Accumulating evidence suggests that a dysregulation in cytokines produced by T-helper (TH) cells is involved in the etiology and progression of inflammatory bowel disease (IBD) (1); however, the underlying immunological correlates of disease pathophysiology remain poorly understood. The 2 main subclasses of TH cells, T-helper 1 (TH1) and T-helper 2 (TH2) cells, secrete subsets of cytokines that mediate various immunological responses (2). TH1 cells secrete the cytokines interleukin (IL)-2, interferon (IFN)-γ, and tumor necrosis factor (TNF)-α to augment an immune response against intracellular pathogens such as viruses. In contrast, TH2 cells secrete IL-4, IL-5, IL-10, and IL-13 (3), mainly to protect the host against extracellular pathogens and promote the synthesis of IgE antibodies in B cells (4). IL-4 is a signature TH2 cytokine responsible for the development of naïve TH cells into a TH2 functional phenotype. IL-6 is a pleiotropic cytokine responsible for T-cell stimulation and proliferation (5) and is primarily secreted by macrophages and monocytes promoting granulocyte and macrophage colony formation (6).
Traditionally, Crohn disease (CD) has been considered a TH1-mediated disease. In many previous studies, cytokine profiles of patients with CD displayed TH1 dominance characterized by higher than normal percentages of IFN-γ and/or lower or undetectable percentages of cells expressing IL-4 in peripheral blood mononuclear cells or intestinal mucosa (2,7–12). Higher TNF-α has also been reported in the intestinal mucosa and serum of patients with CD compared with healthy controls (13–16). A shift toward the TH1 profile may contribute to the onset of CD by activating macrophages that produce proinflammatory cytokines, thereby leading to the intestinal damage seen in CD (17). Recent therapies have targeted these cytokines for CD management, as exemplified by the administration of anti-TNF-α-depleting antibodies (18).
The role of CD4+CD45RO+ T cells in disease progression has been described in patients with CD (19,20). In the mature human immune system, circulating CD4+ T cells can be separated into naïve and memory T cells by their expression of CD45. Naïve T cells express CD45RA and show little or no ability to produce effector cytokines while circulating between the peripheral blood and secondary lymphoid organs. In contrast, memory T cells express CD45RO and, when activated, proliferate in an antigen-specific manner (19,21,22). CD4+CD45RO+ T cells are known for their ability to secrete large quantities of cytokines and their tendency to accumulate at sites of inflammation (eg, luminal surfaces in IBD). Because of these characteristics, it is reasonable to propose that the activity of CD4+CD45RO+ T cells is implicated in the pathogenesis of CD; however, few studies have investigated the association between CD and cytokine production of CD4+CD45RO+ T cells. During childhood the immune system's repertoire is expanded by exposure to antigens. This constant immune expansion may account for some of the differences in cytokine profiles observed between children and adults (23). Therefore, we explored the hypothesis that age- and disease-related cytokine profiles in children differ from adults with CD.
Our prior investigation of cytokine profiles in pediatric patients with CD suggested that lower TH1 cytokine levels were present in newly diagnosed children with CD compared with control subjects (24). In the present pilot study, we aimed to validate this finding and expand the spectrum of cytokines and age of patients with CD and controls to include adults. The intracellular production of IFN-γ, TNF-α, IL-4, and IL-6, cytokines that have been associated with various autoinflammatory conditions including IBD, were analyzed in whole blood by flow cytometry.
Study Population and Sample Collection
The present study was conducted in collaboration between the School of Public Health, University of California, Berkeley (UCB), and the Departments of Pediatrics and Medicine, University of California, San Francisco (UCSF). Written informed consent was obtained from the subjects of legal age or their parents/legal guardians, and assent was obtained from younger subjects in accordance with the protocol approved by the committee on human research at UCSF, and the committee for the protection of human subjects at UCB.
