Clin Exp Med DOI 10.1007/s10238-014-0285-6

ORIGINAL ARTICLE

Myc and AP-1 expression in T cells and T-cell activation in patients after hematopoietic stem cell transplantation Shivtia Trop-Steinberg • Yehudith Azar Rachel Bringer • Reuven Or

•

Received: 25 November 2013 / Accepted: 29 March 2014 Ó Springer-Verlag Italia 2014

Abstract Regeneration of the immune system after hematopoietic stem cell transplantation (HSCT) is a slow process. We attempted to identify problems in the recovery of the immune system by examining expressions of early event cell cycle proteins Myc, Jun, and Fos, as well as DNA binding of Myc, activating protein 1 (AP-1), and CD4 cell activation values, in phytohemagglutinin-activated T lymphocytes taken from patients after HSCT. HSCT patients showed lower protein expression levels of Myc and Jun, as well as Myc and AP-1 DNA-binding levels, as compared to healthy controls. C-Jun was lower in long-term survivors of HSCT than short-term survivors. Adenosine triphosphate (ATP) values in CD4 cells were also lower in HSCT patients than healthy controls, but showed a time-dependent increase post-transplant. Non-surviving patients showed lower levels of both Fos protein and ATP as compared to surviving patients and a negative correlation between Fos values and lymphocyte percentage that was not present in surviving patients. There was a strong positive correlation between Fos values and lymphocyte percentage and between AP-1 values and white blood count, in patients without graft-versus-host disease (GVHD), that did not exist in patients who suffered from GVHD. Patients 2 years post-HSCT showed a positive correlation between AP-1 and Myc DNA-binding protein values, similar to those values found in healthy controls. Our study identified S. Trop-Steinberg Faculty of Medicine, The Institute of Medical Research, Hebrew University of Jerusalem, Jerusalem, Israel S. Trop-Steinberg
Y. Azar
R. Bringer
R. Or (&) Department of Bone Marrow Transplantation, Hadassah-Hebrew University Medical Center, Hadassah Medical Organization, Kiryat Hadassah, P.O.B. 12000, 91120 Jerusalem, Israel e-mail: [email protected]

significant factors that account for the delay in immune reconstitution after transplant; this knowledge may improve the management of post-HSCT patients. Keywords

HSCT
c-Myc
c-Jun
c-Fos
AP-1
GVHD

Introduction Immune reconstitution after hematopoietic stem cell transplantation (HSCT) is vital for successful treatment outcomes. T-cell function deficiency seen in patients posttransplant is a major cause of morbidity and mortality. The delay in the recovery of T-cell numbers and function posttransplant leaves patients susceptible to infection [1]. Moreover, in allogeneic transplants, the regeneration of the immune system is furthered delayed by the development of graft-versus-host disease (GVHD). Adult transplant patients appear to have a mild deficiency of phenotypically naı¨ve CD4? T cells and T-cell receptor excision circles (by-products of T-cell receptor gene rearrangement). In vitro stimulated peripheral blood cells from recipients of autologous or allogeneic HSCT show significant T-cell function insufficiency at three distinct levels: cell proliferation, cytokine production, and lytic capacity [2–8]. Proto-oncogenes c-Myc, c-Jun, and c-Fos enable cells to undergo DNA synthesis and advance through the cell cycle [8–10]. A structural and functional analysis demonstrates their role in the signal transduction pathways of growth factors 8. The proto-oncogenes C-Myc, c-Jun, and c-Fos present in T cells are involved in the production and lytic capacity of CD8 T lymphocytes and in the control of cellular proliferation and activation [5, 8, 9, 11, 12]. These proteins, found in the cytoplasm at particular stages of cell growth, vary in their nuclear localization

123

Clin Exp Med

according to the proliferation state of the cells [13, 14]. The transport of the protein product of c-Fos proto-oncogene from the cytoplasm into the nucleus, where it functions as part of the AP-1 transcription complex, is not spontaneous but depends on the continuous stimulation of cells by serum factors [14]. Activating protein 1 (AP-1) describes a group of structurally and functionally related members of the Jun and Fos protein families [15], which are involved in the synthesis and assembly of Fos and/or Jun proteins. AP-1 is also involved in the control of both basal and inducible transcription of several genes containing AP-1 sites. In human Jurkat T-cell line, Myc, Jun, and Fos are hub genes in network and have documented roles in cell proliferation, differentiation, and activation responses [16]. In addition, they activate IL-2 and IL-2R expression [17, 18]. C-Myc plays an important role in various physiological processes including cell growth, proliferation, loss of differentiation, and cell death (apoptosis) [19]. In T lymphocytes, c-Myc induction is biphasic and associated with both early and late activation events [20]. C-Myc is involved in early hematopoietic development and is reported to control the self-renewal of hematopoietic stem cells [21]. Upon induction of pre-TCR signaling, c-Myc is rapidly up-regulated and is required for the proliferation, but not the differentiation or survival signals, emanating from the pre-TCR [21]. In T-cell metabolic transcriptome, c-Myc drives metabolic reprogramming in activated primary T lymphocytes [22]. Deletion of Myc markedly inhibited activation induced glycolysis and glutaminolysis in the T cells [22]. Stimulation of the TCR/ CD3 complex on normal peripheral blood human T cells leads to the induction of c-myc mRNA expression, and abundant c-myc mRNA is expressed throughout the proliferative response [23]. Quiescent T cells treated with antisense c-Myc oligonucleotides failed to enter S phase when stimulated with mitogens [23]. In T lymphocytes, AP-1 composed of c-Jun and c-Fos heterodimers is essential in the TCR signaling pathway, lymphokine production, and lymphokine receptor signaling [24]. AP-1 is efficiently activated by CD28 costimulation in the presence of TCR signaling and plays an important role for IL-2 gene expression and proliferative response [10, 25–29]. Fos and Jun proteins have been shown to be components of the transcriptional factor NFAT in both murine and human T cells [30]. Studies in Jurkat cells have shown that different combinations of NFAT and AP-1 protein tune the level of IL-2 transcription at different points during T-cell stimulation [31]. However, in the absence of AP-1, NFAT imposes a genetic program of lymphocyte anergy [32]. Anergized human CD4? T cells display a highly specific reduction in binding at the AP-1 binding site in the IL-2 gene promoter [33].

123

The current study investigated how impaired T-cell immune function post-HSCT originates in the inappropriate expression and activity of c-Myc and AP-1 proteins, which parallels an insufficiency of adenosine triphosphate (ATP) in the T cell. We examined the expression levels of these proteins and their activity post-phytohemagglutinin (PHA) incubation in peripheral T cells and T-cell activity during the recovery period in patients post-allogeneic HSCT and compared them to those of healthy controls. In order to examine the T-cell activity during the recovery period in patients post-allogeneic HSCT, we used a diagnostic assay that measures intracellular ATP (iATP) concentration in CD4? cells following PHA incubation (Immuknow Cylex Immune Cell Function Assay kit). This activation assay intracellular adenosine triphosphate (iATP) levels is correlated with cell proliferation and is known to be an early indication of the response to immune stimuli [34, 35].

