Establishment and Analysis of Acute Antibody-Mediated Rejection in Murine Cardiac Transplantation

Summary

This study established a practical mouse model of acute antibody-mediated rejection after cardiac transplantation, characterized by robust DSA production, typical pathological changes, and moderate allograft survival.

Abstract

A major contributor to allograft failure in cardiac transplant recipients is antibody-mediated rejection (AMR). The mouse AMR model, therefore, serves as a vital tool for deciphering its underlying mechanisms and fostering the development of innovative treatments. In this study, the recipient mice were divided into three groups: non-sensitized (NS), pre-sensitized (PS), and pre-sensitized with cyclosporine A treatment (PS + CsA). The NS group, which exhibited a mean allograft survival of 6.8 ± 0.7 days, showed no rise in serum DSA levels and negative allograft C4d staining within four days post-cardiac transplantation (CT), suggesting a pathology dominated by acute cellular rejection. This study established an acute AMR model by pre-sensitizing recipients with skin transplantation (ST) one week before CT, thereby pre-activating the immune system. This approach successfully induced a severe AMR phenotype, as evidenced by a short allograft survival of 2.8 ± 0.4 days, a significant rise in DSA-IgG levels post-ST and post-CT, and early pathological hallmarks of vasculitis and extensive C4d deposition within 12 h of CT. Nevertheless, the extreme severity of this model constrains its broader application. To minimize the concurrent T cell activation induced by ST and establish a more specific acute AMR model, this study administered cyclosporine A. Consequently, the PS + CsA group exhibited an allograft survival time of 5.2 ± 0.4 days. Serum DSA-IgG levels were significantly elevated by day 7 post-ST and remained high within five days post-CT. Pathological assessment on day 2 post-CT confirmed significant vasculitis and C4d deposition, findings which collectively meet the diagnostic criteria for moderately severe acute AMR. In conclusion, this study established a highly practical and translatable mouse model of acute AMR following CT, defined by robust DSA production, characteristic pathological changes, and moderate allograft survival.

Introduction

Cardiac transplantation (CT) remains the gold-standard therapy for end-stage heart disease¹. Although median post-transplant survival now exceeds 13 years, long-term graft failure remains inevitable². Antibody-mediated rejection (AMR) is a major contributor to late graft loss after CT³. Studies have shown that the incidence of graft loss due to AMR exceeds that caused by T cell-mediated rejection, and the risk of graft failure in late-onset AMR is approximately twice that of early AMR^4,5. Therefore, early identification and timely intervention for AMR are critical to improving graft survival.

AMR is driven by donor-specific antibodies (DSA), which cause graft failure through complement activation, vascular inflammation, and ischemic injury⁶. DSA binding to vascular endothelium initiates endothelial damage, promotes thrombosis, and induces both acute and chronic inflammatory responses. The principal mechanisms of DSA-mediated injury involve complement-dependent cytotoxicity and antibody-dependent cellular cytotoxicity pathways⁷. Although targeted therapies such as the anti-C5 antibody eculizumab and the B-cell depleting agent rituximab have been developed, clinical outcomes in late-stage AMR remain unsatisfactory^7,8,9. Therefore, further mechanistic investigation using acute animal models is essential to better understand disease progression and identify new therapeutic targets.

The mouse vascularized heterotopic CT model is a well-established platform for studying CT, particularly in ischemia-reperfusion injury and rejection^10,11,12. Graft function is typically monitored by daily abdominal palpation of the heartbeat intensity, which reflects functional status and defines graft failure¹³. However, conventional models often fail to detect AMR onset shortly after transplantation and therefore require pre-sensitization to induce DSA formation and trigger acute AMR. Furthermore, mouse strains differ in immune reactivity; for example, C57BL/6 mice exhibit stronger complement activation, whereas BALB/c mice favor Th2-skewed responses^14,15. Consequently, both the pre-sensitization strategy and recipient strain selection are critical determinants of model performance.

In previous studies, this study team used different rat strains for renal transplantation following skin transplantation (ST), established an AMR model, and analyzed its characteristics, accumulating considerable experience^16,17,18. In 2021, this team further established an ST-induced acute AMR mouse model in CT for fundamental research, which demonstrated great stability¹⁹. Unlike the relatively inefficient pre-sensitization by allogeneic blood transfusion²⁰ or the more complex and time-consuming spleen lymphocyte infusion²¹, ST offers a robust, stable, and simpler sensitization protocol. This model facilitates early post-transplant detection of elevated serum DSA levels and observable allograft injury from acute AMR, making it particularly suitable for interventional studies on acute AMR. However, the acute AMR induced by ST is typically severe, causing substantial tissue damage that often masks the progression of chronic injury. This study aims to further provide a detailed description of the methodology and procedures used to establish this mode, analyze the immunological and pathological characteristics of the model, and summarize practical experience.

