Carátula del artículo
Relationships Between Phenotype and Function of Blood CD4+
T-Cells and Ascending Thoracic Aortic Aneurysm: an Experimental
Study
Silverio Sbrana
CNR Institute of Clinical
Physiology, Italy
Kaushal Kishore Tiwari drkaushalkt@yahoo.com
"G. Monasterio" Foundation, Italy
Scuola Superiore S. Anna, Italy
Teaching Hospital, Nepal
Stefano Bevilacqua
"G. Monasterio" Foundation, Italy
Paola Giungato
CNR Institute of Biomedical
Technologies, Italy
Enkel Kallushi
"G. Monasterio" Foundation, Italy
Marco Solinas
"G. Monasterio" Foundation, Italy
Anna Maria Mazzone
"G. Monasterio" Foundation, Italy
Brazilian Journal of Cardiovascular Surgery, vol. 34, no. 1, pp. 8-16, 2019
Sociedade Brasileira de Cirurgia Cardiovascular
Received: 01 October 2018
Accepted: 02 October 2018
Funding
Funding source: National Research Council (CNR) of Italy
Funding statement: Financial support: This study was funded by the National Research Council (CNR) of Italy, RSTL (Curiosity Driven Research Program)
DG.RSTL.035.006-035.
Table.
Abbreviations, acronyms symbols
INTRODUCTION
The non-familial ascending thoracic aortic dilatations and thoracic aortic aneurysms
(TAA) are frequent in individuals older than 65 years of age (approximately 6-9%),
with a risk of rupture or dissection ranging from 2 to 3.5 cases per 100,000
patients/year[1].
The progression from aortic dilatation to aneurysm is a multifactorial process
partially undiscovered. Besides other well-known mechanisms[2], the role of chronic
immune-mediated inflammations in defining biomechanical properties of the aortic
wall is still to be determined. In particular, the study of cellular and molecular
mechanisms leading to aortic fibrosis, considered the histopathological marker of an
altered vascular remodeling process[3], might be an important target in understanding the
individual's susceptibility to non-syndromic ascending thoracic aortic dilatation
and TAA formation. Recent findings in humans confirmed the key role of
myofibroblasts (MF) in the extracellular matrix (ECM) proteolysis and deposition
(fibrosis)[4]. MF
activity is modulated by a wide array of pro- and anti-fibrotic cytokines and growth
factors released by mononuclear immune cells infiltrating the inflamed
tissue[5].
Especially, the cytokines produced by CD4+ T-lymphocytes play a causative
role in the initiation and progression of fibrosis associated with pathological
conditions such as systemic sclerosis, atherosclerosis, and the use of silicone
mammary implants[6].
Given the crucial role of distinct CD4+ T-lymphocyte subsets in normal
immune-regulation[7], the evaluation of their dynamic functional balance in terms
of cell frequency ratio, either expressing a particular surface phenotype or
producing selective signature cytokines, is an important tool to investigate whether
a clinical pathological condition and its outcome are associated with a particular
T-helper functional perturbation[8,9].
To our knowledge, so far, no studies have related phenotype and function of
peripheral blood CD4+ T-lymphocytes with the presence of TAA. Therefore,
we have evaluated the presence of possible relationships between this pathological
condition and: a) the expression of chemokine receptors and activation markers
(CCR5, CXCR3, CX3CR1, CD25) on total blood CD4+ T-cell and on the
pro-inflammatory/cytotoxic subset CD4+CD28-, known to be
involved in vascular inflammation[10]; b) the cytokine production (interferon gamma [IFN-g,
interleukin [IL]-17A, IL-21, IL-10) by in vitro stimulated
CD4+ T-cells. Moreover, given their both beneficial and harmful roles
in several clinical settings, including vascular injury[11], we quantified the circulating
fraction of the CD4+CD25highFoxP3+ naturally
occurring regulatory T-cells (Treg). On this basis, newly established phenotypic and
functional blood CD4+ T-lymphocyte ratios, that overcome the traditional
Th1/Th2 paradigm[12],
have been calculated and related to the presence of an aortic aneurysm. We also
measured the circulating levels of several cytokines (IFN-γ, IL-6, IL-10,
IL-17A, IL-23, transforming growth factor beta [TGF-β]) and chemokines
(RANTES, CX3CL1) known to influence CD4+ T-cells function and
migration[13-15].