Patients and methods Patients Fifty-nine adult patients who had received an allogeneic transplantation between May 2000 and February 2007 in the Department of Bone Marrow transplantation at Hadassah-Hebrew University Medical Center were examined in the course of this study. All patients provided written informed consent approved by the Hospital Ethical Committee, and all procedures were in accordance with the Helsinki Declaration of 1975. For each experiment, all HSCT patients were tested once. Some patients participated in more than one experiment within the study. Chimerism data were taken randomly from 1/3 of the allogeneic HCST patients who participated in the study using FACS analyses. In all tested patients, blood cells were from donor origin. Subjects were divided into the following: In the protein study/binding to DNA study groups, Con: healthy volunteer controls (n = 10/18); 3 months to 2 years: patients 3 months to under 2 years post-transplantation (n = 22/16); 2–8 year: patients from 2 to 8 years post-transplantation (n = 11/9); GVHD: patients with GVHD (n = 21/12); No GVHD: patients with no GVHD (n = 10/9); alive: patients who survive until the end of the study (3 years) (n = 25); dead: patients who did not survive until the end of the study (3 years) (n = 7). In the ATP study groups, Con: healthy volunteer controls (n = 13); 3–6 months: patients 3–6 months post-transplantation (n = 24); 9–12 months: patients 9–12 months post-transplantation (n = 9); 15–24 months: patients from 15 months to 2 years post-

Clin Exp Med Table 1 Characteristics of patients and control in the study Characteristics

Patients (n = 59)

Control (n = 41)

Age range (years)

8–65

18–62

Age average/median (years)

31/33

39/36

Gender (male/female)

30/29

25/16

Primary disease AML

21

CML

5

ALL

11

MDS

6

NHL

7

MM

2

HL SAA

1 3

ALD

3

Donors HLA-identical sibling

31

HLA-mismatched family

14

Unrelated

14

Conditioning regimen FLU/BU/ATG

39

TBI/TT/CY

17

FLU/CY/T

3

Acute GVHD Grade I–II

29

Grade III–IV

12

Chronic GVHD Limited

16

Extensive

16

AML acute myeloid leukemia, CML chronic myelogenous leukemia, ALL acute lymphoid leukemia, MDS myelodysplastic syndrome; NHL non-Hodgkin’s lymphoma, MM multiple myeloma, HL Hodgkin’s lymphoma, SAA severe aplastic anemia, ALD adrenoleukodystrophy, FLU fludarabin, BU Busulfex/busulfan, ATG anti-thymocyte globulin, TBI total body irradiation, TT thiotepa, CY cytoxan, T thymoglobulin

transplantation (n = 8); 9–24 months: patients from 9 months to 2 years post-transplantation (n = 17); alive: patients who survive until the end of the study (2 years)/ patients who survived 6 months post-transplant (n = 23/ 16); dead: patients who did not survive until the end of the study (2 years)/patients who did not survive within 3–6 months post-transplant (n = 10/9). Patient characteristics and demographics are described in Table 1. Patients with hematological malignancies were conditioned with intravenous (IV) fludarabine (30 mg/m2/ day on days -10 to -5) and oral busulfan (1 mg/kg 4 times daily on days -6 and -5); patients receiving antihuman T-lymphocyte globulin were given doses of 5–10 mg/kg per day on days -4 to -1 (protocol FLU/BU/ ATG). Patients conditioned with total body irradiation

(TBI) received a total dose of 12 Gy in six fractions, IV thiotepa (5 mg/kg day for 2 days), and IV cyclophosphamide (60 mg/kg/day for 2 days) (protocol TBI/TT/CY). Three patients with severe aplastic anemia were conditioned with IV fludarabine (30 mg/m2 per day for 6 days), cyclophosphamide (60 mg/kg/day for 2 days), and thymoglobulin (2.5 mg/kg for 4 days) (protocol FLU/CY/T). GVHD prophylaxis consisted of IV cyclosporine 3 mg/kg/ day from day -4. For transplanted patients not showing signs of GVHD, cyclosporine was tapered off beginning on day 50 with discontinuation of the drug within 40 days. For patients with acute GVHD, methylprednisolone 2 mg/kg/ day was added to cyclosporine. Patients with extensive chronic GVHD were treated with various combinations of immunosuppressive medications. The number of patients who suffered from acute and chronic GVHD was graded according to an international score. Blood samples Peripheral blood samples were obtained from allogeneic patients after HSCT as part of the regular post-transplant immune monitoring process as well as from healthy volunteers. Blood was sampled once at 3 months post-HSCT or at different times up to 2 years post-transplantation. In longterm HSCT patients (2–8 years post-transplant), blood was sampled once during a routine follow-up visit. T-cell enrichment The obtained blood samples (20 ml each) were mixed with 10 % anticoagulant citrate–phosphate–dextrose (CPD) solution (Baxter Healthcare Corporation, USA). Two milliliter RosetteSep lymphoid (CD3?) enrichment cocktail (StemCell Technologies, Canada) was added, followed by a 20-min incubation at room temperature. The blood was diluted two folds in RPMI 1640 and applied over 15 ml FicollTM sodium diatrizoate (Sigma, USA). The samples were centrifuged for 25 min at 1,2009g at room temperature. The T-cell-enriched plasma interface was collected and washed twice with PBS. Flow cytometer analysis to evaluate T-cell purity Cells (0.5 9 106) derived from the T-cell enrichment process were suspended in 100 ll PBS and incubated with 0.5 lg FITC antihuman CD3 (Southern Biotechnology Associates, USA) for 1 h at 4 °C. They were then washed twice with PBS and centrifuged for 4 min at 1,2009g. The cells were re-suspended in 0.5 ml PBS and analyzed by FACScan (Becton–Dickinson, Belgium). CD3? T-cell purity was 95–97 % (‘‘Appendix’’).

123

Clin Exp Med

In vitro T-cell activation for the protein study Isolated T cells were incubated with 10 lg/ml phytohemagglutinin-P (PHA-P) (Sigma) (1.5 9 106 cells/ml) in a 25-cm2 flask in RPMI medium supplemented with 5 % FCS, 2 mM L-glutamine, 100 unit/ml penicillin, and 100 mg/ml streptomycin for 48 h at 37 °C, 5 % CO2 with humidity. Nuclear and cytoplasmic protein isolation Nuclear and cytoplasmic protein isolation was prepared using the NE-PER nuclear and cytoplasmic extraction reagents kit (Pierce, Rockford, IL) supplemented with protease inhibitor according to the instructions provided by the manufacturer. The protein concentration was determined with Coomassie Plus Protein Assay Reagent (Pierce, Rockford, IL). Western blot analysis The nuclear and cytoplasmic proteins were separated by SDS-polyacrylamide gel electrophoresis (PAGE) on 12 % gels and electroblotted to nitrocellulose membrane. The proteins were blocked with PBS/0.05 % Tween-20 containing 5 % non-fat milk solids and incubated overnight with primary antibodies as follows: mouse anti-c-Myc (1:100 dilution), rabbit anti-c-Jun (1:100) (Santa Cruz, USA), rabbit anti-c-Fos (1:100) (Abcam, UK), rabbit antitubulin (1:100) (Santa Cruz, USA). Primary antibody was detected after incubation of 1 h with HRP-conjugated goat anti-mouse IgG 1:1 or HRP-conjugated goat anti-rabbit IgG 1:1 (Jackson, USA). The proteins were detected by the ECL Western blotting analysis system (Amersham Pharmacia, Arlington Height, IL). Results were normalized with tubulin values, and all tubulin values were normalized with the same tubulin control volunteer, whose value was determined as 1. Results were analyzed using 1.38X ImageJ Software (Wayne Rasband, National Institutes of Health, USA). DNA probes and electrophoretic mobility shift assay (EMSA) The oligonucleotides used as DNA probes were as follows: Forward oligo: MYC-EMSA: GGAAGCAGACCACGTG GTCTGCTTCC (Sigma Genosys Packing Slip); reverse oligo: MYC-EMSA: GGAAGCAGACCACGTGGTCTGC TTCC (Sigma Genosys Packing Slip.). Forward oligo: AP1-EMSA: GTCAGTCAGTGACTCAATCGGTCA (Sigma Genosys Packing Slip); reverse oligo: AP-1-EMSA: TGACCGATTGAGTCACTGACTGAC (Sigma Genosys Packing Slip). Electrophoretic mobility shift assay was performed using the electrophoretic mobility shift assay