Ten children and 10 adults were enrolled at UCSF from July 2008 to May 2009. Five subjects in each age group diagnosed as having CD and an equal number of healthy controls were enrolled. All of the subjects were approached as they presented for endoscopic procedures, and pediatric subjects and controls were matched for age. Inclusion criteria for controls were no preexisting acute or chronic illnesses or inflammatory disorders. At the time of enrollment, most patients with CD had some symptoms and others were asymptomatic. Body mass index (BMI; weight (kg)/height (m)2) was determined by medical record review or patient report. Laboratory analyses included measurement of red blood cell folate levels and hematocrit. Samples were obtained from all of the subjects in a fasting state. Peripheral blood samples were kept at a cool temperature with ice packs during transfer and delivered to the laboratory at UCB for processing within 4 hours of collection.
Whole-blood samples were diluted 1:1 with culture media (RPMI 1640) in a 12 × 75 mm fluorescence-activated cell sorting tubes. Phorbol-12-myristate 13-acetate (2.5 ng/mL; Sigma Chemical Co, St Louis, MO) and ionomycin (1 μg/mL, Sigma Chemical) were added for lymphocyte activation. Cultures were incubated in the presence of brefeldin-A (10 ng/mL, Sigma Chemical), a transport inhibitor that prevents cytokine release from cells, at 37°C and 5% CO2 for 4 hours.
Following the incubation, 200 μL of whole blood was pipetted into a 12 × 75 mm fluorescence-activated cell sorting tube containing monoclonal antibodies specific for TH cell surface antigen CD4 (CD4-PerCP, Becton Dickinson, San Diego, CA). For CD4+CD45RO+ T cells and their cytokine measurement, incubated blood samples were added to tubes containing monoclonal antibodies for CD4 (CD4-fluorescein isothiocyanate [FITC], Becton Dickinson) and monoclonal antibodies for CD45RO (CD45RO-phycoerythrin [PE]-Cy 5, Becton Dickinson). After a 10-minute incubation at room temperature in the dark, 1% paraformaldehyde was added to stabilize the monoclonal antibody–surface antigen complex for 5 minutes. Red blood cells present in the mixture were then lysed using 3 mL of 1× lysing solution (Becton Dickinson) for 8 minutes. Following centrifugation at 1930 rpm for 5 minutes, the supernatant was aspirated, with 1× permeabilizing solution added to the pellet, and incubated for 10 minutes at room temperature in the dark. The mixture was then washed using 3 mL of wash buffer (1% bovine serum albumin, 0.1% NaN3, 1× phosphate buffered saline) and centrifuged for 5 minutes at 2370 rpm. After aspirating the supernatant, 20 μL of TH1 and TH2 cytokine-specific antibodies (IFN-γ-FITC/IL-4-PE, Becton Dickinson; TNF-α-FITC/IL-6-PE, Becton Dickinson) were added to the experimental tubes and the appropriate isotype control monoclonal antibodies (20 μL, immunoglobulin [Ig] G2/IgG1, Becton Dickinson) were added to the control tubes and incubated for 30 minutes at room temperature in the dark. To identify cytokine-producing CD4+CD45RO+ cells, the samples were stained with 20 μL of monoclonal antibodies specific for each cytokine (IFN-γ-PE; IL-4-PE; TNF-α-PE; IL-6-PE, Becton Dickinson) in separate tubes and appropriate isotype control antibodies were used. Subsequently, the tubes were washed with 3 mL of the buffer for 3 minutes and centrifuged at 2520 rpm for 5 minutes. Finally, after aspirating the supernatant, the cells were resuspended in 500 μL of 1% paraformaldehyde and stored at 4°C until flow cytometry analysis.
Cytokine Level Detection by Flow Cytometry
The stained cells were detected by Becton-Coulter EPICS XL flow cytometer (Becton Dickinson) as previously described and illustrated (24) to calculate the percentage of TH1- and TH2-positive cells from CD4+ and CD4+CD45RO+ cells.
