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

Summary

This article describes the establishment of a rat model of acute antibody-mediated rejection (AAMR) in kidney transplantation. The model is induced by presensitizing recipient Lewis rats with skin grafts from donor Brown Norway rats 14 days prior to kidney transplantation, providing a robust platform for studying AAMR pathophysiology.

Abstract

Acute antibody-mediated rejection (AAMR) remains a major obstacle to long-term graft survival in kidney transplantation. The pathophysiology of AAMR is primarily driven by donor-specific antibodies (DSA) that trigger complement activation, leading to endothelial cell injury, vasculitis, and graft dysfunction. Despite advancements in immunosuppressive therapies, the treatment of AAMR remains suboptimal. Therefore, the development of reliable animal models for AAMR is crucial for understanding its mechanisms and evaluating potential therapeutic interventions. In this study, a rat model of AAMR was established by sensitizing recipient Lewis rats with donor Brown Norway rat skin grafts for 14 days prior to kidney transplantation. No immunosuppressive agents were administered to the recipients throughout the study. Survival analysis demonstrated significantly reduced graft survival in the presensitized allogeneic group compared with the non-presensitized group (6.2 ± 1.1 d vs 10.0 ± 0.7 d, P < 0.001). The levels of serum DSA-IgG significantly increased from the 7th day after skin transplantation and continued to rise until the 5th day after kidney transplantation, and the levels of serum DSA-IgM significantly increased on the 7th and 14th days after skin transplantation. Starting 6 h after kidney transplantation, the allografts began to show signs of glomerulitis, peritubular capillaritis, and C4d deposition in capillaries. Starting from 3 days after kidney transplantation, significant allograft damage and tubular necrosis were observed. These changes gradually worsened over time and are all consistent with the characteristics of AAMR. This model effectively recapitulates the key features of AAMR, providing a robust platform for future studies on underlying mechanisms and potential treatments.

Introduction

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.

Protocol

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.

Results

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 o...

Discussion

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 on...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-CD31 Monoclonal Antibody	Abcam (UK)	ab64543	For immunohistochemistry (labeling vascular endothelial cells) Clone number: TLD-3A12
Anti-C4d  Antibody	Hycult Biotech (Netherlands)	HP8034	For immunohistochemistry (detecting complement deposition), dilution is 1:50 Clone number: NA (Polyclonal)
FITC-Conjugated Anti-Rat IgG Antibody	Abcam (UK)	ab6840	Used for flow cytometric quantification of DSA-IgG levels Clone number: Polyclonal
Gas Anesthesia System	Nanjing Calvin Biotechnology Co., Ltd.	KW-MZJ-4	Equipped with isoflurane for general anesthesia Clone number: NA (Polyclonal)
PE-Conjugated Anti-Rat IgM Antibody	Bio Legend	408918	Used for flow cytometric quantification of DSA-IgM levels Clone number: MRM-47
Stereotaxic Surgical Microscope	Beijing Zhongtian Guangzheng Technology Co., Ltd.	TS-39NK	Used for performing rat renal transplantation surgery Clone number: NA
Vascular Bulldog Clamps	ROBOZ SURGICAL INSTRUMENT CO.	RS-5481T	Used to block blood flow during renal transplantation Clone number: NA