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Ann Thorac Surg 2000;70:1238-1245
© 2000 The Society of Thoracic Surgeons

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Original articles: cardiovascular

T lymphocytes mediate leaflet destruction and allograft aortic valve failure in rats

^a Department of Surgery and Dalhousie University, Halifax, Nova Scotia, USA
^b Department of Microbiology and Immunology, Dalhousie University, Halifax, Nova Scotia, Canada

Address reprint requests to Dr Ross, IWK Grace Health Centre 5850/5980 University Ave, Halifax, NS B3J 3G9, Canada;
e-mail: dross{at}iwkgrace.ns.ca

	Abstract

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
Background. Allograft heart valves are commonly used in cardiac surgery but ultimately fail. This situation is most acute in children. This study addresses the role of T cell–mediated immune damage in allograft valve failure.

Methods. Syngeneic (Lewis to Lewis) or allogeneic (Brown Norway to Lewis) aortic valve grafts were implanted infrarenally into Lewis rat recipients (n = 24). Allogeneic valve grafts were also implanted into T cell–deficient rats (nude; n = 12). At 7, 14, and 28 days the valves were explanted and examined for structural integrity and cellular infiltration.

Results. Syngeneic grafts maintained normal leaflet structure with little leaflet immune infiltration. Allografts showed leaflet infiltration (7 days), significant leaflet thickening, progressively decreased cellularity (14 days), and leaflet destruction (28 days). Infiltrates contained CD43+, CD3+, and CD8+ cells. Allografts in T cell–deficient rats showed none of the above changes and maintained normal structural integrity.

Conclusions. Allograft heart valves in the rat model undergo T cell–mediated immune rejection, resulting in structural failure.

	Introduction

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
Allograft valves were introduced for aortic valve replacement in 1962 [1] and for reconstruction of the right ventricular outflow tract in 1966 [2]. Allografts have many advantages over commercially available mechanical and bioprosthetic valves: low transvalvar gradients, low rates of thromboembolism, and resistance to infection [3]. They are used extensively for children who are prone to rapidly calcifing xenograft valves and for whom warfarin use is difficult. Despite their advantages, allograft valves ultimately do fail in the aortic position. The freedom from valve-related death or reoperation at 10 years is approximately 70%, although there is some evidence of improved outcome using cryopreserved valves [4, 5]. Aortic allografts in children, in particular, have been reported to deteriorate at an unacceptably high rate. In children less than 3 years of age allograft failure may be as high as 70%, with replacement necessary at a mean interval of 1.9 years after the original operation [6].

The mechanisms involved in graft failure remain unclear despite mounting evidence that aortic valve allografts are antigenic and elicit an immune response [7–9]. Animal models have shown that allogeneic valved con-duits induce accelerated rejection of subsequent donor-specific skin grafts [10]. This indicates that valve allografts induce donor-specific allosensitization. Aortic valve allografts have been shown to result in systemic sensitization, as measured by increased splenic mixed lymphocyte reaction and increased frequency of splenic cytotoxic T lymphocyte (CTL) precursors in rats [11]. Data from our laboratory have demonstrated that allogeneic, but not syngeneic, valve grafts undergo progressive destruction over time. There is also evidence of immune activation by valve grafts from human studies. Donor-specific immune antibody to class I and II human leukocyte antigens and donor-specific systemic T cell alloreactivity have been demonstrated after valve graft implantation [12, 13].

Based on this evidence of immune activation by valve grafts and on the current knowledge of solid organ allograft rejection, we hypothesized that T lymphocytes and, in particular, CTL are the effector cells involved in allograft valve destruction. This is supported by some results obtained from human autopsy specimens that have identified T lymphocytic infiltration in failed allografts [14].

The availability of genetically identical inbred strains make small animal models essential in evaluating the role of immunologic processes involved in allograft transplantation. Rat models of infrarenal aortic valve implantation are well described and have provided important advances in our understanding of allograft failure [7, 15]. The objective of the current study was to evaluate the role of T cell–mediated immune damage in allograft valve failure in a rat model.

	Material and methods

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
Animals
Inbred male Brown Norway (RT1.Aⁿ), Lewis (RT1.A^l), and nude (homozygous rnu/rnu) rats weighing 250 to 350 g were purchased from Harlan Sprague Dawley (Indianapolis, IN) and housed in the Medical Sciences animal care facility with food and water ad libitum for 1 week before experimentation in accordance with the guidelines of the Canadian Council of Animal Care [16].

