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

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.

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.

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 widen the therapeutic window. Results demonstrated that the PS + CsA group had significantly prolonged cardiac allograft survival compared to the PS group (5.2 ± 0.4 days vs. 2.8 ± 0.4 days, P < 0.01, Figure 2A).

Levels of DSA (IgG and IgM)
This study next quantified the levels of DSA in the serum of mice from each group. In the NS group, a significant increase in the MFI of DSA-IgG was observed only at 7 days after CT (574.3 ± 42.72 vs. 410.5 ± 57.97, P < 0.05), while no significant change in DSA-IgM MFI was detected at any time point (Figure 2B). In the PS group, serum DSA levels gradually increased following ST. DSA-IgG MFI was significantly elevated by day 7 after ST (823.0 ± 61.47 vs. 444.5 ± 49.13, P < 0.001) and remained high from day 3 after CT onward (778.8 ± 134.4 vs. 444.5 ± 49.13, P < 0.001). DSA-IgM also showed a significant increase by day 3 after CT (79.25 ± 10.81 vs. 53.5 ± 5.0, P < 0.05) (Figure 2C). In the PS + CsA group, although overall DSA MFI values were lower than those in the PS group, DSA-IgG MFI was still significantly higher at day 7 after ST (642.5 ± 33.76 vs. 444.3 ± 52.73, P < 0.001) and remained elevated from day 5 after CT (656.5 ± 45.04 vs. 444.3 ± 52.73, P < 0.001) (Figure 2D). In contrast, no significant changes in DSA-IgM were observed throughout the post-ST period.

Pathological characteristics of cardiac grafts
Finally, this study conducted a comparative analysis of the graft pathological characteristics in each group, with grading performed according to the International Society for Heart and Lung Transplantation (ISHLT) standards.

In the NS group, focal inflammatory cell infiltration was observed in the cardiac allografts starting on postoperative day 1. The majority of cardiomyocytes showed no significant abnormalities, consistent with grade 1R acute T cell-mediated rejection (aTCMR). H&E staining revealed no significant capillary dilation or mononuclear cell infiltration within the microvasculature, and immunohistochemistry showed no evident C4d deposition in the microvessels. By postoperative day 2, multifocal inflammatory infiltration was noted, accompanied by cardiomyocyte necrosis, interstitial edema, and microvascular dilation, while C4d staining remained faint. The injury progressively worsened, and by day 5, diffuse infiltration of mononuclear cells and eosinophils was observed, along with extensive cardiomyocyte damage and interstitial edema. The pathological changes were primarily characterized by aTCMR, accompanied by only scant C4d deposition on immunohistochemistry and inconspicuous microvascular damage on H&E staining.

In the PS group, as early as 12 h post-transplantation, immunohistochemistry revealed prominent C4d deposition along the microvascular walls. By postoperative day 3, marked endothelial swelling, nuclear pyknosis, and severe interstitial edema were evident. In addition, aTCMR persisted throughout, characterized by diffuse inflammatory infiltration and cardiomyocyte injury in the myocardium, potentially confounding mechanistic analyses of AMR.

To mitigate the influence of aTCMR, recipient mice in the PS + CsA group received subcutaneous CsA starting on the day of ST. In this group, inflammatory cell infiltration in the myocardium was markedly reduced at 12 h compared to the PS group, with detectable C4d deposition. By postoperative day 1, inflammatory infiltration gradually emerged and increased over time; however, the severity of aTCMR remained lower than in the PS group, indicating that CsA effectively suppressed aTCMR in the acute AMR model. Moreover, significant microvascular inflammation and C4d deposition were observed in this group by day 3, confirming that CsA did not interfere with the development of acute AMR (Figure 3, and Figure 4A,B).

Figure 1: Experimental grouping design. The study employed three experimental groups: (1) Non-sensitized group: Balb/c mice were used as the donors and C57BL/6 mice as the recipients for CT. (2) Pre-sensitized group: Balb/c mice were used as the donors and C57BL/6 mice as the recipients for ST, followed by CT one week later. (3) Pre-sensitized + CsA group: Balb/c mice were used as the donors and C57BL/6 mice as the recipients for ST, followed by CT one week later. From the day of ST, the recipient mice were injected subcutaneously with cyclosporine A at 20 mg/(kg·d) until graft failure or graft specimen collection. CT, cardiac transplantation; ST, skin transplantation; CsA, cyclosporine A. Please click here to view a larger version of this figure.