123

(EMSA) kit (Invitrogen Detection Technologies) according to the instructions provided by the manufacturer. The binding reactions were carried out in a total volume of 10 ll with 1 lg/ll Forward and 1 lg/ll reverse oligos and 50 lg/ll nuclear protein. The reaction mixture was separated by non-denaturing PAGE on 8 % gels, and the nucleic acids were stained with SYBR Green EMSA nucleic acid gel stain (Invitrogen detection technologies). The stained nucleic acids were visualized with an appropriate photographic filter. Results were analyzed using 1.38X ImageJ software (Wayne Rasband, National Institutes of Health, USA) and were normalized with the same control, whose value determined as 1. Western blot for Gel-shift (EMSA) In order to prove that the bands are Myc’s and AP-1’s, EMSA gels were electroblotted to the nitrocellulose membrane. As described above, the blots were incubated with specific primary antibodies: mouse anti-c-Myc, rabbit anti-c-Jun (Santa Cruz, USA), and rabbit anti-c-Fos (Abcam, UK) and detected with HRP-conjugated goat antimouse IgG or HRP-conjugated goat anti-rabbit IgG (Jackson, USA). Since the EMSA system by which the DNA is dyed directly by SYBR Green does not allow for analysis by the cold probe method, we used the appropriate protein’s antibodies to prove that the high molecular band is the binder of the DNA to the appropriate protein. T-cell activation T-cell activation was measured by determining the increase in intracellular ATP (iATP) concentration in CD4? cells from allogeneic HSCT patients and healthy controls following PHA incubation, using the Immuknow Cylex Immune Cell Function Assay kit (CYLEX Inc., Colombia, MD, USA) [1, 19–21]. In 2002, the Food and Drug Administration (FDA) approved an in vitro assay designed to measure increases in intracellular ATP (iATP) following CD4 cell activation for use in assessing immune function status in transplant patients to monitor lymphocyte function [34]. This ATP assay is a structured kit, for CD4 cells only, that influences all immune functions [34]. Briefly, blood was diluted 1:4 with RPMI 1640 medium. To 100 ll of the diluted blood, 25 ll of a solution of phytohemagglutinin L (PHA) from the kit was added and the mixture was cultured overnight at 37 °C in an incubator with 5 % CO2. Magnetic beads coated with mouse anti-CD4 monoclonal antibodies were added to the mixture, which was then cultured for an additional 30 min at 4 °C. The CD4 cells were collected by a magnet held in position for 15 min. Attached cells were washed and lysed, and iATP release was

Clin Exp Med Table 2 Hematological data of patients Normal values

3–6 months (n = 47)a

9–12 months (n = 29)a

WBC (109/L)

4.5–11

4.6 ± 0.5

Lym %

20–40

12 ± 1.8

13.3 ± 2.6

8.3 ± 1.5

8.7 ± 0.8

MO %

2–13

50 ± 4.2

44 ± 5

29 ± 5.4

58 ± 3.2

NE %

40–75

34 ± 3.4

38 ± 4

43.3 ± 4.2

6.4 ± 1.3*

15–18 months (n = 22)a

2–8 years (n = 19)

7 ± 0.9

9 ± 0.9*

31 ± 3

* 3–6 versus 9–12 months, p \ 0.02, 3–6 months versus 2–8 years p \ 0.04 a

Some of the patients did not participate in the study

measured. The previously suggested guidelines for ascertaining the degree of immunosuppression were used, and iATP concentrations were classified as follows: strong (iATP C 525 ng/ml), moderate or normal (iATP = 226–524 ng/ml), and low (iATP B 225 ng/ml) [19]. Hematologic data Hematological data were taken from hospital medical records. Hematological data for the healthy controls were not measured, but the normal ranges are given. T-cell count was measured in the peripheral blood samples from patients and controls. Statistical methods

Table 3 T-cell count in patients and control groups T-cell count 106 Mean

SE

p*

p**

0.002

Control (n = 12)

2.7

0.09

3–6 m (n = 19)

0.2

0.05

3.5 9 10-6

9–12 m (n = 19)

0.3

0.04

5 9 10-6

0.01

-6

0.03

15–18 m (n = 17)

0.4

0.08

7 9 10

2–8 y (n = 21)

0.5

0.08

1 9 10-6

m months post-HSCT, y years post-HSCT p values are versus * control, ** 2–8 years group (T test)

T-cell activation in vitro post-PHA incubation

Comparison between groups was performed using the Student’s t test and chi-square analysis. If the distribution of the outcome variables showed a marked asymmetry with extreme values, the nonparametric Mann–Whitney test was used. For correlation statistics, the Pearson test was used.

T cells post-PHA incubation in HSCT patients exhibited lower levels of activity in culture than those in healthy controls (Fig. 1a). The comparison between patients and healthy controls was only done post-PHA incubation since cells lacking PHA deteriorate physiologically within 2 days and become abnormal.

Results

Effect of PHA stimulation on c-Myc expression in HSCT patients and controls

Hematologic data White blood count (WBC) significantly increased in the allogeneic patients with time post-transplant. Lymphocyte percentage in HSCT patients was lower than the normal range; however, monocyte percentage was higher than the normal. This result may have been caused by atypical and activated lymphocytes that were classified as monocytes. Neutrophil percentages of post-HSCT were on the lower limit of the normal range and in some patients even below normal (Table 2). T-cell count T-cell count significantly increased with time post-transplant in the allogeneic patients, but was still significantly lower than in healthy controls (Table 3).