Aortic valve allograft implantation
The technique of transplanting aortic valve allograft was performed as first described by Yankah and colleagues [7]. An intraperitoneal injection of sodium pentobarbital (65 mg/kg) was used to anesthetize the rats.

Donor operation
A midline upper abdominal incision was made and extended as a median sternotomy. The aortic valve containing some myocardium and approximately 8 mm of ascending aorta were dissected free, and the coronary ostia were ligated with 9-0 nylon sutures (Sharpoint, Reading, PA). The graft was then rinsed with cold isotonic saline. Both the donor and recipient operations were performed using a Weck OM-1206 mounted operating microscope (Zeiss, Oberkochen, Germany).

Recipient operation
A midline laparotomy was performed, the bowel eviscerated to the right, and the abdominal aorta exposed. The aorta was mobilized from the level of the renal artery to the aortic bifurcation and divided between clamps. The aortic valve allograft was then anastamosed between the stumps of the recipient aorta using interrupted 9-0 nylon sutures (Sharpoint), flushing intermittently with heparinized saline. Once hemostasis was secured the abdominal contents were returned into the peritoneal cavity and the wound closed in layers. Strain combinations for experimentation consisted of syngeneic (Lewis to Lewis; n = 12), allogeneic (BN to Lewis; n = 12), and athymic (BN to nude; n = 12) transplants.

Tissue analysis
Tissue was harvested for histology at 7, 14, and 28 days and was immersion-fixed (10% formaldehyde) at 4°C for 12 hours. Grafts were then paraffin-embedded and 5-µm sections were cut for histology (hematoxylin and eosin, Verhoeff elastin), immunocytochemistry, and TUNEL studies.

Tissue sections were stained for histology using hematoxylin and eosin (Harris hematoxylin) or Verhoeff elastin stain (hematoxylin 5%, ferric chloride 10%, Lugol’s iodine). Tissue slides from each animal were then examined by light microscopy (Nikon, Optophot, Canada Inc) and images digitalized and captured (JVC digital camera, TK1070U). Morphometric analysis was carried out using Adobe Photoshop (Microsoft, Redmond, WA) and National Institutes of Health Image analyzer software on a Power PC (G3, Apple Computer, Cuppertino, CA) to measure surface area (mm²) and cell counts per area.

In situ detection of apoptosis
A commercial assay (ApopTag In Situ Apoptosis Detection Kit) (Oncor, Gaithersburg, MD) using terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick-end labeling (TUNEL) was used to detect nuclear DNA fragmentation indicative of apoptosis. Tissues were fixed and embedded in paraffin as described for histology, and 5-µm sections were collected onto silinated glass microscope slides. Sections were deparaffinized through a graded alcohol series into 0.1 mol/L phosphate buffered saline (PBS). Tissue sections were covered with 100 µg/mL Rnase A (Boehringer Mannheim, Laval, Quebec) in 2 x SSC buffer and incubated in a humidified chamber for 40 minutes at 37°C. Slides were washed three times in 2 x SSC buffer. Endogenous peroxidase was quenched in fresh 0.15% hydrogen peroxide in 0.1 mol/L PBS for 25 minutes at RT, rinsed in 0.1 mol/L PBS, and covered with prewarmed 37°C proteinase K (Boehringer Mannheim) (20 µg/mL in Tris-EDTA buffer, pH 8.0). Slides were incubated at 37°C for 1 minute and then on ice in chilled 2 mg/mL glycine in 0.1 mol/L PBS for 15 minutes. After 10 minutes in -20°C ethanol:acetic acid (2:1), slides were rinsed with 2 x SSC buffer and sections were covered with equilibration buffer (ApopTag Kit, Intergen, Purchase, NY) for 10 minutes. Sections were treated with TdT enzyme with digoxigenin-labeled dUTP (ApopTag Kit), as suggested by the supplier, in a humidified chamber at 37°C for 2 hours. The reaction was stopped by three 10-minute washes in 37°C 2 x SSC buffer. The sections were covered with peroxidase-labeled antidigoxigenin antibody with 0.1% Tween-20 for 1 hour at 37°C in a humidified chamber. Slides were washed in 0.1 mol/L PBS. Color development was achieved by exposing sections to freshly prepared 0.01% hydrogen peroxide using 0.06% 3.3' diaminobenzidine as the chromogen, for up to 20 minutes at room temperature. Sections were counterstained with methyl green, dehydrated, and mounted. Normal rat intestine was used as a positive control tissue. For negative controls, distilled water was substituted for the TdT enzyme.