For each sample, control tubes containing fluorochrome-equivalent IgG2/IgG1 isotype controls were run to detect non-specific binding, followed by runs of the experimental tubes containing IFN-γ/IL-4- and TNF-α/IL-6-specific antibodies. During each run of the sample, 5000 CD4+ cells were counted. The lymphocyte population was marked by placing a circular gate around the dense aggregation of lymphocytes on the forward-scatter/side-scatter histogram based on the size and granularity of the cells. A 1-plot histogram was used to identify CD4+ cells by placing a linear gate on the right peak of the histogram. The percentage of CD4+ cells was calculated as the number of CD4+ cells over the total lymphocyte population. TH1 cells and TH2 cells were identified by using a 2-plot histogram, in which the IFN-γ- or TNF-α-positive cells were found in the lower right quadrant, and the IL-4- or IL-6-positive cells were found in the upper left quadrant. The percentage of TH1-positive cells was determined by dividing the number of IFN-γ- or TNF-α-positive cells by CD4-positive cells, and the percentage of TH2-positive cells was determined by dividing the number of IL-4- or IL-6-positive cells by CD4-positive cells.
For each sample, isotype-matched FITC, PE-Cy5, and PE-labeled monoclonal antibodies were used as controls. Upon gating for the lymphocyte population in the forward-scatter/side-scatter histogram, a 2-plot histogram was used to select 5000 CD4+CD45RO+ cells in the upper right quadrant. A 1-plot histogram was used to identify IFN-γ-, IL-4-, TNF-α-, and IL-6-positive cells in separate tubes. The percentage of CD4+CD45RO+ cells was calculated as the number of CD4+CD45RO+ cells over the total lymphocyte population. The percentage of cytokine-producing cells was determined by dividing the number of IFN-γ, IL-4, TNF-α, or IL-6 cells by the number of CD4+CD45RO+ cells. Flow cytometry results are presented as percentages of cells.
Statistical analyses were performed using STATA 11.0 (StataCorp, College Station, TX). Data were analyzed using Mann-Whitney and Kruskal-Wallis tests to compare subjects with CD with subjects without CD and adults versus children. Because of the small sample size, data are expressed as both mean with standard deviation and median with range. Values of P < 0.05 were considered statistically significant.
Demographic characteristics and clinical parameters of the study participants are shown in Tables 1 and 2. The median age of adult patients with CD was slightly lower compared with healthy controls, but the difference was not statistically significant (P = 0.14). Children ranged from 9 to 15 years old.
No differences were observed in BMI levels between healthy adults and children and between adults and children with CD. All 4 of the pediatric subjects with BMI <18.5 were still within the normal range for their age (lowest z score was −1.6 and highest was 0.5). No differences were found between patients with CD and healthy controls by age, sex, BMI, folate levels, or hematocrit.
Effects of CD on Cytokine Profiles
The distributions of cytokine profiles (CD4+, CD4+CD45RO+, IFN-γ, TNF-α, IL-4, IL-6) by age and disease status are shown in Table 3.
CD4+ T Cells
No differences were found in the frequency of CD4+ cells between patients with CD and healthy controls in both adult and pediatric groups. TNF-α analyses revealed that pediatric patients with CD had higher frequencies of TNF-α (23.3% ± 7.6%) compared with the controls (11.3% ± 2.2%; P = 0.009) (Fig. 1B). Similarly, the percentages of TNF-α were increased in adult patients with CD (30.5% ± 10.6%) compared with controls (16.5% ± 4.9%; P = 0.047). Percentages of cells expressing IL-4 were also higher in pediatric patients with CD (0.4% ± 0.2%) than in pediatric controls (0.1% ± 0.1%; P = 0.036, whereas IFN-γ-expressing cells were less common in pediatric patients with CD (4.66% ± 0.9%) compared with pediatric controls (9.1% ± 4.4%; P = 0.009) (Fig. 1A and C)). In our previous study, newly diagnosed pediatric patients with CD also possessed lower IFN-γ compared with healthy controls, but the IL-4 did not differ significantly between the 2 groups (24). In the present study, IL-6 percentages did not differ between the 2 pediatric groups. No differences in IFN-γ, IL-4, and IL-6 percentages were found between adult patients with CD and controls.
Noteworthy is that TNF-α expression was more prevalent in the patients whether they were being treated with anti-TNF medications compared with the control group subjects, independent of age (Fig. 2).
CD4+CD45RO+ T Cells
When only the memory T-cell population cytokine profile was considered, no significant difference was found in CD4+CD45RO+ cells between patients with CD and controls in both age groups; however, pediatric patients with CD had decreased percentages of IFN-γ cells (7.1% ± 0.9%) compared with controls (16.4% ± 7.0%; P = 0.009. No difference was found in TNF-α, IL-4, and IL-6 between patients with CD and controls in both the adult and pediatric groups.
