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(Arteriosclerosis, Thrombosis, and Vascular Biology. 2005;25:1347.)
© 2005 American Heart Association, Inc.

Vascular Biology

High Prevalence of Circulating CD4⁺CD28^– T-Cells in Patients With Small Abdominal Aortic Aneurysms

Christina Duftner; Rüdiger Seiler; Peter Klein-Weigel; Heike Göbel; Christian Goldberger; Christian Ihling; Gustav Fraedrich; Michael Schirmer

From the Departments of Internal Medicine (C.D., C.G., M.S.) and Vascular Surgery (R.S., P.K.-W., G.F.), Innsbruck Medical University, Innsbruck, Austria; and the Department of Pathology (H.G., C.I.), University Hospital, Freiburg, Germany.

Correspondence to Michael Schirmer, MD, Assoc Prof, Clinical Department of Internal Medicine, Clinical Division of General Internal Medicine, Innsbruck Medical University, Anichstrasse 35, 6020 Innsbruck, Austria. E-mail michael.schirmer{at}uibk.ac.at

	Abstract

Top Abstract Introduction Materials and Methods Results Discussion References

 
Objective— To assess the possible role of proinflammatory CD28^– T cells in abdominal aortic aneurysms (AAAs). Animal studies and human tissue studies suggest a role for interferon (IFN)-–producing T cells in the development and progression of AAAs.

Methods and Results— Fluorescence-activated cells sorter analysis of peripheral blood samples and measurement of AAA size using sonography were performed in 101 AAA patients and 38 healthy controls. Peripheral percentages of CD28^– T cells of the CD3⁺CD4⁺ and the CD3⁺CD8⁺ were enriched in AAA patients with 7.8±8.8% and 41.9±15.7% compared with healthy controls with 2.2±6.1% and 24.9±15.5%, respectively (P=0.002 and P<0.001, respectively). Both CD4⁺CD28^– and CD8⁺CD28^– T cells produced large amounts of IFN- and perforin. Patients with small AAAs (<4 cm) showed higher peripheral levels of CD4⁺CD28^– T cells than those with larger AAAs (P=0.025). Immunohistological examinations revealed 39.1±17.2% CD4⁺CD28^– and 44.0±13.8% CD8⁺CD28^– in AAA tissue specimens with inflammatory infiltrates.

Conclusions— IFN-– and perforin-producing CD28^– T cells are present in the periphery and the vessel wall of a majority of AAAs. This observation in humans favors the concept of a T cell–mediated pathophysiology of AAAs, especially during the early development of AAAs.

Animal studies and tissue studies suggest a role for interferon (IFN)-–producing T cells in the development of abdominal aortic aneurysms (AAAs). This study shows an increased prevalence of circulating IFN-–producing CD28^– T cells especially in smaller AAAs, and thus supports the concept of a T cell–mediated pathophysiology of AAAs.

Key Words: aortic disease • aneurysms • leukocytes • immune system • human

	Introduction

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Abdominal aortic aneurysm (AAA) is a common disease with a prevalence of 3% of individuals aged 60 years and older and is a potentially lethal disorder causing 15 000 deaths annually in the United States.¹

Recent human tissue studies and animal models have led now to a paradigm shift in the pathogenetic concept of AAAs. Rather than a simple degenerative process, the majority of AAAs has proven to be a complex and dynamic remodeling process. In brief, studies of human AAA tissues have identified extensive inflammatory infiltrates in both the media and the adventitia,² leading to an increased expression of proinflammatory cytokines and C-reactive protein (CRP) in aneurysmal tissue.^3–5 The presence of vascular-associated lymphoid tissue (VALT) with lymphoid follicles and lymph node–like structures in the adventitia of AAAs suggests a role for immunocompetent and antigen presenting cells not only in atherosclerosis⁶ but also in AAA disease.² The role for T cells has been further supported by immunogenetic findings with associations between AAA and human leukocyte antigen class II molecules.⁷ Infiltration of the adventitia usually occurs together with medial thinning. This inflammatory process triggers the production of metalloproteinases and apoptosis of medial smooth muscle cells thus explaining the disruption of the orderly lamellar structure in aortic aneurysms.