METHODS
Patients and Blood Samples
We have enrolled 20 patients undergoing surgery for aortic valve disease (AVD).
TAA group (n=10) included patients undergoing aortic valve surgery with TAA
surgery and AVD group (n=10) included patients undergoing surgery only for AVD,
like stenosis or regurgitation, or both. Both groups underwent surgery by
aortotomy. All patients had an indication for aortic valve surgery, according to
current guidelines. TAA surgery was performed, if recommended by current
guidelines. Exclusion criteria were the presence of genetic disorders,
autoimmune and chronic inflammatory diseases, cancer, or hematological diseases.
We enrolled patients with normal, ectasic or entirely aneurysmatic ascending
aorta. Comorbidities, risk factors, and medical therapies were recorded, with
particular attention to statins and/or angiotensin-converting enzyme (ACE)
inhibitors, known to exert also immunomodulating effects[16-18]. Mean and peak aortic gradients were
measured by Doppler echocardiography and used to define the grade of stenosis.
Aortic valve lesions were classified as predominant stenosis (mean transvalvular
gradient ≥40 mmHg and grade of regurgitation
<3+/4+), predominant regurgitation (grade of
regurgitation 4+/4+ and mean gradient <40 mmHg), and
steno-regurgitation, in other cases.
Venous blood was collected at the admission in Vacutainer tubes containing
tripotassium ethylenediaminetetraacetic acid (K3-EDTA) or
sodium-heparin as anticoagulants. Plasma was obtained after blood centrifugation
and immediately frozen at -80°C. The protocol was approved by the local ethics
committee and informed consent was obtained from each patient.
Data Acquisition and Analysis
Leukocyte count was carried out with a routine analyzer. Flow cytometry was
performed with a FACS scan instrument equipped with a CellQuest software (Becton
Dickinson). Quantification of lymphocyte surface markers expression and
intracellular cytokines content was based on measurement of fraction
(percentage) of positive events. Enzyme-linked immunosorbent assays (ELISA) were
performed using the ASYS HITECH Microplate Reader (Eugendorf).
Surface Staining
Blood samples were processed as previously described for monocyte
analysis[19].
In brief, K3-EDTA samples were treated with a red cell lysing
solution to isolate leukocytes, and then stained with the following combinations
of phycoerythrin (PE)/cyanine 5 (Cy5) [PC5]-, fluorescein isothiocyanate
(FITC)-, and (PE)-conjugated monoclonal antibodies: CD4/CD28/CD25;
CD4/CD28/CCR5; CD4/CD28/CXCR3; CD4/CD28/CX3CR1. Isotype controls were performed.
The antibodies used were from Becton Dickinson, Pharmingen, Immunotech, R&D
Systems, and MBL International Corporation. The acquisition was stopped after
30,000 CD4+ T-lymphocytes were collected for each sample (Figure 1).
Fig. 1.
Identification of total CD4+ and CD4+CD28-
T-lymphocyte subsets. Top: Selection of total CD4+
T-lymphocytes (R1) based on side scatter (SSC) properties and FL3
(CD4) bright fluorescence (A). R1-derived dot-plot of FL1 (CD28)
versus FL3 (CD4) used to identify and quantify (as percentage) the
CD4+CD28- T-lymphocyte fraction (R2) (B).
Bottom: Example of quantification of CX3CR1 expression. The overlay
of the FL2 fluorescence histograms is used to quantify the CX3CR1
positive cells (as percentage, events in M1, filled histograms) on
the selected total CD4+ (R1 gated) (C) and
CD4+CD28- (R1 + R2 gated) (D) T-cell subsets, when
compared with isotype controls (dotted histograms).