Western blot evaluation of cytosol c-Myc expression from T cells that had been exposed to PHA in vitro showed significantly lower values in the 3–24 months post-HSCT group compared with healthy controls. Significant differences were seen between the number of patients and healthy controls who expressed cytosol c-Myc, with less expression occurring in patients (Figs. 1b, 2a, b). No significant differences were observed in the expression of nuclear c-Myc between patients and control (data not shown). No significant differences were observed in the expression of c-Myc between the patients groups (data not shown). Effect of PHA stimulation on c-Jun expression in HSCT patients and controls Cytosol c-Jun expression from T cells post-PHA incubation was significantly lower in all the patients groups compared

123

Clin Exp Med

Fig. 1 Effect of PHA stimulation on T-cells cultures and on c-Myc, c-Jun, and c-Fos expression in T cells. T cells from HSCT patients and healthy control volunteers were incubated for 48 h with 10 lg/ml phytohemagglutinin (PHA) (1.5 9 106 cells/ml) in RPMI medium with supplement. Nuclear and cytosol c-Myc, c-Jun, c-Fos, and btubulin from HSCT patients and control were measured after 48-h PHA incubation. a PHA stimulation caused different activation patterns in culture in patients and control. A culture from one

3 months post-HSCT patient (from 3 to 24 months group) and a culture from one volunteer from the control group are shown. The two pictures were photographed with an inverted microscope (CK40 Olympus) (950), b bands from one patient 3 months post-HSCT (of the 3–24 months group) and from one of the control group are shown. Detection of c-Myc, c-Jun, and c-Fos proteins was performed by Western blot analysis. Abbreviations: Nu nuclear, Cy cytosol

with healthy controls, though the 3–24 months group showed significantly higher cytosol c-Jun expression than the 2–8 years group. Nuclear c-Jun expression was also lower in all the patients subgroups compared with healthy controls. Significantly fewer patients expressed cytosol and nuclear c-Jun than healthy controls (Figs. 1b, 3a–e).

lymphocyte percentage. In contrast, the GVHD group did not show any such correlation (Table 4). No correlation was seen between hematologic values and c-Myc and c-Jun expression values (data not shown).

Effect of PHA stimulation on c-Fos expression in HSCT patients and control No significant differences were observed in cytosol and nuclear c-Fos expression values in patients versus healthy controls and among the different patient groups (data not shown). No significant difference was observed in the number of patients who expressed c-Fos versus control (Fig. 1b). Protein expression after PHA incubation in HSCT patients who developed GVHD during the study (3 years) and those who did not There was a significant positive correlation in the nonGVHD group between nuclear c-Fos expression and

123

Protein expression after PHA incubation in surviving versus non-surviving patients post-HSCT (3 years) No significant differences were seen in c-Myc expression values between the patient groups and healthy controls and among the patient groups themselves. Cytosol c-Jun expression was significantly lower in both surviving and non-surviving groups as compared to healthy controls. Nuclear c-Fos expression was significantly higher in the surviving group than in the non-surviving group (Fig. 4a–c). Correlation between hematologic values and gene expression: A significant negative correlation was observed in the non-surviving group between cytosol c-Fos expression and lymphocyte percentage. In contrast, the surviving group did not show any significant correlation (Table 4). No correlation was seen between hematologic values and c-Myc and c-Jun expression values (data not shown).

Clin Exp Med

Effect of PHA stimulation on c-Myc binding to DNA in HSCT patients and control Gel-shift (EMSA) evaluation of c-Myc binding to DNA from CD3? T cells following PHA incubation showed significantly lower values in the 2–8 years subgroup than in healthy controls. However, no significant difference was seen between the 3–24 months group and healthy controls. Significantly fewer patients showed c-Myc binding to DNA as compared to healthy controls in the 3–24 months group (Fig. 5b, c). No significant differences were observed in the values of c-Myc binding to DNA between the patient groups. Effect of PHA stimulation on AP-1 binding to DNA in HSCT patients and control

Fig. 2 Effect of PHA stimulation on c-Myc expression in T cells from HSCT patients and control. Cytosol c-Myc from HSCT patients and control were measured after 48 h PHA (10 lg/ml) incubation. a Comparison of cytosol c-Myc value in control, 3–24 months, and 2–8 years. Abbreviation: Cy cytosol. b Comparison of the number of individuals among the control, 3–24 month, and 2–8 year who expressed or did not express cytosol c-Myc. Groups analyzed: con: healthy volunteer control (n = 10); 3–24 months: patients 3–24 months post-transplantation (n = 21); 2–8 years: patients 2–8 years post-transplantation (n = 11). ? expressing DNA-binding proteins. - not expressing DNA-binding proteins. Detection of c-Myc protein was performed by Western blot analysis. Results were normalized with Tubulin values using 1.38X ImageJ software (Wayne Rasband, National Institutes of Health, USA), and all tubulin values were normalized with the same tubulin control volunteer, whose value was determined as 1. Results were expressed as mean of expression value ± SE. Differences were tested by the nonparametric Mann– Whitney test or by the T test if the results were parametric. The number of individuals who expressed or did not express c-Myc was tested by the v2 test. *p B 0.05

Gel-shifts (EMSA) and their blot By measuring the electroblotted EMSA gels, we were able to highlight the c-Myc and AP-1 bands binding to DNA proteins precisely (Fig. 5a).

AP-1 binding to DNA from CD3? T cells following PHA incubation showed significantly lower values in the 3–6 months and 2–8 years groups than in healthy controls. Although AP-1 binding to DNA was seen in fewer patients than in the controls, the results were significant only in the 2–8 years group (Fig. 5b, d). No significant differences were observed in the values of AP-1 binding to DNA between the patients groups. Correlation between AP-1 and c-Myc DNA-binding protein values: Both the 2–8 years group and healthy controls demonstrated a significantly high positive correlation between AP-1 and c-Myc. In contrast, none of the other groups showed such significant correlation (Table 5). c-Myc and AP-1 binding to DNA values post-PHA incubation in allogeneic HSCT patients who did and did not develop GVHD during the study (3 years) Correlation between hematological and DNA-binding protein values: The non-GVHD group showed a significant, positive correlation between AP-1 values and WBC. In contrast, the GVHD group did not have such correlation (Table 6). No correlation was seen between hematological and c-Myc DNA-binding protein values (data not shown). Effect of PHA stimulation on ATP values in CD4 cells from HSCT patients and control ATP values in CD4 cells from HSCT patients and control were measured following PHA incubation. ATP values in 3–6 months group were significantly lower than in healthy controls. The 3–6 months group showed significantly lower ATP values than the 9–24 months group (Fig. 6a, b). No significant differences were seen in ATP values between the GVHD and non-GVHD groups (data not shown).

123

Clin Exp Med

Fig. 3 Effect of PHA stimulation on c-Jun expression in T cells from HSCT patients and control. Nuclear and cytosol c-Jun from HSCT patients and control were measured after 48 h PHA (10 lg/ml) incubation. Comparison of cytosol c-Jun values in: a control, 3–24 month, 2–8 year; b 3–24 month and 2–8 year groups; c comparison of nuclear c-Jun values in control, 3–24 month, 2–8 year; Comparison of the number of individuals among controls, 3–24 months, 2–8 years who expressed or did not express c-Jun; d cytosol c-Jun; e nuclear c-Jun. Groups analyzed: con: healthy volunteer control (n = 10); 3–24 months: patients 3–24 months posttransplantation (n = 21/22 from cytosol and nuclear respectively); 2–8 years: patients 2–8 years post-transplantation (n = 11); Groups analyzed: con: healthy volunteer control (n = 10); 3–24 months: patients 3–24 months post-transplantation (n = 21/22 from cytosol

and nuclear, respectively); 2–8 years: patients 2–8 years posttransplantation (n = 11); ? expressing DNA-binding proteins. - not expressing DNA-binding proteins. Abbreviations Nu nuclear, Cy cytosol. Detection of c-Jun protein was performed by Western blot analysis. Results were normalized with tubulin values using 1.38X ImageJ software (Wayne Rasband, National Institutes of Health, USA), and all tubulin values were normalized with the same tubulin control volunteer, whose value was determined as 1. Results were expressed as mean of expression value ± SE. Differences were tested by the nonparametric Mann–Whitney test, or T test if the results were parametric. The number of individuals who expressed or did not express c-jun was tested by the v2 test. *p B 0.05; **p B 0.005; ***p B 0.0005

Effect of PHA stimulation on ATP values in CD4 cells from patients who did and did not survive to the end of the study (2 years)

survivors and non-survivors had significantly lower ATP values than healthy controls (Fig. 6c). Since most of the patients who did not survive were 3–6 months post-transplant, we compared ATP values in the patients who survived or did not survive for 3–6 months. Significantly higher ATP values were seen

Significantly higher ATP values were seen in the survivor group than in the non-survivor group. However, both

123

Clin Exp Med

in the survivor group than the non-survivor group (Fig. 6d).