Recently, a subgroup of proinflammatory T cells has been identified in patients with immune-mediated disorders including rheumatoid arthritis,^8–9 ankylosing spondylitis,¹⁰ multiple sclerosis,¹¹ Wegener granulomatosis,¹² and unstable angina,¹³ which lack the costimulatory molecule CD28 on their surface and are considered as markers for chronic inflammation and early aging.¹⁴ Under these circumstances the CD28^– T cells are part of the CD4⁺ as well as the CD8⁺ T cell compartment, persist over years, and include most of the oligoclonally expanded T cells. Phenotypically, CD4⁺CD28^– T cells from rheumatoid arthritis and ankylosing spondylitis patients and CD8⁺CD28^– T cells from aged persons, rheumatoid arthritis, and melanoma patients share the expression of various natural killer (NK) cell receptors and lack the expression of the lymphocyte marker CD7.^9,15–16 Functionally these T cells are capable to release large amounts of interferon (IFN)-, perforin, and granzyme B, providing them with the possibility to lyse target cells.¹⁷ In the pathogenesis of coronary arteriosclerosis, the critical role of IFN- was already investigated in a mouse heart transplant model, and IFN- knockout recipients did not develop thickening of the arterial intima despite a T lymphocyte and macrophage infiltrate in the parenchyma.¹⁸

Therefore we examined in this study whether proinflammatory IFN-–producing CD4⁺CD28^– and CD8⁺CD28^– T cells would be enriched in patients with AAAs peripherally and locally. Additionally, the characteristics of these cells and a possible association between the percentages of CD28^– T cells and AAA size were studied.

	Materials and Methods

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Patients and Ethical Concerns
In a prospective design, all consecutive patients older than 50 and younger than 80 years of age with a diameter of the abdominal aorta larger than 3 cm were considered for recruitment into the study. Patients and controls with a history of neoplasm, recent acute infection, or history of any other immune-mediated chronic disease possibly related to elevated peripheral levels of CD28^– T cells were excluded from the study. In 49 consecutive AAA patients, a second blood draw was performed during follow-up. Maximal diameters of AAA were measured by sonography. Sonography was not performed in the controls. In our cohort 26.6% of AAA patients were treated by aneurysmectomy. The protocol was approved by the local ethics committee of the Medical Faculty of Innsbruck, Austria.

Immunostaining and Flow Cytometry
Surface staining of peripheral blood mononuclear cells (PBMCs) was performed using fluorescein isothiocyanate (FITC)-conjugated anti-CD4, anti-CD8, anti-CD7, anti-CD57, anti-CD45RO, anti-CD45RA, phycoerythrin-conjugated anti-CD28, and peridinin chlorophyll protein-conjugated anti-CD3, anti-CD4, or anti-CD8 monoclonal antibodies (all from Becton Dickinson, San Diego, Calif). For detection of apoptotic cells, cells were stained with Annexin V (Becton Dickinson). For intracellular staining of IFN- and perforin, stimulated cells were stained with FITC-conjugated anti–IFN-, anti-perforin, and control immunoglobulin, respectively (Becton Dickinson). For more information please see the expanded Methods (available online at http://atvb.ahajournals.org).

Immunohistochemistry
Serial sections were stained for CD4 (mouse anti-human monoclonal antibody; DAKO Cytomation, Denmark), CD8 (mouse anti-human monoclonal antibody; DAKO Cytomation), and CD28 (goat anti-human polyclonal antibody; R&D Systems Inc, Minneapolis, Minn) according to standard protocols using a three step avidin-biotin complex method.¹⁹ In brief, after fixation of the sections in acetone, endogenous peroxidase was blocked. Nonspecific background was blocked by normal goat (DAKO Cytomation) or normal donkey serum (Jackson ImmunoResearch Laboratory Inc, West Grove, Penn). After application of the primary antibody, the slides were incubated with the respective biotinylated secondary antibodies and eventually incubated with a horseradish-peroxidase (HRP)-labeled avidin–biotin complex (SABC; DAKO Cytomation). The enzyme was detected by 3-amino-9-ethyl-carbazole (AEC; Sigma) yielding a brownish-reddish color. Sections were counter-stained with hematoxylin. For double staining, donkey anti-mouse bound alkaline phosphatase was developed with fast blue BB salt (Sigma) and HRP with AEC. For more details please see the expanded Methods.