Intracellular Cytokines
A whole blood staining procedure for intracellular cytokines detection was
carried out as described[20]. In brief, heparinized samples were diluted 1:1 with
RPMI (acronym for Roswell Park Memorial Institute) medium complete medium and
incubated at a density of 2x106 leukocytes/ml for 5 hours, at 37°C.
Phorbol myristate acetate (PMA) (Sigma Chemical Co.) (50 ng/ml) and ionomycin (1
mg/ml) were used as in vitro activators, in the presence of
brefeldin A (eBioscience Inc.) (3 mg/ml final) as secretion blocking agent.
Samples were then recovered, treated with a red cell lysing solution, and the
isolated white blood cells (WBC) were stained with a combination of PC5- and
FITC-conjugated monoclonal antibodies against the markers CD3 and CD4,
respectively. Then, the cells were fixed with formaldehyde (2%) and
permeabilized with saponin (0.5%). Cytokine staining was performed with
PE-labeled monoclonal antibodies against IL-17A, IL-21, IFN-γ and IL-10.
Isotype controls were performed. The acquisition was stopped after 30,000
CD3+ T-lymphocytes were collected for each sample (Figure 2).
Fig. 2.
Analysis of intracellular interferon gamma (IFN-y) production by
CD4+ T-lymphocytes in activated whole blood samples,
as example of cytokine-producing cells quantification by dot-plot
quadrant statistics. Top: Selection of CD3+ T-lymphocytes
(R1) based on side scatter (SSC) properties and FL3 (CD3) bright
fluorescence (A). R1-derived dot-plot of FL2 (IFN-γ) vs. FL1
(CD4) used to select the CD4+ T-lymphocyte fraction (R2)
(B). Bottom: Quantification (as percentage) of CD4+
T-cells producing IFN-γ (upper right quadrant, R1 + R2 gated)
(D), when compared with isotype control (upper right quadrant, R1 +
R2 gated) (C).
CD4+CD25highFoxP3+ Regulatory
T-cells
Blood samples were processed as published[21], with minor modifications. In brief,
K3-EDTA samples were lysed as described before and the isolated
WBC stained with PC5- and PE-conjugated monoclonal antibodies against the
markers CD4 and CD25, respectively. Then, cells were fixed, permeabilized, and
stained with an Alexa Fluor 488-conjugated anti-human forkhead-box-P3 (FoxP3)
monoclonal antibody (eBioscience Inc.). Isotype controls were performed. Treg
cells fraction was quantified into the 2% of CD4+CD25high
double positive events, as reported[22]. The acquisition was stopped after 30,000
CD4+ T-lymphocytes were collected for each sample (Figure 3).
Fig. 3.
Gating sequence used to quantify the frequency of regulatory
T-cells (Treg). Top: Identification of CD4+ T-lymphocytes
(R1) based on side scatter (SSC) properties and FL3 (CD4) bright
fluorescence (A). R1-derived dot-plot of FL2 (CD25) vs. FL3 (CD4)
used to select the CD25high positive events at the top 2%
of CD4+CD25+ T-cells (R2) (B). Bottom: The
overlay of Alexa Fluor-488 FL1 fluorescence histograms is used to
quantify the percentage of FoxP3+ cells (events in M1,
filled histogram) (C), when compared with isotype control (dotted
histogram). By multiplying the above 2 percentages and dividing them
by 100, the CD4+CD25highFoxP3+
(Treg) cell fraction is calculated.
ELISA
Plasma levels of IFN-γ, IL-6, IL-10, IL-17A, IL-23, TGF-β, RANTES,
and CX3CL1 were quantified by ELISA kits (R&D Systems) according to the
manufacturer's protocol. Minimum detectable concentrations were 15.6 pg/ml
(IFN-γ, 0.156 pg/ml (IL-6), 0.78 pg/ml (IL-10), 31.25 pg/ml (IL-17A),
39.0 pg/ml (IL-23), 0.031 ng/ml (TGF-β, 0.031 ng/ml (RANTES), and 0.156
ng/ml (CX3CL1).