Figure 2: Cardiac allograft survival and DSA dynamics. (A) Cardiac allograft survival in different transplant groups, including the NS group, PS group, and PS + CsA group (n = 4/group). (B) DSA-IgG and IgM levels in the NS group at different time points (n = 4/group). (C) DSA-IgG and IgM levels in the PS group at different time points (n = 4/group). (D) DSA-IgG and IgM levels in the PS + CsA group at different time points (n = 4/group). CT, cardiac transplantation; ST, skin transplantation; CsA, cyclosporine A; DSA, donor-specific antibody; MFI, mean fluorescence intensity; NS, non-sensitized; PS, pre-sensitized. NS = no significance; *P<0.05; **P < 0.01; ***P < 0.001. Please click here to view a larger version of this figure.

Figure 3: Pathological characteristics of cardiac grafts in the NS group, PS group, and PS + CsA group. Grafts are collected at 6 h, 12 h, 1 day, 2 days, 3 days, 4 days, and 5 days after cardiac transplantation. After collection, the grafts are stained with hematoxylin-eosin and C4d, and representative images are used for display. Pathological changes of the grafts are observed under a 400x light microscope. Scale bar: 50 µm. Please click here to view a larger version of this figure.

Figure 4: Pathologic AMR grade and C4d grade in the NS group, PS group, and PS + CsA group. (A) Cardiac graft pathologic injury due to AMR was graded in the NS, PS, and PS + CsA groups according to ISHLT standards. (B) The intensity of C4d deposition in cardiac grafts was graded across the NS, PS, and PS + CsA groups based on ISHLT standards. AMR, antibody-mediated rejection; CsA, cyclosporine A; NS, non-sensitized; PS, pre-sensitized. Please click here to view a larger version of this figure.

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 developed typical AMR lesions, including microvascular injury and C4d deposition, as early as postoperative day 2, accompanied by a marked increase in DSA. Compared with the PS group, PS + CsA recipients exhibited prolonged graft survival and reduced aTCMR severity. Collectively, these findings confirm the successful establishment of a reproducible and specific acute AMR model, providing a practical platform for future mechanistic and therapeutic studies.

The mouse CT model has a research history of over 50 years, progressing from nonvascularized to vascularized techniques²⁶. Compared with kidney or intestinal transplantation, its surgical procedure is technically less demanding, and graft function can be directly assessed by postoperative observation and palpation. Therefore, this model is widely applied in mechanistic studies of solid organ transplantation. Because mice of the same strain share identical MHC genotypes and do not experience rejection, transplantation is generally performed between different strains²⁷. In this study, C57BL/6 mice, characterized by high complement activity, were selected as recipients. In conventional CT models, AMR typically develops later and presents as a chronic process, posing challenges for experimental analysis. To establish an early-onset acute AMR model, this study used ST for pre-sensitization to induce DSA production prior to CT.Tail skin from the donor mouse is routinely used for transplantation and must include both epidermis and dermis, with subcutaneous adipose tissue thoroughly removed. The graft size should closely match the recipient bed to avoid necrosis or poor vascularization. After surgery, the wound is bandaged to prevent graft displacement from gnawing, and serum DSA levels are monitored regularly to evaluate sensitization. If DSA shows no significant rise, technical errors are first excluded before inspecting the graft for rejection signs such as redness, scabbing, or shrinkage. In the absence of both clinical rejection and detectable DSA, a donor of a different strain may be selected or the observation period extended prior to repeat DSA testing. In the study, serum DSA levels in the PS group were significantly higher than those in the NS group on day 7 after ST.