Statistics
Univariate variance analysis with adjustment for differences in age and sex between healthy controls and AAA patients, the two-sided Spearman-Rho, the Kruskal-Wallis, the paired t test, and regression analysis by receiver operating curves were performed as indicated using the SPSS program, version 11.0. For further details please see the expanded Methods.

	Results

Top Abstract Introduction Materials and Methods Results Discussion References

 
A total of 101 AAA patients (69.4±7.4 years old) and 38 healthy controls (60.7±8.9 years old) without a history of neoplasm, recent acute infection, or history of any other immune-mediated chronic disease were included into the study for peripheral fluorescence-activated cells sorter (FACS) analyses. In 90 patients diameters of AAAs were measured sonographically to be 5.0±1.2 cm (range, 3.0 to 8.7). Levels of the CRP were increased to 2.8±4.8 mg/dL (n=85; range, 0.2 to 30.2, normal levels <0.7 mg/dL) and the erythrocyte sedimentation rate (ESR) to 20.4±23.2 mm/h (n=38, range, 2.0 to 95.0, normal controls <15 mm/h). Patients’ characteristics are summarized in Table I (available online at http://atvb.ahajournals.org). Out of our cohort of AAA patients, 30.3% of patients had a history of coronary heart disease and 23.2% of patients a history of peripheral arterial occlusive disease.

Prevalence of Circulating CD4⁺CD28^– and CD8⁺CD28^– T-Cells in AAA Patients
In the peripheral blood, percentages of CD28^– of the CD3⁺CD4⁺ and the CD3⁺CD8⁺ cells were enriched to 7.8±8.8% and 41.9±15.7% in AAA patients compared with healthy controls with 2.2±6.1% and 24.9±15.5%, respectively (P=0.002 and P0.001, respectively, after adjustment for age and sex; Figure 1A). 95% of the age-matched healthy controls had CD3⁺CD4⁺CD28^– T cells lower than 3.5% and CD3⁺CD8⁺CD28^– T cells lower than 54.7%. Taking these levels as limits for normal accumulation of CD28^– T cells, 60.4% of the AAA patients had elevated levels of CD4⁺CD28^– T cells and 19.8% showed increased levels of CD8⁺CD28^– T cells in the peripheral blood. Receiver operating curves (ROC) as a regression analysis model revealed comparable specificity and sensitivity of CD4⁺CD28^– and CD8⁺CD28^– T cells for AAA disease, with a calculated area under the curve (AUC) of 0.795 and 0.799, respectively. Using the two-sided Spearman-Rho test to analyze a possible dependency of circulating levels of CD4⁺CD28^– and CD8⁺CD28^– T cells, a correlation was found between these two T cell subsets (R=0.530, P<0.001).

View larger version (59K): [in this window] [in a new window]   Figure 1. Peripheral and local presence of CD4+CD28– and CD8+CD28– T cells in AAA disease. A, Peripheral circulating levels of CD28– T cells out of the CD3+CD4+ and CD3+CD8+ T cell compartments are increased in AAA patients (•; 7.8±8.8% and 41.9±15.7%) compared with healthy controls (; 2.2±6.1% and 24.9±15.5%; P=0.002 and P0.001). The mean is shown as a continuos and dotted line, respectively. Age- and sex-adjusted univariate variance analysis was used to compare peripheral levels of CD4+ and CD8+CD28– T cells in controls and AAA patients. P<0.01 is considered as highly significant. B, Hematoxylin-Eosin staining of a transmural section of AAAs reveals lymphocytic infiltration within the outer part of the media and the adventitia (red circle) and an advanced plaque of the intima (100-fold magnification). C, Immunohistological examinations reveal 39.1±17.2% CD4+ T cells lacking the costimulatory signal CD28 and 44.0±13.8% CD8+CD28– T cells within lymphocytic infiltrations in the adventitia. T cells with CD4+ or CD8+ positivity but without staining for CD28 are shown in blue (black arrows) and double-stained CD4+/CD8+ and CD28+ T cells are shown in dark blue/violet (white arrows, 400-fold magnification).