Statistical Analysis
The calculation of mean values and standard errors of the mean (± SEM), as
well as the determination of linear correlations among continuous variables, was
carried out with the StatView 5.0 software. The existence of statistically
significant differences between groups was explored by analysis of variance
(ANOVA) Student's t-test for unpaired data. Chi-squares statistical analysis was
used for comparison between nominal variables. P<0.05 was
considered statistically significant.
RESULTS
General Findings
Mean aortic diameters were 35.50 ± 1.65 mm (range 27-43 mm) and 48.90
± 1.18 mm (range 45-56 mm) in AVD and TAA groups, respectively
(P<0.0001). At the operation, 8 patients had an isolated
ascending aorta replacement and 2 underwent combined aortic root surgery.
Clinical and surgical characteristics of patients are reported in Table 1.
Table 1.
Patients' clinical characteristics (n=20).
The mean frequency (± SEM) of CD4+ T-lymphocytes was
46.39±1.99; their absolute number (number of cells/ml) was
953.34±61.85, accordingly with published CD4+ T-cell reference
values during aging[23]. The mean percentage (± SEM) of
CD4+ T-cells belonging to the pro-inflammatory/cytotoxic subset
CD4+CD28- was 5.56±1.33; this value, if
referred to the age's range of our patients, is also in accord with literature
data[24].
Cumulative analysis of surface markers expression on total CD4+ and
CD4+CD28- T-cell subsets are reported in Table 2.
Table 2.
Surface marker expression on blood total CD4+and
CD4+CD28- T-cell subsets (percentage of positivity, mean
±SEM) (20 patients).
The measurement of plasma cytokines/chemokines is reported in Table 3.
Table 3.
Blood levels of cytokines and chemokines (mean ± SEM) (20
patients).
No relationship has been found between the plasma levels of cytokines/chemokines
and CD4+ T-cells phenotypes, intracellular cytokines levels, and
presence of TAA, respectively. Cumulative quantification of intracellular
cytokines is reported in Table 4; also,
these data are in accordance with published T-cell cytokines levels in normal
aging[25].
Table 4.
Intracellular cytokine production by "in vitro"timulated CD4+ T-lymphocytes in whole blood samples(percentage of positivity, mean ± SEM) (20 patients).
Relationships among CD4+ T-cells Phenotypic Subsets
The accumulation of CD28- T-cells in aging is driven by a repeated
cellular activation, leading also to a modulation of CD4+ T-cells
chemokine receptors expression[24,26].
In our patients, a tightly positive correlation between the circulating fraction
of CD4+CD28- T-cells and the frequency of CD4+
T-lymphocytes carrying the fractalkine receptor CX3CR1
(CD4+/CX3CR1+) (P<0.0001, R=0.934)
has been observed.
Relationships among CD4+ T-cells Phenotype/Function and
TAA
The sum of blood CD4+CD28±CD4+/CX3CR1+
T-cells fractions was significantly lower in the TAA group than in the AVD group
(Figure 4).
Fig. 4.
Total fraction (percentage) of circulating
CD4+CD28±CD4+/CX3CR1+
T-cells in AVD (white bar) and TAA (black bar) patients. Statistical
comparison between percentages (mean ± SEM) was performed
with ANOVA Student's t-test for unpaired data. Statistically
significant differences (P<0.05) were detected. The functional
balance (ratio) between the blood frequency of CD4+
T-lymphocytes producing IFN-γ vs. the total fraction of
IL-17A+IL-21 cytokine-producing CD4+ T-cells was
significantly lower in the TAA group (Figure 5A).
ANOVA=analysis of variance; AVD=aortic valve disease;
IFN-γ=interferon gamma; IL=interleukin; SEM=standard error of
the mean; TAA=thoracic aortic aneurysm
For each patient, the sum of
CD4+CD28±CD4+/CX3CR1+
T-cell fractions correlated positively with the corresponding values of the
above-mentioned cytokine ratio (Figure
5).
Fig. 5.