Effects of Age on Cytokine Profiles
The effect of age on cytokine profiles was investigated by comparing the CD4+, CD4+CD45RO+, TH1, and TH2 in children and adults who were either patients with CD or controls.
CD4+ T Cells
Among patients with CD, the percentage of CD4+CD45RO+, IFN-γ+ cells was >3-fold higher in adults (15.1% ± 3.8%) than in pediatric subjects (4.6% ± 0.9%; P = 0.009) (Fig. 1A). In contrast, IL-4 percentages were lower in adults (0.17% ± 0.1%) compared with pediatric subjects (0.4% ± 0.2%; P = 0.047) (Fig. 1C). No significant differences in the CD4+, TNF-α, and IL-6 were observed between the 2 age groups. Among healthy controls, the percentages of TH1 cells (IFN-γ and TNF-α) were slightly higher in adults and no differences in CD4+, IL-4, and IL-6 were observed.
CD4+CD45RO+ T Cells
The comparison of CD4+CD45RO+ T cell between pediatric (8.1% ± 1.6%) and adult patients with CD (14.1% ± 3.5%) showed a notable increase with age (P = 0.009) (Fig. 1D). In addition, percentages of CD4+CD45RO+IFN-γ+ cells were significantly higher in adult patients with CD (14.2% ± 5.8%) than in pediatric patients with CD (7.1% ± 0.9%; P = 0.016). No differences were observed for TNF-α-, IL-4-, and IL-6-expressing CD4+CD45RO+ cells between the 2 age groups.
We measured the intracellular production of IFN-γ, TNF-α, IL-4, and IL-6 from CD4+ and CD4+CD45RO+ T cells in the peripheral blood of adult and pediatric patients with CD and age-matched controls. To distinguish between the effect or capabilities of the CD4+CD45RO+ memory T cells and the broader class of CD4+ T cells, we obtained a separate set of flow cytometric data showing cytokine expression in the CD4+CD45RO+ memory T-cell compartment and compared the profiles of patients with CD to the controls in both age groups.
The mean IFN-γ levels from CD4+ T cells were significantly lower in pediatric patients with CD than in controls. This supports our previous observation as well as other published data in treatment-naïve pediatric patients with CD (24,25). These results, however, are in contrast to findings from other studies that have evaluated the TH1/TH2 profiles in the intestinal mucosa and/or peripheral blood associated with CD. In adults, increased expression of IFN-γ was found in the peripheral blood or intestinal tissue of adult patients with CD compared with controls, in line with the accepted dogma of a TH1-mediated mechanism of disease (7,9,26). Similar to our study, IL-4 production has been reported to be inconsistent among patients with CD.
We found that TNF-α expression was increased in pediatric patients with CD, whereas levels of IFN-γ were decreased. Secretion of proinflammatory cytokines such as TNF-α in patients with CD is triggered by the TH1 response originating from an increase in IFN-γ levels from lymphocytes and macrophages and leads to continual immune activation. Because this aberrant immune activation contributes to inflammation in IBD (27), it is puzzling that the peripheral levels of TNF-α in our children with CD were elevated despite an apparent downregulation of a TH1 response (ie, decreased levels of IFN-γ and increased IL-4 levels). This observation may result from our relatively small number of subjects; however, the findings are consistent with previous reports (24,25).
IL-6 has been associated with CD because of its capability to induce synthesis of acute phase proteins and inflammatory chemical pathways (6). In the present study, however, we did not find any significant differences in the intracellular production of IL-6 between patients with CD and controls in both age groups. Previous studies suggest that adult patients with CD have higher serum or plasma levels of IL-6 compared with controls (6,27). In our study, production of IL-6 was measured only from circulating CD4+ T cells and their subsets. The predominant sources of IL-6 in patients with IBD during inflammation are activated monocytes, macrophages, and, to a lesser extent, epithelial cells (5). Our data may provide evidence that peripheral CD4+ T-cell production of IL-6 is not a determining factor in the pathogenesis of pediatric and adult CD. It is necessary to evaluate IL-6 expression from other cell types to gain a better understanding of its role in CD pathogenesis.