Immunohistochemistry
Sections were deparaffinized. Endogenous peroxidase was quenched (0.06% hydrogen peroxide and methanol (HOOH/MeOH)), nonspecific staining blocked with normal equine serum, and subsequently incubated with primary mAb for 1 hour at 37°C. The antibodies used were anti-CD43 (W3/13, Cedarlane, Mississauga, Ontario) specific for a cell surface molecule present on most rat leukocytes, anti-CD3 (R73, Cedarlane) specific for ß T cell receptor –positive bearing cells, and anti-CD8 (MRC OX-8, Cedarlane) specific for a cell surface molecule present on CD8 + CTL. Sections were then washed in PBS containing 1% BSA (Sigma, Oakville, Ontario, Canada) three times before incubation with biotinylated secondary antibody and labeling with peroxidase avidin/biotin complex using 3.3' diaminobenzidine as the chromogen (Vector, Burlingame, CA).

Statistical analysis
Apoptotic cell counts and number of labeled leukocytes were expressed as a function of leaflet surface area (in square millimeters) and analyzed as continuous variables. Means were obtained from 4 animals in each group. Data are reported as mean and standard error of the mean. An unpaired two-tailed Student’s t test was used to assess statistical significance, with a p value of less than 0.05 being the limit of significance.

	Results

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
Transplanted allografts were harvested and evaluated by light microscopy for stuctural integrity of leaflets at 7, 14, and 28 days after implantation in a syngeneic (Lewis to Lewis) or an allogeneic (BN to Lewis) environment. A total of 36 animals received aortic valve transplants (4 animals/group). A representative control aortic graft at low magnification is illustrated in Figure 1. By serial cross-sectional preparation the leaflets were easily identified. Leaflets were visualized in all syngeneic transplants for the entire duration of this study (28 days). In contrast, allogeneic implants were gradually destroyed such that, at 28 days, only 50% of the animals had an identifiable remnant of leaflet tissue. Previous work in our laboratory has demonstrated the complete absence of leaflets in allogeneic transplants 56 days after implantation [15].

View larger version (148K): [in this window] [in a new window]   Fig 1. Photomicrograph of control syngeneic (Lewis-Lewis) valve graft implant 14 days after implantation. Cross-section of aortic root showing three thin cellular leaflets representing part of three cusps of a normal aortic valve (x40) (arrow = leaflet).

View larger version (123K): [in this window] [in a new window]   Fig 2. Photomicrographs of syngeneic (Lewis-Lewis) valve grafts at (A) 7 days, (B) 14 days, and (C) 28 days after implantation. Photomicrographs of allogeneic (BN-Lewis) valve grafts at (D) 7 days, (E) 14 days, and (F) 28 days after implantation (x200) (arrow = leaflet; ALLO = allogeneic; SYN = syngeneic).

View larger version (105K): [in this window] [in a new window]   Fig 3. Photomicrographs of immunocytochemical staining at 7 days after implantation in syngeneic (Lewis-Lewis) (A) anti-CD43, (B) anti-CD3, (C) anti-CD8 and allogeneic (BN-Lewis), (D) anti-CD43, (E) anti-CD3, and (F) anti-CD8 with a secondary immunoperoxidase reaction (x200) (arrow = leaflet; ALLO = allogeneic; SYN = syngeneic).

View larger version (126K): [in this window] [in a new window]   Fig 4. Photomicrographs of immunocytochemical staining at 14 days after implantation in syngeneic (Lewis-Lewis) (A) anti-CD43, (B) anti-CD3, (C) anti-CD8 and allogeneic (BN-Lewis), (D) anti-CD43, (E) anti-CD3, and (F) anti-CD8 with a secondary immunoperoxidase reaction (x200) (arrow = leaflet; ALLO = allogeneic; SYN = syngeneic).

View larger version (19K): [in this window] [in a new window]   Fig 5. Graph comparing the number of immunolabeled cells in syngeneic (Lewis-Lewis) as compared to allogeneic (BN-Lewis) leaflets 7 days after implantation using anti-CD43, anti-CD3, and anti-CD8 mAb and secondary immunoperoxidase reaction. Counts are expressed as number of cells per square millimeter of leaflet area (4 animals/group).

View larger version (14K): [in this window] [in a new window]   Fig 6. Histogram comparing the number of apoptotic cells (in situ TUNEL assay) in leaflets from syngeneic (Lewis-Lewis) and allogeneic (BN-Lewis) transplants. Counts are expressed as number of cells per square millimeter of leaflet area (4 animals/group; p = 0.006).