(A) Ratio between the blood frequency of CD4+
T-lymphocytes producing IFN-γ vs. the total fraction of
IL-17A+IL-21 cytokine-producing CD4+ T-cells in AVD
(white bar) and TAA (black bar) patients. Statistical comparison
between ratios (mean ± SEM) was performed with ANOVA
Student's t-test for unpaired data. Statistically significant
differences (P<0.05) were detected. (B) Linear regression between
the individual fractions (percentages) of circulating
CD4+CD28±CD4+/CX3CR1+
T-cells (x axis) and the corresponding IFN-γ/(IL-17A+IL-21)
cytokine ratios (y axis) in all patients.
ANOVA=analysis of variance; AVD=aortic valve disease;
IFN-γ=interferon gamma; IL=interleukin; SEM=standard error of
the mean; TAA=thoracic aortic aneurysm
The mean value (± SEM) of blood Treg, expressed as a percentage of total
CD4+ T-cells, was 1.19±0.09; no statistically significant
differences have been observed between the 2 groups of patients. The frequency
ratio of
CD4+CD28±CD4+/CX3CR1+
T-cells vs. circulating Treg was significantly lower in the TAA
group than in the AVD group (Figure 6).
Fig. 6.
Frequency ratio of blood
CD4+CD28±CD4+/CX3CR1+
T-cells vs. Treg in AVD (white bar) and TAA (black bar) patients.
Statistical comparison between ratios (mean ± SEM) was
performed with ANOVA Student's t-test for unpaired data.
Statistically significant differences (P<0.05) were
detected.
ANOVA=analysis of variance; AVD=aortic valve disease; SEM=standard
error of the mean; TAA=thoracic aortic aneurysm; Treg=regulatory
T-cells
CD4+ T-cells Phenotype/Function, Aortic Valve Pathology, and
Clinical Features
No correlation has been found between all the above-mentioned phenotypic and
functional CD4+ T-cell features and the patients' ages. Student's
t-test and linear regression statistical analysis did not show significant
associations between immunological parameters and patients' clinical
characteristics, such as the prevalent type of valve pathology, aortic valve
mean gradient values, associated risk factors, and current medical therapies.
The mean functional ratio of IFN-γ vs. IL-17A+IL-21
cytokine-producing CD4+ T-cells was significantly higher
(P=0.0228) in female (2.144±279) than in male
patients (1.398±0.147). On the other hand, the Chi-squares statistical
analysis did not evidence a significant association between sex differences and
the presence of TAA.
DISCUSSION
Clinical and experimental studies carried out in abdominal aortic aneurysms by
histological and immunohistochemical procedures proved that a predominant
Th2-mediated immune response, mainly driven by IL-4, IL-5, or IL-10 cytokines,
induces severe aneurysm formations[27]. On the other hand, a prevalent Th1-mediated immune
response, sustained by the infiltration of mononuclear cells releasing IFN-γ
and IL-12, has been demonstrated to be responsible for transmural inflammation and
external vessel wall dilatation in ascending TAA[28]. Nevertheless, independently of their
localization, both types of dilating aortic lesions are characterized by common
histopathological findings, also including evident modifications of ECM turnover
with overexpressed collagen deposition and fibrosis[2-4]. As known, the ECM remodeling, with fibroblasts activation and
development of fibrosis, occurring in pathological conditions, such as systemic
sclerosis, atherosclerosis, parasitic infections, and after the use of silicone
mammary implants, is suppressed by a locally polarized Th1 IFN-γ-driven
immune response[5,6].
In spite of the histological demonstration of a prevalent Th1-mediated immune
response into the wall of dilated and/or aneurysmatic ascending thoracic aortas, so
far little is known about the systemic immunological status of these patients.
Therefore, in our study, we evaluated several phenotypic and functional features of
peripheral blood CD4+ T-lymphocytes in patients undergoing a cardiac
operation for aortic valve replacement associated, or not, with elective surgery for
TAA.
We have found out that the cumulative CD4+ T-cell fraction calculated by
percentage addition of circulating CD4+CD28- T-cells, a subset
of cytotoxic T-lymphocytes producing large amount of IFN-γ[29], plus CD4+ T-cells
carrying the fractalkine receptor CX3CR1 (CD4+/CX3CR1+),
preferentially expressed on Th1 IFN-γ-producing cells[30], is significantly lower in the
TAA group than in the AVD group.
