibility. Vascular thrombosis at the anastomosis site is a frequent cause of graft failure; this is typically prevented by performing thorough local irrigation of the vessel lumens with heparinized saline. This step not only clears debris but also hydraulically reveals the intima, facilitating precise intimal eversion during the 10-0 suture placement. Ureteral complications, such as obstruction or leakage, can be mitigated by utilizing a "no-touch" dissection technique, where only the peri-ureteral connective tissue is handled, rather than the ureter itself, to preserve the delicate adventitial blood supply. Furthermore, ureter length must be optimized by trimming excess tissue to ensure a straight, tension-free trajectory to the bladder, thereby preventing kinking caused by redundancy. Hind limb paralysis may occur due to prolonged aortic clamping; to avoid this, the aortic incision should not exceed 40% of the circumference. Finally, to prevent hypothermia-induced mortality, it is essential to maintain the animal's body temperature using a heating pad throughout the perioperative period.

Although this model effectively recapitulates the key pathological features of acute AMR, some limitations remain. First, the model primarily relies on ST for immune sensitization, which may differ from certain clinical scenarios. Future studies could explore alternative sensitization methods, such as direct antibody injection or different transplantation models, to further validate the generalizability and reliability of this model. Second, while a detailed assessment of post-transplant pathological changes was conducted, a deeper analysis of immune cell dynamics, particularly the role of B cells, plasma cells, and macrophages in AAMR, requires further investigation. Future research using this model may focus on exploring the roles of B cells, plasma cells, and macrophages in AAMR pathogenesis through targeted immunosuppressive therapies. Additionally, developing alternative pathological models, including those using porcine or primate kidneys, could provide more clinically relevant insights and better simulate human organ transplantation. This protocol shares conceptual similarities with previously established murine models of AMR. For example, Bickerstaff et al. successfully established an acute humoral rejection model in mice using skin presensitization, demonstrating key features such as C3d deposition and elevated alloantibodies⁴⁴. While murine models have significantly advanced our understanding of AMR mechanisms, rat models offer specific advantages in terms of surgical reproducibility and physiological scale. The larger vessel caliber in rats facilitates precise vascular anastomosis with reduced incidence of technical thrombosis, and the larger blood volume allows for serial serum sampling to monitor DSA kinetics without compromising animal health, as demonstrated in this study.

In summary, this study developed a stable and reliable rat model of AAMR, offering a robust experimental tool for studying AAMR pathology and treatment strategies. Through further research and the integration of novel immunotherapies, this model may provide new theoretical foundations and practical guidelines for the early diagnosis and personalized treatment of AAMR.