Protocol

All animal experiments were conducted in strict accordance with Guangdong experimental animal management regulations, the Declaration of Helsinki, and the 3Rs principles. The experimental protocol was approved by the Committee of Guangzhou Jennio Biotech Co., Ltd. (approval number JENNIO-IACUC-2024-A027). Male Balb/c and C57BL/6 mice (6-8 weeks old, weighing 20-25 g), all specific pathogen-free (SPF) grade, were obtained from a commercial source. The reagents and equipment used in this study are detailed in the Table of Materials.

1. Animal preparation

1. House the SPF grade mouse in the appropriate animal facility sytem.
2. Divide the mice into three experimental groups as described in Figure 1
   1. Non-sensitized group (NS): Use Balb/c mice as the donors and C57BL/6 mice as the recipients for CT.
   2. Pre-sensitized group (PS): Use Balb/c mice as the donors and C57BL/6 mice as the recipients for ST, followed by CT one week later.
   3. Pre-sensitized + cyclosporine A group (PS + CsA): Use Balb/c mice as the donors and C57BL/6 mice as the recipients for ST, followed by CT one week later. From the day of ST, inject recipient mice subcutaneously with CsA at 2 mg/(kg·d) until graft failure or graft specimen collection.

2. Preoperative preparation

1. Ensure that the donor mice are not deprived of food or water before surgery, while the recipient mice are fasted for 8 h but not deprived of water.
2. Anesthesia
   1. Place the mouse in an anesthesia induction chamber for induction (Isoflurane concentration: 3%-4%; Flow rate: 300-500 mL/min) (following institutionally approved protocols). Shake the induction chamber to check if the animal is fully anesthetized.
      NOTE: If the animal is flipped onto its side and does not attempt to return to a prone position, it indicates complete anesthesia.
   2. After adjusting the maintenance concentration on the surgical platform, remove the mouse from the induction chamber, fix its nose in the anesthesia mask (Isoflurane concentration: 1.0%-2.5%; Flow rate: 100-200 mL/min), and check whether the mouse is fully anesthetized (press the paw or tail with a finger; if there is no response, it indicates full anesthesia).

3. Presensitization via ST (for Groups 2 and 3)

1. Donor tail skin harvesting
   1. Disinfect the tail skin of the anesthetized donor mice with 75% alcohol.
   2. Use a scalpel to amputate the tail (at the distal 1/3), fix the stump with forceps in the left hand, and use the scalpel in the right hand to cut through the skin between the tail veins, exposing the underlying white tendon. Then, fix the tailbone with forceps, use curved forceps to clamp the skin at the incision, and carefully strip it off.
   3. Place the peeled skin into a beaker containing sterile ice-cold saline, with the tissue facing down. Cut a 1.0 cm × 1.0 cm piece of skin for later use.
2. Recipient dorsal ST
   1. Shave the fur from the dorsal area of the anesthetized recipient mice using a hair clipper, disinfect with 75% alcohol, and place a sterile drape.
   2. Use a scalpel to incise the skin, grip the skin edges with toothed forceps, and cut a piece of skin slightly larger than 1.0 cm × 1.0 cm with tissue scissors to create the graft bed.
   3. Place the donor mouse skin graft on the graft bed with the tissue side down. After trimming the graft, suture the donor skin to the recipient skin under a microscope using 8-0 nylon thread, ensuring the donor skin is evenly placed on the graft bed.
   4. Cover the wound with a sterile adhesive bandage, and place the mouse in a housing cage after it regains consciousness.