A prevalent pro-fibrotic IL-17A/IL-21-driven polarization of blood CD4+
T-lymphocytes in the TAA group seems to be demonstrated, in our study, by the
significantly lower mean ratio observed between the frequency of CD4+
T-lymphocytes producing the anti-fibrotic IFN-γ vs. the
total fraction of IL-17A+IL-21 pro-fibrotic cytokine-producing CD4+
T-cells[6,31,32].
Previous papers demonstrated that human fibrocytes are potent antigen-presenting
cells (APC) capable of priming naive T-cells in situ[33]. Since the
requirement of a more broad T-cell/APC cross-talk via CD40-CD40L interactions for
generation of Th17- than for Th1 IFN-γ-mediated inflammatory
responses[34,35], our data suggest that the
higher extent of fibrotic tissue may orientate aneurysmatic aorta tissue-resident
T-cells towards a prevalent production of IL-17A and IL-21, so creating a
self-maintaining loop for further fibrocyte priming, collagen production, and
pro-fibrotic tissue remodeling.
The close correlation existing between the individual values of the
IFN-γ/(IL-17A + IL-21) functional ratio and the
CD4+CD28±CD4+/CX3CR1+ T-cell
fraction indicates that the association of these last phenotypic T-cell subsets is
strictly involved in the establishment of a prevalently IFN-γ-oriented blood
CD4+ T-cell polarization in AVD subjects without TAA.
Moreover, while previous papers showed that Treg and IL-10-producing CD4+
T-lymphocytes separately suppress collagen deposition and tissue fibrosis in several
chronic inflammatory conditions[5], apparently cooperating with the anti-fibrotic action of
IFN-10-producing CD4+ T-cells, the significantly lower ratio of total
CD4+CD28±
CD4+/CX3CR1+ T-cell vs. Treg observed in
our TAA patients suggests that a tissue IFN-γ counter-acting effect of Treg,
mainly sustained by IL-10 production[36], could be detrimental for this specific pathological
condition.
Previous papers have shown that circulating T-cells represent an important repository
pool to reveal tissue-resident T-cell functional abnormalities in immune-mediated
connective pathologies, such as systemic sclerosis, characterized by a deregulated
fibroblast activation leading to fibrosis of internal organs[37]. Moreover, it has been
demonstrated that the evaluation of balanced dynamic inter-relationships among
different phenotypic/functional characteristics of blood CD4+ T-cell
subsets can identify unique immunological features correlating with the clinical
outcome and therapeutic interventions in selected cardiovascular
patients[9].
CONCLUSION
We conclude that there is a presence of an immunological imbalance in the form of
fibrocyte activation and Treg differentiation leading to the development of an
aortic aneurysm in patients with AVD. On this basis, eventually, a targeted
therapeutic model could be developed if it is confirmed in a large number of
patients with TAA, including subjects with aortic rupture/dissection.
Tabla 6.
Authors’ roles & responsibilities
ACKNOWLEDGMENTS
The authors wish to express their appreciation to the personnel of the "G.
Pasquinucci" Heart Hospital operating theater, intensive care unit, and laboratory
departments.
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Notes
Notes
This study was carried out at the College of Medical Sciences, Teaching Hospital,
Bharatpur, Chitwan, Nepal.
Financial support: This study was funded by the National Research Council (CNR)
of Italy, RSTL (Curiosity Driven Research Program) DG.RSTL.035.006-035.
Fast Track
Conflict of interest declaration
No conflict of interest.
Author notes
Correspondence Address: Kaushal K. Tiwari,
http://orcid.org/0000-0002-1345-3223, Department of Cardiothoracic and Vascular
Surgery, College of Medical Sciences, Teaching Hospital, P.O. Box 23,
Bharatpur-10, Chitwan, Nepal - Zip Code: 44207. E-mail:
drkaushalkt@yahoo.com