View larger version (109K): [in this window] [in a new window]   Fig 7. Photomicrographs of BN valve grafts implanted into nude (rnu/rnu) animals and evaluated at 7 days (A),14 days (B), and 28 days (C) after implantation (x200). Photomicrographs of immunocytochemical staining with anti-CD3 mAb and anti-CD8 mAb and evaluated at 7 days (D and G), 14 days (E and H), and 28 days (F and I) after implantation in nude recipients with a secondary immunoperoxidase reaction (x200). (arrow = leaflet).

	Comment

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

We have shown that, as early as 7 days after implantation, there is extensive lymphocytic infiltration into allogeneic leaflets. These infiltrates are predominantly T lymphocytes (CD3+) with a majority being CD8+ cells, suggesting active CTL. This indicates that, by 7 days posttransplant, immune activation has occurred and that there has been transit of alloreactive CTL to the valve graft. Moreover, these CTL have made their way into the leaflet tissue where they will come into contact with the allogeneic stromal cells. This result extends the work of others who have shown systemic sensitization of lymphocytes by aortic valve grafts in both rodents and humans [11, 12]. In this study this systemic sensitization is demonstrated to be related to local T cell effector populations within the leaflets. Recent work by O’Brien and coworkers supports the concept of immune activation targeted to the allograft [5]. However, their study did not extend to events in the leaflet tissue but were restricted mostly to the arterial conduit, where they confirmed the characteristics of chronic rejection that have been well described in aortic interposition graft models [18, 19].

The infiltration of T cells and, in particular, of CD8+ CTL into the valve leaflets provides strong circumstantial evidence that valve destruction is the result of a T cell–mediated immune response. We have confirmed this suggestion by demonstrating that, in the absence of a functional T cell compartment (in nude rats), cellular infiltration and valve destruction do not occur. This both confirms that the overall response is T-cell orchestrated and provides strong evidence to support the suggestion that the infiltration is causally related to the valve destruction. This hypothesis is further supported by our demonstration that apoptosis is ongoing in the allogeneic valve leaflets during the time of maximal CD8+ T-cell infiltration.

This study suggests that valve destruction is mediated by T-cell activation and the generation of alloreactive CTL that eliminate leaflet stromal cells by apoptotic mechanisms, which is the primary mode of destruction used by CTL. In support of this hypothesis, the gradual loss of cellular elements in allografted valves has been previously suggested to occur by apoptotic cell death [20]. Further examination of the presence of CTL mediators of apoptosis such as granzyme B, perforin, and Fas ligand will be required to confirm this hypothesis. In addition, techniques need to be devised to differentiate between stromal cell apoptosis and the inflammatory cell apoptosis that will certainly be occuring coincident with valve destruction.

Given that allograft valves are not matched for human leukocyte antigen type and that no immunosuppressive regimen is used, it would seem logical that these valves would incite a harmful immune response, particularly with the mounting evidence that aortic allograft valves are antigenic [7–11]. It is therefore surprising that very little effort has been made to understand the sequence of events that occurs after implantation, and whether modification of the host response could extend the longevity of the valves. Standard clinical practice does not attempt to minimize or to control such a response. Some clinicians have attempted to match valves for ABO blood group compatibility; however, there is no evidence that this is beneficial [21]. Immunosuppressive or antiinflammatory agents have been sporadically tried but no rigorous, prospective study has been performed [6, 22]. The risk–benefit ratio of a permanent immunosuppressive regimen versus reoperation for allograft failure may indeed be too high. However, it is possible that these nonvascularized allograft valves may require only short-course immunosuppressive treatment after transplantation to limit immune-mediated injury. To date, very few efforts have been made to evaluate the impact of immunosuppressive treatment on the structural integrity of allograft valves. In a rat model, cyclosporine treatment was shown to result in the preservation of cellular elements after allotransplantation, but little reference was made to the structural integrity of the valves [23].

In summary, aortic valve allograft failure in a heterotopic rat model is a T cell–mediated rejection event that appears to involve alloreactive cytotoxic T lymphocytes. Taken together, these study findings provide a starting point for the development of novel therapies for allograft valve failure. Given the nature of the graft it is likely that these therapies will not be identical to current therapies for solid organ allografts, but may have conceptual similarities.

	Acknowledgments

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 
This work was supported by The Toronto Hospital for Sick Children Research Foundation.

	References

Top Abstract Introduction Material and methods Results Comment Acknowledgments References

 

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This Article Abstract Full Text (PDF) Alert me when this article is cited Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Add to Personal Folders Download to citation manager Author home page(s): David B. Ross Permission Requests Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Legare, J. F. Articles by Ross, D. B. Search for Related Content PubMed PubMed Citation Articles by Legare, J. F. Articles by Ross, D. B.