4. CT

1. Donor heart harvesting
   1. Shave the fur from the chest and abdomen of the anesthetized donor mice using a hair clipper, disinfect with 75% alcohol, and place a sterile drape.
   2. Make a midline abdominal incision with a scalpel, exposing the upper segment of the inferior vena cava, and inject 1 mL of 50 U/mL heparinized cold saline (4 °C) into the inferior vena cava using a syringe.
   3. Use tissue scissors to open the chest cavity and expose the heart. Isolate the ascending aorta and sever it before its first branch; isolate the pulmonary artery and transect it at its bifurcation; ligate the inferior vena cava and the connective tissue at the pulmonary hilum using 6-0 non-traumatic sutures; cut the distal tissue at the ligation site with tissue scissors and remove the heart.
   4. Quickly place the harvested heart into 2 mL of hypertonic citrate adenine (HCA) solution and store it in an ice-water mixture for later use. Euthanize the donor mouse using the cervical dislocation method.
2. Recipient CT
   1. Shave the fur from the abdomen of the anesthetized recipient mice using a hair clipper, disinfect with 75% alcohol, and place a sterile drape.
   2. Lift the skin with toothed forceps and make a midline incision in the abdominal wall from the xiphoid process to the bladder base using tissue scissors. Place sterile, moist gauze on both sides of the incision to protect it.
   3. Use sterile cotton swabs to push the intestinal loops to the right side of the incision and wrap them with moist gauze. After bluntly dissecting the peritoneum, expose the inferior vena cava and abdominal aorta, ligate the lumbar veins, and use a vascular clamp to stop blood flow to the distal and proximal ends of the aorta and vena cava.
   4. Under a microscope, use micro-scissors to incise the venous wall by about 2 mm, matching the diameter of the donor cardiac pulmonary artery; similarly, incise the arterial wall by 1-2 mm, matching the diameter of the donor cardiac ascending aorta.
   5. Take the donor heart and perform an end-to-side anastomosis between the donor cardiac pulmonary artery and the recipient's inferior vena cava using 12-0 non-traumatic sutures. Similarly, perform end-to-side anastomosis between the donor cardiac ascending aorta and the recipient's abdominal aorta (when tying the last knot, use saline to flush air from the lumen; intermittently drip 4 °C saline onto the surface of the donor heart during anastomosis to cool it).
   6. Release the vascular clamp and restore blood supply to the heart (normal filling of the coronary arteries is visible, the heart turns pink, and it resumes beating within 1-2 min). After reinserting the intestinal loops, close the abdominal cavity with 4-0 nylon sutures.
      NOTE: If intraoperative bleeding exceeds 0.5 mL, administer normal saline subcutaneously to maintain fluid volume.
   7. Postoperatively, place the mice on a 37 °C warming platform for monitoring. Once the recipient mouse regains consciousness, place it back into the housing cage. The mouse resumes normal eating and drinking postoperatively.

5. Postoperative care and monitoring

1. General condition observation
   1. Ensure daily observation of the mouse's pain status after surgery, whether there is reduced appetite, decreased activity, excessive licking or scratching of the wound, or increased aggression.
   2. Ensure daily observation of the surgical wound after surgery by checking for redness, swelling, exudate, or purulent infection symptoms. If necessary, clean and disinfect the wound.
2. Observation of cardiac graft survival
   1. Ensure daily observation and palpation to check and record the strength of the transplanted cardiac heartbeat. If necessary, anesthetize the mouse and make an incision to observe the cardiac beating.
      NOTE: When the heartbeat of the transplanted heart stops completely, it is recorded as graft failure, and the survival time is noted.