Discussion The majority of allogeneic HSCT patients have an abnormal T-cell population both in cell number and cell Table 4 Correlation between c-Fos expression and lymphocyte % from HSCT patients Group

r

p

Nuclear c-Fos GVHD (n = 21)

0.06

ns

No GVHD (n = 10)

0.9

0.01

Cytosol c-Fos Survivors (n = 25)

-0.05

ns

Non-survivors (n = 7)

-0.8

0.05

Fig. 4 Effect of PHA stimulation on c-Myc, c-Jun, and c-Fos expression in T cells from HSCT patients who survived and did not survive to the end of the 3-year study. C-Myc, c-Jun, and c-Fos values from T cells from HSCT patients who survived and did not survive to the end of the 3-year study and from control were measured after 48 h PHA (10 lg/ml) incubation: a cytosol and nuclear Fos values in survivors and non-survivors; b cytosol c-Myc, c-Jun, and c-Fos values in survivors, non-survivors, and control; c nuclear c-Myc, c-Jun, and c-Fos values in survivors and non-survivors. Groups analyzed: con:

function. Immune reconstitution is an important component of successful HSCT; however, full recovery of the immune system takes a long time. The prognosis for patients with defective T-cell reconstitution is very poor when compared to those who achieve full T-cell reconstitution [1–7, 36]. Our results demonstrate that patients after HSCT develop qualitative abnormalities in the early event protein cell cycle in T cells, which may account for prolonged impairment of immune reconstitution. These defects may be partially responsible for the deficient CD4 activation. In the current study, we investigated the expression and activity of early event cell cycle proteins c-Myc, c-Jun, and c-Fos in T cells, by evaluating expression levels as well as Myc and AP-1 DNA-binding values. These proteins in both murine and human cells are involved in G0–G1 transition, lymphocyte proliferation, and lymphocyte development, differentiation, and function [8, 9, 16, 25, 30, 37–43]. AP-1 (Fos/Jun) induces a large number of cytokine, as well as other genes, that are central to the productive immune

healthy volunteer control (n = 10); alive: patients who survived to the end of the 3-year study (n = 25); dead: patients who did not survive to the end of the 3-year study (n = 7); detection of c-Myc, c-Jun, and c-Fos proteins was performed by Western blot analysis. Nu nuclear, Cy cytosol. Results were normalized with tubulin values using 1.38X ImageJ software (Wayne Rasband, National Institutes of Health, USA), and all tubulin values were normalized with the same tubulin control volunteer, whose value was determined as 1. Results were expressed as mean of expression value ± SE. *p B 0.05

123

Clin Exp Med

Fig. 5 Effect of PHA stimulation on c-Myc and AP-1 DNA-binding protein values in T cells from HSCT patients and control. C-Myc and AP-1 from HSCT patients and control were measured by Gel-shift (EMSA) after 48 h PHA (10 lg/ml) incubation. a Bands from 3 patients 3 months post-HSCT (from the 3 to 24 months group) and one of the control group are shown, as well as gel without protein. Blots are shown to the right of the Gel-shift. By measuring EMSA gels to their electroblot, we showed the exact bands of c-Myc and AP1 binding to DNA proteins. b c-Myc and AP-1 values from control, 3–24 months, and 2–8 years are shown. Comparison of the number of individuals who expressed or did not express DNA-binding proteins in control, 3–24 months, 2–8 years groups c Myc, d AP-1. Groups

analyzed: con: healthy volunteer control (n = 18); 3–24 months: patients 3–24 months post-transplantation (n = 16); 2–8 years: patients 2–8 years post-transplantation (n = 9); ? expressing DNAbinding proteins. - not expressing DNA-binding proteins. Results were analyzed using 1.38X ImageJ software (Wayne Rasband, National Institutes of Health, USA) and were normalized with the same control, whose value was determined as 1. Results were expressed as mean of expression value ± SE. Differences were tested by the nonparametric Mann–Whitney test, or T test if the results were parametric. The number of individuals that expressed or did not express DNA-binding proteins was tested by the v2 test. *p B 0.05; **p B 0.005

response [32] and also maintain various combinations with NFAT protein to tune the level of IL-2 transcription at various points during T-cell stimulation in Jurkat cells [31]. These proteins were examined due to their important involvement in T-cell function, and the limited information in the current literature that suggests their expression postHSCT for hematological malignant diseases. In addition, CD4 cell activity was evaluated by measuring intracellular

ATP. We demonstrated that c-Myc and c-Jun expression as well as AP-1 and c-Myc DNA-binding protein values were significantly lower in all allogeneic HSCT patients as compared to healthy controls and that c-Jun protein expression values were lower in long-term recovering patients as compared to short-term recovering (up to 2 years post-transplant) patients. This observation suggests that the immunity reconstitution process requires increased

123

Clin Exp Med Table 5 Correlation between AP-1 and c-Myc-binding protein values in HSCT patients and control Group

AP-1 versus c-Myc r

p

3–24 m (n = 16)

0.4

n.s

2–8 y (n = 7)

0.95

0.01

Control (n = 18)

0.86

0.01

m months post-HSCT, y years post-HSCT

Table 6 Correlation between AP-1 binding protein values and WBC from HSCT patients with and without GVHD Group

AP-1 versus WBC r

GVHD (n = 12) No GVHD (n = 9)

p

-0.2 0.8

n.s 0.01

protein values soon after transplant in order to compensate for the above-mentioned immune disorders. A similar pattern of results emerged in a concurrent study [44] in which we found different activation timing in c-jun and c-fos gene expression in patients and healthy controls; c-jun and c-fos expression values in the allogeneic patients differed from each other and increased with time, except for the 2–8 year post-transplant patients, in whom the expression values decreased. These results strongly suggest a different pattern of recovery between short-term- and long-term recovering patients. In the present study, there were no initial significant differences in c-Fos expression between patients and healthy controls. However, long-term surviving patients showed higher levels of c-Fos expression than patients who died within 3 years post-transplant. These observations suggest that defects in AP-1 activation are the cause of impairment of the immune system [25, 40, 42]. Similar observations have been seen in elderly patients undergoing HSCT, where changes in c-Jun mRNA expression and an imbalance in the ratio of c-Fos/c-Jun were noted [25, 39, 40, 45]. After thermal injury in human cells, the expression of mRNA for c-fos is markedly reduced as compared with sham control subjects [30]. These changes relate to the down-regulation of IL-2 formation and defective AP-1 activation [25, 39, 40, 45]. It has been suggested that binding of the Jun protein to the enhancer region of the IL-2 gene in the absence of c-Fos product may act as a negative regulator of transcription; initiation of transcription will not occur until levels of Fos protein increase [30]. The immune reconstitution process in patients most likely requires increased protein expression, and its absence may be a predictor of higher morbidity and