The increase of peripheral levels of CD4⁺ and CD8⁺CD28^– T cells was independent from coexisting peripheral arterial occlusive disease (5.9±6.6% and 41.8±17.3%, respectively) or coronary heart disease (9.0±8.6% and 44.6±17.1%) compared with patients with AAAs alone (7.5±9.2% and 39.7±13.9%, respectively).

Follow-Up and Apoptosis of Peripheral CD4⁺ and CD8⁺CD28^– T-Cells
During follow-up of 14.5±10.5 months, the levels of CD3⁺CD4⁺CD28^– T cells increased from 7.8±8.8% to 11.1±9.2% and levels of CD3⁺CD8⁺CD28^– T cells from 41.9±15.7% to 46.4±16.6% (n=49, P=0.006 and P=0.082, respectively). After staining with Annexin V, both CD4⁺ and CD8⁺CD28^– T cells showed reduced spontaneous apoptosis compared with their CD28⁺ T cell counterparts (3.3±1.8% and 4.2±2.7% to 19.2±9.2% and 17.7±9.9% Annexin V⁺ cells, P=0.007 and P=0.012, respectively).

Phenotypic and Functional Characterization of CD4⁺ and CD8⁺CD28^– T-Cells
For phenotypic characterization of CD28^– T cells, surface expression of CD7, CD57, CD45RA, and CD45RO were compared between CD28^– T cells and their CD28⁺ T cell counterparts (Figure 2A through 2H). The CD7 molecule, which is involved in T cell activation, is present on most normal human T cells under physiological conditions, but not on NK cells.²⁰ CD7 surface expression was low on CD4⁺CD28^– T cells in comparison with CD4⁺CD28⁺ T cells (4.7±3.6% versus 52.6±18.0% positive cells, P=0.001; Figure 2A), whereas CD7 was less expressed on CD8⁺CD28^– T cells compared with their CD8⁺CD28⁺ T cell counterparts (67.2±19.1% versus 81.9±10.6% positive cells, P=0.020; Figure 2C). CD57 is a 110-kDa glycoprotein that is typically presented by NK cells. CD57 expression was increased on CD4⁺CD28^– T cells compared with CD4⁺CD28⁺ T cells (67.2±23.2% versus 2.5±3.2% positive cells, P=0.001; Figure 2B). CD57 was expressed on 70.7±10.7% of CD8⁺CD28^– T cells, compared with 4.2±1.8% of CD8⁺CD28⁺ T cells (P<0.001; Figure 2D). CD45RA, the marker for naïve T cells, was expressed on 53.4±26.4% of CD4⁺CD28^– T cells and on 19.7±10.4% of CD4⁺CD28⁺ T cells (P=0.010; Figure 2E). On CD8⁺CD28^– T cells, surface expression of CD45RA was 72.7±14.8% compared with 20.8±16.4% on CD8⁺CD28⁺ T cells (P<0.001; Figure 2G). CD45 RO, the marker for memory T cells, was expressed on 42.4±24.1% of CD4⁺CD28^– T cells and 72.6±10.9% of CD4⁺CD28⁺ T cells (P=0.009; Figure 2F). On CD8⁺CD28^– T cells, surface expression of CD45RO was 10.9±10.8% compared with 59.2±17.3% on CD8⁺CD28⁺ T cells (P=0.001; Figure 2H). Representative FACS plots are shown as an example for phenotypic expression of CD7, CD57, CD45RA and CD45RO in Figure I (available online at http://atvb.ahajournals.org).

View larger version (21K): [in this window] [in a new window]   Figure 2. Phenotypic characterization of CD4+CD28– and CD8+CD28– T cells by triple-color FACS analysis for CD7 (A) and CD57 (B) on CD4+ T cells, CD7 (C) and CD57 (D) on CD8+ T cells, CD45RA (E) and CD45RO (F) on CD4+ T cells, and CD45RA (G) and CD45RO (H) on CD8+ T cells. Data are shown as mean±SD. Probability values were calculated using the paired t test to compare the CD28+ with the CD28– compartment with P<0.05 considered as significant and P<0.01 as highly significant.