6. Postoperative assessments

1. DSA level detection
   1. In the NS group, obtain blood samples on postoperative days 0, 4, and 7 after CT to monitor DSA levels following transplantation. In the PS group, collect samples on days 0, 4, and 7 after ST and on days 1 and 3 after CT to assess DSA dynamics after pre-sensitization. In the PS+CsA group, collect blood on days 0, 4, and 7 after ST and on days 1, 3, and 5 after CT to evaluate the effects of CsA on DSA levels.
   2. Centrifuge the whole blood sample from the recipient mouse and collect the serum (2000 x g, 10 min, 4 °C); dilute the serum 10 times with PBS and incubate it with donor mouse spleen cells at 37 °C for 30 min.
   3. After centrifugation at 350 × g and 4 °C for 5 min, discard the supernatant. Wash the cells by gently pipetting in 1 mL of PBS, and repeat this wash step twice after subsequent centrifugation.
   4. Incubate the cells with fluorescein isothiocyanate (FITC) anti-mouse IgG antibody and phycoerythrin (PE) anti-mouse IgM Antibody at 4 °C for 1 h.
   5. After washing the cells with PBS, resuspend them to a concentration of 5 × 10⁶ /mL.
   6. Use flow cytometry to detect the two antibodies separately and analyze the mean fluorescence intensity (MFI) to assess the DSA levels (IgG, IgM) in the recipient mouse (Parameter Settings: Acquire 10,000 events for analysis. Use FSCA vs. SSCA to exclude debris, then apply FSCA vs. FSCH gating to remove doublets. Include a negative control and adjust the voltage of each channel to position the negative population within the 10⁰-10¹ range).
2. Histopathological examination of cardiac graft
   1. Collect grafts at 6 h, 12 h, 1 day, 2 days, 3 days, 4 days, and 5 days after CT. After collection, fix the grafts in 4% paraformaldehyde, routinely embed in paraffin, section, and stain with hematoxylin-eosin (H&E). Observe the pathological changes of the grafts under a 400x light microscope.
3. C4d staining
   1. Cut the paraffin-embedded specimens into 4 µm sections, flatten the sections in water at 42 °C, and dry them in an oven at 60 °C for 30 min.
   2. Deparaffinize and hydrate the tissue by sequentially using xylene I, xylene II, ethanol, and deionized water.
   3. Immerse the sections in EDTA antigen retrieval solution, heat at 95-100°C for approximately 15 min, and cool to room temperature.
   4. Incubate the sections with 3% hydrogen peroxide solution at room temperature for 10-15 min to inactivate endogenous peroxidase.
   5. Dilute the anti-C4d antibodies (1:200) with antibody diluent and incubate with the sections at 4 °C overnight. After incubation, wash the sections with PBS.
   6. Add the corresponding secondary antibodies and incubate at room temperature for 30 min, then wash with PBS.
   7. Add DAB chromogen solution and incubate at room temperature for 30 s for coloration.
   8. Perform counterstaining with hematoxylin, dehydrate with ethanol, immerse in xylene, and mount with neutral resin.
   9. Observe the staining results of the grafts under a 400x light microscope.

7. Statistical analysis

1. Use mean ± standard deviation to present all data.
2. Use the t-test for group comparisons of continuous data with a normal distribution, while use the Mann-Whitney U test for group comparisons of continuous data that do not follow a normal distribution.
3. Use Kaplan-Meier analysis to compare the survival time of cardiac allografts between groups.
4. Use SPSS version 27.0 to statistically analyze all data, and consider a P-value of <0.05 as statistically significant.

Results

Cardiac allograft survival time
Upon successful establishment of the mouse model, graft survival times across groups were assessed using Kaplan-Meier analysis. As anticipated, mice in the PS group exhibited significantly shorter mean survival of cardiac allografts compared to those in the NS group (2.8 ± 0.4 days vs. 6.8 ± 0.7 days, P < 0.01), attributable to acute AMR. To facilitate future evaluation of therapeutic strategies for acute AMR, CsA was administered to prolong allograft survival and w...

Discussion

Currently, treatments for late-stage AMR remain ineffective, and reliable methods for its early diagnosis are lacking^22,23,24,25. To address this gap, this study established an acute AMR mouse model to facilitate mechanistic investigation. Recipients were pre-sensitized with ST and administered CsA to suppress concomitant aTCMR. Pathological analysis showed that allografts in the PS+CsA group d...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-C4d Monoclonal Antibody	Hycult Biotech (Netherlands)	HP8034	For immunohistochemistry (detecting complement deposition)
Complete set of microsurgical instruments	Guangzhou Qihua Medical Equipment Co., Ltd.		Used for performing mouse cardiac transplantation surgery
Continuous zoom stereoscopic surgical microscope	Beijing Zhongtian Guangzheng Technology Co., Ltd.	TS-39NK	Used for performing mouse cardiac transplantation surgery
Cyclosporine A	Novartis Pharmaceuticals (Switzerland)	H20100673	Used for inhibiting TCMR in recipient mice
FITC anti-mouse IgG Antibody	Bio Legend (USA)	406001	Used for flow cytometric quantification of DSA-IgG levels
Flow cytometer	Becton Dickinson, USA	BD FACScalibur	Used for assessment of serum DSA levels in recipient mice
PE anti-mouse IgM Antibody	Bio Legend (USA)	406507	Used for flow cytometric quantification of DSA-IgM levels
Small animal gas anesthesia system	Anhui Zhenghua Bio-Instrument Equipment Co., Ltd.	ZH-MZJ	Equipped with isoflurane for general anesthesia
Vascular Bulldog Clamps	ROBOZ SURGICAL INSTRUMENT CO.	RS-5481T	Used to block blood flow during cardiac transplantation