mortality rates. Mitogenic or antigenic stimulation of the TCR complex and appropriate accessory receptors lead to full activation and binding of the AP-1 complex to the IL-2 enhancer region and, presumably, to transcriptional activation of the IL-2 gene [30]. However, in the absence of AP-1, NFAT imposes a genetic program of lymphocyte anergy [32]. Anergized human CD4? T cells display a highly specific reduction in binding at the AP-1 binding site in the IL-2 gene promoter [33]. The strong positive correlation between AP-1 and c-Myc DNA-binding proteins in the 2–8 years group, which is similar to healthy controls but was not present in the 3–24 months patient group, may indicate the presence of a T-cell reconstitution process that is either impaired or lacking in short-term recovering patients post-HSCT. Since AP-1 and c-Myc are required for efficient lymphocyte functioning, their up-regulation is indispensable, and it is therefore essential that both proteins be activated at the same time. Lower ATP levels in CD4 cells in patients compared with healthy controls that showed a time-dependent increase post-transplant confirm our assumption of a partial T-cell reconstitution process post-transplant that was not observed in more recently transplanted patients. The increasing levels were parallel to the up-regulation of lymphocyte percentage and T-cell counts in long-term recovering patients post-transplant and further confirms our hypothesis. Lower levels of ATP in non-surviving patients versus surviving patients suggest that the up-regulation of ATP in patients over time post-transplant might be necessary for survival and its absence, a predictor of mortality. T-cell activation, as measured by determining the increase in intracellular ATP, has been perceived as an early indicator of a response to immune stimuli. Lymphocytes, stimulated by mitogens or antigens, divide in response to a series of activation events, which include clustering of cell surface receptors, increased uptake of metabolites and ions, increased turnover of phospholipids, synthesis of cytokines, and changes in intracellular ATP levels [34]. The intracellular adenosine triphosphate (iATP) levels in the assay are correlated with cell proliferation and expected to be earlier indicators of the response to immune stimuli [34, 35]. It may be utilized in the monitoring of infectious diseases, vaccine efficacy, transplant acceptance, and response to cancer therapy [34, 35]. ATP assay for measuring T-cell activation achieves results within 24 h, provides a faster, easier-to-use method of measuring T-cell activation in response to a variety of stimuli than proliferation testing, and has the additional advantage of identifying the specific T-cell subset involved [34]. This assay also provides an independent measure of the strength of a patient’s immune system and not just as a

123

Clin Exp Med

Fig. 6 Effect of PHA stimulation on ATP values in CD4 cells from HSCT patients and control. Intracellular ATP (iATP) values (ng/ml) from CD4 cells from HSCT patients and control were measured after overnight PHA incubation, using the Immuknow Cylex Immune Cell Function Assay kit. a ATP values from control, 3–6, 9–12, and 15–24 months groups. b ATP values from 3 to 6 months and 9–24 months groups. c Patients who did not survive to the end of the 2-year study period and patients who survive. d Patients who did not survive 3–6 months post-transplant and patients who survive. Groups analyzed in a, b: con: healthy volunteer control (n = 13); 3–6 months: patients 3–6 months post-transplantation (n = 24); 9–12 months: patients 9–12 months post-transplantation (n = 9);

15–24 months: patients 15–24 months post-transplantation (n = 8); 9–24 months: patients 9–24 months post-transplantation (n = 17). Groups analyzed in c: con: healthy volunteer control (n = 13); alive: patients who survived to the end of the study period (n = 23); dead: patients who did not survive to the end of the study period (n = 10). Groups analyzed in d con: healthy volunteer control (n = 13); alive: patients who survived 6 months post-transplant (n = 16); dead: patients who died within 3–6 months post-transplant (n = 9). iATP was measured using the ImmuknowTM test. Results were expressed as mean of expression value ± SE. *p B 0.05; **p B 0.005; ***p B 0.0005

diagnostic tool to predict infection [34, 46]. Although in patients undergoing organ transplantation the immune system is pharmacologically suppressed to achieve tolerance, in HSCT patients, the immune system is, in fact, the transplanted organ, and our use of the ATP assay was to measure the gradually improving function of the new graft by analysis of their ATP levels parallel with their WBC counts. It has been shown that low levels of ATP may indicate a higher risk of infection, whereas increased ATP levels are compatible with rejection of the graft in the context of organ donation. Others have reported higher levels of ATP

in post-organ transplant patients that rejected their transplant versus those who engrafted [47-50]. For HSCT patients, high ATP levels might theoretically signal the risk of GVHD [49]. In our study, there were no significant differences in ATP levels between patients suffering from GVHD group versus patients who did not suffer from GVHD. Though, the high variability among the patient population examined might require a larger sample of patients to determine whether there are significant differences between these groups. We evaluated CD4 T-lymphocyte responses via assessment of increases in the levels of ATP synthesis in patients

123

Clin Exp Med

and healthy controls. As immune reconstitution takes on a more significant role in the assessment of disease management, the measurement of cellular immune function has become very important. CD4 cells, which include effector cells Th1 and Th2, play a critical role in almost all adaptive immune responses. Th1 cells activate macrophages to kill intracellular parasites, and Th2 cells mediate humoral immune response against extracellular pathogens [48]. CD4 cells are critical, in order to create a proper immune system. Our study evaluated the rehabilitation of CD4 cells via ATP levels, since most of the immune activity depends directly or indirectly on the formation of ATP [47, 49, 50]. A positive correlation between both nuclear c-Fos values and lymphocyte percentage (Ly%) and between AP-1 and WBC values was observed in patients with nonGVHD, but not in patients with GVHD. This observation might be inductive of a different recovery patterns in these subgroups of patients. A negative correlation observed between cytosol c-Fos values and Ly% in non-surviving patients that was not observed in surviving patients may be a factor in an immune impairment pattern that predicts a fatal outcome. The importance of c-Myc, c-Jun, and c-Fos is derived from their involvement in the development, proliferation, and activation of lymphocytes, through which they influence the efficient functioning of the immune system [8–10, 51–53]. Our study demonstrated that successively increasing levels of early event cell cycle proteins during the first 2 years post-transplant, in addition to increasing ATP levels to resemble healthy controls, may predict an immune stabilization after HSCT. It is possible that an increase in protein expression in patients while increasing ATP levels could predict success of treatment or illustrate a recovery process, whereas a decrease in these parameters could indicate a higher risk of morbidity and mortality due to incomplete immune recovery. By including our research parameters in the routine post-HSCT follow-up, we believe it may be possible to monitor the rehabilitation of the patient’s immune system more reliably and simultaneously improve treatment, as well as alert the physician to the need for alternative therapy, if necessary. Our research parameters can provide additional clinical information in addition to the usual hematological parameters. We hope that the results of our study together with clinical parameters will help create a more precise diagnostic/prognostic tool for patients receiving HSCT. Further research could investigate whether there is a correlation between cytokine secretion and c-Myc, c-Jun, and c-Fos expression and whether the phosphorylation status of c-Jun contributes to the difference in AP1 activation between patients and healthy controls. In addition, the role of NFk B transcription factor activation and

nuclear translocation as a major regulator of T-cell proliferation and survival should also be further investigated as a possible significant parameter. In conclusion, the results of the present study suggest that there is a different pattern in T-cell protein levels and activity (c-Myc and AP-1) in HSCT patients as compared to healthy controls. The pattern changes with time posttransplant and also in accordance with the patient’s hematological clinical status. The molecular monitoring of the immune reconstitution using c-Fos and AP-1 levels combined with ATP levels and clinical parameters may improve early detection of immune function failure. This follow-up system may allow for the development of more efficient treatments and new interventions designed to hasten desirable T-cell regeneration. Acknowledgments The authors of this paper wish to thank the staff at the bone marrow transplant outpatient clinic for their assistance in the collection of blood samples and clinical follow-up data and the staff members at our laboratory. We also thank Aviva Yoselis of Israel Health Consulting for her editorial assistance. Conflict of interest

None.