From the functional perspective, production of IFN- was more frequent in CD4⁺CD28^– than in CD4⁺CD28⁺ T cells (34.5±20.0% versus 14.6±8.1% IFN-–producing cells, P=0.003; Figure 3A) and more frequent in CD8⁺CD28^– than in CD8⁺CD28⁺ T cells (32.3±41.8% versus 14.5±25.1% IFN-–producing cells, P>0.05; Figure 3D). Interleukin (IL)-4–producing cells were low in both CD4⁺CD28^– and CD4⁺CD28⁺ T cells (1.9±1.5% versus 1.2±1.0% IL-4–producing cells, P>0.05; Figure 3B) and in CD8⁺CD28^– and CD8⁺CD28⁺ T cells (2.9±2.0% versus 1.8±1.6% IL-4–producing cells, P>0.05; Figure 3E). Perforin-producing cells, however, were more frequent in CD4⁺CD28^– than in CD4⁺CD28⁺ T cells (38.6±21.3% versus 1.6±1.1% perforin-positive cells, P=0.004; Figure 3C) and more frequent in CD8⁺CD28^– than in CD8⁺CD28⁺ T cell subsets (49.0±26.4% versus 4.6±4.4% perforin-positive cells, P=0.043; Figure 3F).

View larger version (17K): [in this window] [in a new window]   Figure 3. Functional characterization of CD4+CD28– and CD8+CD28– T cells by intracellular cytokine production. Production of IFN- is shown in stimulated CD4+ T cells (A) and CD8+ T cells (D), production of IL-4 in CD4+ T cells (B) and CD8+ T cells (E), and production of perforin in CD4+ T cells (C) and CD8+ T cells (F). Data are shown as mean±SD. Probability values were calculated using the paired t test to compare the CD28+ with the CD28– compartment with P<0.05 considered as significant and P<0.01 as highly significant.

Aneurysm Dimension and CD4⁺ and CD8⁺CD28^– T-Cells
To test a possible association between the peripheral levels of CD4⁺ and CD8⁺CD28^– T cells and the size of AAAs, maximal AAA diameters were subdivided into 3 groups: small (<4 cm, n=17), intermediate (4 to 6 cm, n=56), and large AAAs (>6 cm, n=17). Patients with small AAAs (<4 cm) showed higher peripheral levels of CD4⁺CD28^– T cells (11.7±9.0%) than those patients with intermediate (6.6±7.7%, P=0.025) and large AAAs (6.7±5.6%, P=0.065; Figure 4A). No correlation was detected between the peripheral levels of CD8⁺CD28^– T cells and the maximal AAA diameter (Figure 4B). Levels of CD4⁺ and CD8⁺CD28^– T cells did not correlate with age, CRP, or ESR.

View larger version (14K): [in this window] [in a new window]   Figure 4. Aneurysm dimension (diameter measured by sonography) and peripheral levels of CD4+ and CD8+CD28– T cells. A, Patients with small AAAs (<4 cm) show highest peripheral levels of CD4+CD28– T cells. B, Peripheral levels of CD8+CD28– T cells do not depend on AAA diameter. Whiskers box plots show 50% of cases within the boxes and all data excluding mavericks between the end points of the whiskers (lines). Probability values were calculated using the Kruskal–Wallis test to compare small, medium, and large-sized AAAs, with P<0.05 considered as significant.

CD4⁺ and CD8⁺CD28^– T-Cells in AAA Tissue Sections
For immunohistological studies, single and double staining of CD4, CD8, and CD28 were performed to examine the local presence of these CD4⁺ and CD8⁺CD28^– T cells in cryo-frozen tissue specimens from AAA patients who underwent surgery. Of 13 specimens from individual AAA patients, 7 specimens showed inflammatory infiltrations with lymphocytes predominantly in the outer part of the media and the adventitia (Figure 1B) and were used for further double staining evaluations. In the other specimens the media consisted of scar tissue with calcifications or missing adventitial tissue and immunohistochemistry showing only few scattered inflammatory cells and single smooth muscle cells. The ratio between CD4⁺ and CD8⁺ T cells within the lymphocytic aggregates was calculated to be 3.9±2.7. In our tissue specimens with inflammatory infiltrates, the levels of CD4⁺CD28^– and CD8⁺CD28^– T cells were 39.1±17.2% and 44.0±13.8%, respectively (Figure 1C). Although not significant, CD4⁺CD28^– T cells were more frequent in the tissue (23.4±26.1%) than the peripheral blood (8.9±14.1%), whereas CD8⁺CD28^– T cells appeared less frequent in tissue specimens (23.4±25.1%) than in the peripheral blood (44.0±15.0%, NS).