CT was performed following confirmed elevation of DSA in recipient mice. In the study, 6 h after CT, the PS group exhibited markedly greater C4d deposition in cardiac tissue than the NS group, confirming that ST effectively induces DSA and accelerates the onset of acute AMR after CT. Vascular anastomosis constitutes the most critical step in CT. This procedure involves adequate mobilization of the recipient abdominal aorta and inferior vena cava, along with careful positioning of the donor heart to prevent vascular torsion during suturing. The anastomosis is typically performed using a two-point positioning technique with six interrupted sutures. Prior to restoring blood flow, the anastomotic site is compressed with a sterile cotton swab and is released gradually only after confirming the absence of significant bleeding. Should anastomotic bleeding occur, compression is maintained until hemostasis is achieved. Graft pulsation is then assessed daily via abdominal palpation. If pulsation is absent, the abdomen is reopened to evaluate graft viability.

In the PS group, acute AMR manifested as early as 12 h post-transplantation. The process begins when antigen-presenting cells recognize donor HLA and activate CD4⁺ T cell-dependent B cell responses, leading to DSA production^28,29. Subsequently, DSA binding to donor endothelial cells triggers complement activation: IgM and IgG1-3 initiate the classical pathway, as evidenced by C4d deposition, whereas IgG4 activates the alternative pathway, forming the membrane attack complex (C5b-9) that directly damages the endothelium^30,31,32. Complement activation further amplifies inflammation through opsonization (C3b, C4b) and recruitment of phagocytes mediated by anaphylatoxins (C3a, C5a)^33,34. These mechanisms were supported by our histopathological findings of microvascular endothelial swelling, mononuclear infiltration, and C4d deposition within the allografts.

However, during acute allograft rejection, aTCMR often predominates³⁵. In this study, the PS group showed inflammatory cell infiltration and cardiomyocyte injury in the allografts, consistent with aTCMR pathology. The excessive rejection intensity in this group led to rapid graft loss, limiting subsequent mechanistic investigation. CsA, a calcineurin inhibitor, is routinely used for post-transplant immunosuppression and effectively suppresses T cell activation, proliferation, and cytokine release³⁶. To prolong graft survival in pre-sensitized recipients and minimize T cell-mediated interference, CsA was administered subcutaneously from the day of ST until graft harvest or failure. In the PS+CsA group, graft survival was prolonged to 6 days, accompanied by markedly reduced inflammatory cell infiltration. This approach expanded the experimental window for acute AMR, decreased aTCMR interference, and created optimal conditions for exploring AMR pathogenesis and potential interventions.

Compared with other methods for establishing acute AMR animal models, the ST combined with subcutaneous CsA injection offers higher efficiency, simplicity, and excellent specificity. In 2006, Qian et al. pre-sensitized rats via nonautologous blood transfusion three weeks before CT, yet acute rejection only occurred on the fourth postoperative day, reflecting a delayed and relatively mild response²⁰. In 2022, Mang et al. injected Lewis rat splenic lymphocytes into Brown Norway rats two weeks before liver transplantation; although the pre-sensitized group showed more severe rejection than the non-sensitized group when examined two weeks post-transplantation, the protocol remains complex and timeconsuming²¹. In contrast, ST as a pre-sensitization approach requires only basic surgical skills and is far simpler to perform. Moreover, it induces a robust sensitization, with significantly elevated serum DSA levels observed as early as 7 days after transplantation and severe cardiac allograft rejection occurring in the early post-transplant period, thereby facilitating interventional studies.

It must be acknowledged that this animal model has several inherent limitations. Although CsA was used as a T cell inhibitor, residual aTCMR effects could not be entirely excluded. Future studies could employ more potent and targeted T lymphocyte-depleting agents to enhance model specificity³⁷. Additionally, in the pre-sensitized groups, none of the cardiac allografts survived beyond 6 days, and pathological progression from acute to chronic lesions was not observed. This limitation restricts the investigation of mechanisms underlying the acute-to-chronic transition and the development of preventive strategies. Using recipient strains with slightly lower complement activity instead of C57BL/6 mice may help extend graft survival and broaden the model's applicability.

In summary, this study provides a practical experience summary and pathological analysis for the establishment of the mouse CT acute AMR model, and demonstrates the feasibility of using ST as a pre-sensitization method and proposes CsA as an effective drug to extend recipient mouse survival time and suppress T cell interference. In future studies, targeted T lymphocyte-depleting agents could be used instead of CsA to enhance model specificity, while recipient strains with moderately lower complement activity may help prolong graft survival and adapt the model to various experimental objectives.