Appendix: Evaluation of CD31 T-cell purity Flow cytometry analysis of the enriched cells after the RosetteSep cocktail showed a CD3? T-cell purity of 95–97 %.

References 1. Jimenez M, Ercilla G, Martinez C (2007) Immune reconstitution after allogeneic stem cell transplantation with reduced-intensity conditioning regimens. Leukemia 21:1628–1637 2. Brugnoni D, Airo P, Pennacchio M, Carella G, Malagoli A, Ugazio AG, Porta F, Cattaneo R (1999) Immune reconstitution after bone marrow transplantation for combined immunodeficiencies: down-modulation of Bcl-2 and high expression of

123

Clin Exp Med

3.

4. 5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

CD95/Fas account for increased susceptibility to spontaneous and activation-induced lymphocyte cell death. Bone Marrow Transpl 23:451–457 Storek J, Joseph A, Espino G, Dawson MA, Douek DC, Sullivan KM, Flowers ME, Martin P, Mathioudakis G, Nash RA, Storb R, Appelbaum FR, Maloney DG (2001) Immunity of patients surviving 20 to 30 years after allogeneic or syngeneic bone marrow transplantation. Blood 98:3505–3512 Parkman R, Weinberg KI (1997) Immunological reconstitution following bone marrow transplantation. Immunol Rev 157:73–78 Guillaume T, Rubinstein DB, Symann M (1998) Immune reconstitution and immunotherapy after autologous hematopoietic stem cell transplantation. Blood 92:1471–1490 Poulin JF, Sylvestre M, Champagne P, Dion ML, Kettaf N, Dumont A, Lainesse M, Fontaine P, Roy DC, Perreault C, Sekaly RP, Cheynier R (2003) Evidence for adequate thymic function but impaired naive T-cell survival following allogeneic hematopoietic stem cell transplantation in the absence of chronic graftversus-host disease. Blood 102:4600–4607 Davison GM, Novitzky N, Kline A, Thomas V, Abrahams L, Hale G, Waldmann H (2000) Immune reconstitution after allogeneic bone marrow transplantation depleted of T cells. Transplantation 69:1341–1347 Cleveland JL, Rapp UR, Farrar WL (1987) Role of c-Myc and other genes in interleukin 2 regulated CT6 T lymphocytes and their malignant variants. J Immunol 138:3495–3504 Grausz JD, Fradelizi D, Dautry F, Monier R, Lehn P (1986) Modulation of c-fos and c-myc mRNA levels in normal human lymphocytes by calcium ionophore A23187 and phorbol ester. Eur J Immunol 16:1217–1221 Modiano JF, Mayor J, Ball C, Chitko-McKown CG, Sakata N, Domenico-Hahn J, Lucas JJ, Gelfand EW (1999) Quantitative and qualitative signals determine T-cell cycle entry and progression. Cell Immunol 197:19–29 Shapira MY, Tsirigotis P, Resnick IB, Or R, Abdul-Hai A, Slavin S (2007) Allogeneic hematopoietic stem cell transplantation in the elderly. Crit Rev Oncol Hematol 64:49–63 Petersen SL (2007) Alloreactivity as therapeutic principle in the treatment of hematologic malignancies. Studies of clinical and immunologic aspects of allogeneic hematopoietic cell transplantation with nonmyeloablative conditioning. Dan Med Bull 54:112–139 Vriz S, Lemaitre JM, Leibovici M, Thierry N, Mechali M (1992) Comparative analysis of the intracellular localization of c-Myc, c-Fos, and replicative proteins during cell cycle progression. Mol Cell Biol 12:3548–3555 Roux P, Blanchard JM, Fernandez A, Lamb N, Jeanteur P, Piechaczyk M (1990) Nuclear localization of c-Fos, but not v-Fos proteins, is controlled by extracellular signals. Cell 63:341–351 Hartenstein B, Teurich S, Hess J, Schenkel J, Schorpp-Kistner M, Angel P (2002) Th2 cell-specific cytokine expression and allergen-induced airway inflammation depend on JunB. EMBO J 21:6321–6329 Lwin WW, Park K, Wauson M, Gao Q, Finn PW, Perkins D, Khanna A (2012) Systems biology approach to transplant tolerance: proof of concept experiments using RNA Interference (RNAi) to knock down hub genes in Jurkat and HeLa Cells In Vitro. J Surg Res 176:e41–e46. doi:10.1016/j.jss.2011.12.002 Foletta VC, Segal DH, Cohen DR (1998) Transcriptional regulation in the immune system: all roads lead to AP-1. J Leukoc Biol 63:139–152 Hughes-Fulforda M, Suganoa TE, Schopperd T, Lia CF, Boonyaratanakornkita JB, Cogolid A (2005) Early immune response and regulation of IL-2 receptor subunits. Cell Signal 17:1111–1124 Tanaka H, Matsumura I, Ezoe S, Satoh Y, Sakamaki T, Albanese C, Machii T, Pestell RG, Kanakura Y (2002) E2F1 and c-Myc

123

20.

21.

22.

23.

24.

25.

26.

27.

28. 29.

30.

31.

32.

33.

34.

35.

36.