	Discussion

Top Abstract Introduction Materials and Methods Results Discussion References

 
The present study shows for the first time that peripheral percentages of CD28^– of the CD3⁺CD4⁺ and the CD3⁺CD8⁺ T cells are expanded in the peripheral blood of AAA patients even without history or clinical signs of any other immune-mediated diseases. This observation further supports the concept of chronic inflammation including T cells in the pathophysiology of AAAs, and further underlines the systemic nature of many AAAs. Both CD4⁺ and CD8⁺CD28^– T cells in the peripheral blood showed reduced spontaneous apoptosis compared with their normal CD28⁺ counterpart and persisted in the peripheral blood over months. This observation could explain the perpetuation of AAA disease after an initial trigger as proposed for rheumatoid arthritis.²¹

The crucial role for IFN-–producing T cells in the development of AAAs has been shown in a recent animal study with CD4^–/– and IFN-^–/– knock-out mice demonstrating the necessity of IFN- or IFN-–producing lymphocytes for the development of AAAs.²² Also in the development of atherosclerosis IFN- was shown to play an essential role in animal models, and immunofluorescent studies of human carotid specimens revealed activated T cells as a possible source of IFN- in the atherosclerotic plaques.^23–25 Indeed, human CD28^– T cells produce IFN- which then activates macrophages to produce proinflammatory cytokines and results in oxidative damage with lipid peroxidation attacking smooth muscle cells and matrix components of the vessel wall. Thus the disease process is perpetuated and cytotoxic features result in thinning of the arterial wall and aneurysm formation. Besides, both CD4⁺ and CD8⁺CD28^– T cells release large amounts of perforin, and thus have to be considered as cytotoxic. Cytotoxic T cells are suspected to participate in tissue injury, and in vitro experiments already showed effective killing of endothelial cells by cytotoxic CD4⁺CD28^– T cells.¹⁷ Recently a Th2-predominant inflammation and lack of IFN- signaling in the host were reported to induce murine aneurysms in allografted aortas.²⁶ The IFN- receptor knockout mice show increased levels of IL-4 with markedly increased levels of matrix metalloproteinase (MMP)-9 and MMP-12. We wonder whether the concomitant development of a Th1 response as indicated by the prominent increase in IFN- expression might still have an effect on cells within the aortic graft and thus could trigger aneurysm formation.²⁷ In our experiments both CD4⁺CD28^– and CD8⁺CD28^– T cells lack the intracellular production of IL-4 analyzed by FACS after stimulation (Figure 3B and 3E).

Indeed we found that these specific T cells were higher in the smaller AAAs (<4 cm) than in larger AAAs (4 cm). The correlation between CD28^– T cells and the size of the AAAs parallels the findings by Hamano et al that tumor necrosis factor alpha (TNF-) and macrophages are more present in small-sized AAAs than in large-sized AAAs.⁵ Taken together it appears that in the development of AAA the immune-mediated process occurs earlier in the disease with involvement of proinflammatory T cells, whereas the larger AAAs represent more an advanced stage with a minor role of immune-mediated processes. Thus in the early phase of AAA development the risk of AAA rupture is low,^28–29 but the T cell–mediated process leads to macrophage activation with TNF- production and subsequent elastin and collagen degradation and thus precedes the crucial changes of the AAA wall with potential rupture of the AAA in the later phase.