potentiate apoptosis through inhibition of NF-kappaB activity that facilitates MnSOD-mediated ROS elimination. Mol Cell 9:1017–1029 Shipp MA, Reinherz EL (1987) Differential expression of nuclear proto-oncogenes in T cells triggered with mitogenic and nonmitogenic T3 and T11 activation signals. J immunol 139:2143–2148 Dose M, Khan I, Guo Z, Kovalovsky D, Krueger A, Boehmer HV, Khazaie K, Gounari F (2006) c-Myc mediates pre-TCRinduced proliferation but not developmental progression. Blood 108:2669–2677 Wang R, Dillon CP, Zhichang SL, Milasta S, Carter R, Finkelstein D, McCormick LL, Fitzgerald P, Chi H, Munger J, Green DR (2011) The transcription Ffctor Myc controls metabolic reprogramming upon T Lymphocyte activation. Immunity 35:871–882 Lindsten T, June CH, Thompson CB (1988) Multiple mechanisms regulate c-myc gene expression during normal T cell activation. EMBO J 7:2787–2794 Qiao X, Pham DN, Luo H, Wu J (2010) Ran overexpression leads to diminished T Cell responses and selectively modulates nuclear levels of c-Jun and c-Fos. J Biol Chem 285:5488–5496. doi:10. 1074/jbc.M109.058024 Hughes CC, Pober JS (1993) Costimulation of peripheral blood T cell activation by human endothelial cells. Enhanced IL-2 transcription correlates with increased c-fos synthesis and increased Fos content of AP-1. J Immunol 150:3148–3160 Whisler RL, Chen M, Beiqing L, Carle KW (1997) Impaired induction of c-fos/c-jun genes and of transcriptional regulatory proteins binding distinct c-fos/c-jun promoter elements in activated human T cells during aging. Cell Immunol 175:41–50 Whisler RL, Beiqing L, Wu LC, Chen M (1993) Reduced activation of transcriptional factor AP-1 among peripheral blood T cells from elderly humans after PHA stimulation: restorative effect of phorbol diesters. Cell Immunol 152:96–109 Eferl R, Wagner EF (2003) AP-1: a double-edged sword in tumorigenesis. Nat Rev Cancer 3:859–868. doi:10.1038/nrc1209 Watanabe M, Nakajima S, Ohnuki K, Ogawa S, Yamashita M, Nakayama T, Murakami Y, Tanabe K, Abe R (2012) AP-1 is involved in ICOS gene expression downstream of TCR/CD28 and cytokine receptor signaling. Eur J Immunol 42:1850–1862 Horgan AF, Mendez MV, O’Riordain DS, Holzheimer RG, Mannick JA, Rodrick ML (1994) Altered gene transcription after burn injury results in depressed T-lymphocyte activation. Ann Surg 220:342–352 Walters ED, Drullinger LF, Kugel JF, Goodrich JA (2013) NFATc2 recruits cJun homodimers to an NFAT site to synergistically activate interleukin-2 transcription. Mol Immunol 56:48–56 Macia F, Garcıa-Cozar F, Sin-Hyeog I, Horton HF, Byrne MC, Rao A (2002) Transcriptional mechanisms underlying lymphocyte tolerance. Cell 109:719–731 Heisel O, Keown P (2001) Alteration in transcription factor binding at the IL-2 promotor region un anergized human CD4? T lymphocytes. Transplantation 72:1416–1422 Sottong PR, Rosebrock JA, Britz JA, Kramer TR (2000) Measurement of T-lymphocyte responses in whole-blood cultures using newly synthesized DNA and ATP. Clin Vaccine Immunol 7:307–311 White AG, Raju KT, Keddie S, Abouna GM (1989) Lymphocyte activation: changes in intracellular adenosine triphosphate and deoxyribonucleic acid synthesis. Immunol Lett 22:47–50 Lewin SR, Heller G, Zhang L, Rodrigues E, Skulsky E, van den Brink MR, Small TN, Kernan NA, O’Reilly RJ, Ho DD, Young JW (2002) Direct evidence for new T-cell generation by patients after either T-cell-depleted or unmodified allogeneic hematopoietic stem cell transplantations. Blood 100:2235–2242

Clin Exp Med 37. Selvatici R, Rubini M, Orlando P, Balboni A, Gandini E (1990) C-fos, c-myc and IL-2R mRNA expression in PHA activated T lymphocytes treated with a monoclonal anti-HLA class I antibody (MAb 01.65). Biochem Int 22:397–403 38. Ferrari S, Torelli U, Selleri L, Donelli A, Venturelli D, Narni F, Moretti L, Torelli G (1985) Study of the levels of expression of two oncogenes, c-myc and c-myb, in acute and chronic leukemias of both lymphoid and myeloid lineage. Leuk Res 9:833–842 39. Gamble DA, Schwab R, Weksler ME, Szabo P (1990) Decreased steady state c-myc mRNA in activated T cell cultures from old humans is caused by a smaller proportion of T cells that transcribe the c-myc gene. J Immunol 144:3569–3573 40. Song L, Stephens JM, Kittur S, Collins GD, Nagel JE, Pekala PH, Adler WH (1992) Expression of c-fos, c-jun and jun B in peripheral blood lymphocytes from young and elderly adults. Mech Ageing Dev 65:149–156 41. Shan X, Luo H, Chen H, Daloze P, St-Louis G, Wu J (1993) The effect of rapamycin on c-jun expression in human lymphocytes. Clin Immunol Immunopathol 69:314–317 42. DePalma L, Brown E, Baker R (1998) c-fos and c-jun mRNA expression in activated cord and adult lymphocytes: an analysis by Northern hybridization. Vox Sang 75:134–138 43. King LB, Tolosa E, Lenczowski JM, Lu F, Lind EF, Hunziker R, Petrie HT, Ashwell JD (1999) A dominant-negative mutant of c-Jun inhibits cell cycle progression during the transition of CD4(-)CD8(-) to CD4(?)CD8(?) thymocytes. Int Immunol 11:1203–1216 44. Trop-Steinberg S, Azar Y, Or R (2013) Early cell-cycle gene expression in T-cells after hematopoietic stem cell transplantation. Transpl Immunol 29:146–154. doi:10.1016/j.trim.2013.03. 002 45. Kern JA, Reed JC, Daniele RP, Nowell PC (1986) The role of the accessory cell in mitogen-stimulated human T cell gene expression. J Immunol 137:764–769

46. Kowalski RJ, Mannon RB et al (2006) Assessing relative risks of infection and rejection: a meta-analysis using an immune function assay. Transplantation 82:663 47. Kowalski R, Post D, Schneider MC, Britz J, Thomas J, Deierhoi M, Lobashevsky A, Redfield R, Schweitzer E, Heredia A, Reardon E, Davis C, Bentlejewski C, Fung J, Shapiro R, Zeevi A (2003) Immune cell function testing: an adjunct to therapeutic drug monitoring in transplant patient management. Clin Transplant 17:77–88 48. Sanchez-Velasco P, Rodrigo E, Valero R, Ruiz JC, FernandezFresnedo G, Lopez-Hoyos M, Pinera C, Palomar R, LeyvaCobian F, Arias M (2008) Intracellular ATP concentrations of CD4 cells in kidney transplant patients with and without infection. Clin Transplant 22:55–60 49. Gesundheit B, Budowski E, Israeli M, Shapira MY, Resnick IB, Bringer R, Azar Y, Samuel S, Dray L, Amar A, Kristt D, Or R (2010) Assessment of CD4 T-lymphocyte reactivity by the Cylex ImmuKnow assay in patients following allogeneic hematopoietic SCT. Bone Marrow Transpl 45:527–533 50. Sayegh MH, Turka LA (1998) The role of T-cell costimulatory activation pathways in transplant rejection. N Engl J Med 338:1813–1821 51. Nguyen TN, Kim LJ, Walters RD, Drullinger LF, Lively TN, Kugel JF, Goodrich JA (2010) The C-terminal region of human NFATc2 binds cJun to synergistically activate interleukin-2 transcription. Mol Immunol 47:2314–2322 52. Naito T, Tanaka H, Naoe Y, Taniuchi I (2011) Transcriptional control of T-cell development. Int Immunol 23:661–668 53. Katoh M, Igarashi M, Fukuda H, Nakagama H (2013) Cancer genetics and genomics of human FOX family genes. Cancer Lett 328:198–206

123