An increased prevalence of CD28^– T cells had been proposed for several diseases including unstable angina and vasculitic diseases.^12–13 Therefore we had excluded patients with such diseases, and had tested for possible correlation between CD4⁺CD28^– T cells and other atherosclerotic diseases including peripheral arterial occlusive disease and coronary heart disease in the AAA patients. As coexisting atherosclerotic diseases had no influence on the levels of the CD28^– T cells in AAA patients, the relationship between CD4⁺CD28^– T cells and adventitial inflammation in the AAAs becomes even more obvious. This finding is also in line with the report that CD4⁺CD28^– T cells had been shown to be lower in stable coronary heart disease than in unstable disease.¹³ The higher levels of CD4⁺CD28^– T cells in smaller AAAs with more macrophages and higher levels of TNF- compared with larger AAAs further support the concept of CD4⁺CD28^– T cells correlating with AAA disease activity more than with atherosclerotic disease. As shown by others, exposure to TNF- results in downregulation of CD28 in vitro, which may explain the higher prevalences of CD4⁺CD28^– T cells in smaller AAAs.³⁰ At present we can only speculate about a possible role of IL-12, which can actively reverse downregulation of CD28 in vitro.³¹ The levels of IL-12 in larger AAAs have not been investigated so far.

The definite antigen recognized by the CD28^– T cells remains unclear. Several studies have been performed to define possible triggers of AAAs, but results are still not conclusive.^32–34 T lymphocyte expression of isoforms of CD45 corresponds with their ability to respond to recall antigens. The naïve T cell pool is defined by the marker CD45RA, whereas the responsive or memory T cell pool is defined by CD45RO.^35–36 More recently, however, it has been shown that the CD45RA⁺ T cell population is heterogeneous, as CD45RO⁺ T cells can reconvert back to CD45RA⁺ T cells.³⁷ In our AAA patients, both CD4⁺ and CD8⁺CD28^– T cells revealed high percentages of CD45RA on the cell surface, which confirms their status as highly differentiated cells. Up to now, there is ongoing discussion regarding whether CD28^– T cells are caused by chronic antigen stimulation or whether their characteristic phenotypic and functional properties is a result of premature senescence.³⁸

In the peripheral blood the levels of CD4⁺CD28^– T cells correlated well with the levels of CD8⁺CD28^– T cells. In the AAA tissue, however, CD4⁺CD28^– T cells seemed to be enriched, whereas CD8⁺CD28^– T cells appeared less frequent in the tissue specimens. Both cell types were stained and analyzed by the same technique using a computer assisted cell counting program. Therefore the CD4⁺CD28^– T cells may be more important for the pathogenetic mechanisms of AAA than the CD8⁺CD28^– T cells.

Our study has the limitation that AAA specimens were only available from those patients who underwent aneurysmectomy because of a large-sized AAA. Because CD28^– T cells are less frequent in the large-sized AAAs, the immunohistological findings in this late phase of AAA may underestimate the number of immune-mediated AAAs. This could be explained by the fact that our surgeons take tissue specimens from AAAs from the mid-portion of the AAA, although more TNF- mRNA and TNF-–converting enzyme (TACE) mRNA had been described in the transition zone of AAAs.³⁹ Thus the mid-portion of AAAs reflects more a non- or postinflammatory status, whereas a more active inflammatory status would be expected in the transition zone of AAAs. The source of biopsies could thus explain the lack of T cells in some of our specimens taken from the mid-portion of the AAAs.

In conclusion, these human data support a role for IFN-–producing cytotoxic CD28^– T cells in the pathogenesis of a majority of AAAs as proposed by the CD4^–/– and IFN-^–/– knock-out mouse model. T cell–mediated processes appear to be more important for initiation and initial development of AAA disease (<4 cm) but may lose importance in the larger AAAs after wall destabilization and disintegration.

	Acknowledgments

 
The study was supported by the Research Fund of the Austrian National Bank (P 8835 and P 9715), Vienna, the Medical Research Fund of the Innsbruck Medical University (MFF), and the "Verein zur Förderung der Hämatologie, Onkologie und Immunologie," Innsbruck, Austria.

	Footnotes

 
C.D. and R.S. contributed equally to this work and have to be regarded as co-first authors.

Received November 12, 2004; accepted April 8, 2005.

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