Our analyses reveal that intracellular production of IFN-γ by CD4+CD45RO+ cells is significantly reduced in pediatric patients with CD compared with healthy controls, reflecting the general trend seen in the overall CD4+ T cell population. Thus, it is likely that a downregulation in the activity of circulating memory TH1 cells and their production of IFN-γ occurs in pediatric patients with CD; however, no differences were seen in the intracellular production of TNF-α, IL-4, and IL-6 from CD4+CD45RO+ T cells between the patients with CD and controls in either age group. Previously, De Tena et al reported that the percentages of CD4+CD45RO+ T cells producing TNF-α and IL-6 were higher in adults with active CD than in healthy controls (19). The intracellular production of IFN-γ was also greater in the active CD group compared with controls, although not statistically significant (19). These results are discordant with our findings in both age groups; however, the range and mean of ages of the study participants and the methods of cytokine analysis in the 2 studies were different. Furthermore, the clinical status of remission in most of our patients may have led to a reduction in the effector capabilities of circulating memory T cells to produce cytokines in our patients with CD. An investigation of memory T cells in the intestinal mucosa of pediatric and adult patients with CD may help elucidate the true function of CD4+CD45RO+ T cells in the immunodysregulation associated with CD.
Earlier studies examined the relation between age and cytokine profiles throughout development (23,28); however, with the exception of our previous assessment in children newly diagnosed as having CD, this is the first study to evaluate peripheral cytokine levels with age in patients with CD. We previously reported that although CD4+ T cell frequencies remain constant among pediatric patients with CD and controls with increasing age, the percentage of TH1 cells steadily increases with age in both groups (24). Similarly, in the present study the overall production of TH1 cytokines from CD4+ T cells was generally higher in healthy adults compared with pediatric subjects. Our findings corroborate earlier reports of an increase in TH1 cells with age (23,28,29).
We found that the percentages of CD4+CD45RO+ T cells increased with age in patients with CD. In contrast, no difference was noted between the groups of healthy children and adults. Only a few studies have followed the changes in expression of circulating CD4+CD45RO+ population with age in healthy subjects (30,31).
In summary, our study demonstrates increased production of the proinflammatory cytokine TNF-α in patients with CD in both pediatric and adult groups. Our clinical data also support differences in disease phenotype between adult and pediatric patients with IBD (32). The results suggest that fundamental differences exist between adults and children in the immunological mechanisms of CD pathogenesis. In addition, age can be an important determining factor in cytokine regulation in both healthy and CD-diagnosed subjects. Although our study was limited by sample size and subjects’ heterogeneity, we still detected several statistically significant differences in cytokine levels between groups. It should be noted that functional cytokine analyses were performed after in vitro stimulation with phorbol-12-myristate 13-acetate and ionomycin, which may overstimulate a traditional in vivo process. Furthermore, not all of our patients with CD were newly diagnosed as having the disease, and some had been receiving medical therapy before sample collection, raising the possibility of cytokine profile alterations caused by drug therapy. Nevertheless, the data obtained may help to elucidate immunological changes in patients with CD of varying ages.
Although these investigations have shed light on the peripheral blood cytokine profiles of pediatric and adult patients with CD, further research is necessary. Ex vivo analyses and in situ measurement of mucosal tissue must be performed to determine whether peripheral cytokine profiles correlate with disease activity. Larger sample sizes with a broader age distribution would be ideal to determine whether a correlation does exist, in both diseased and nondiseased patients between age and percentage of TH1 or TH2 secreting CD4+ T cells. Additional analyses on the TH17 subset (eg, IL-17, IL-23, IL-18, transforming growth factor-β) should be included in future analyses.
We are grateful to the laboratory and clinical staff and participants of the study for all of their contributions. The helpful comments of Dr. Karen Huen and Vitaly Volberg are appreciated.
1. Brown SJ, Mayer L. The immune response in inflammatory bowel disease. Am J Gastroenterol 2007; 102:2058–2069.
2. Parronchi P, Romagnani P, Annunziato F, et al. Type 1 T-helper cell predominance and interleukin-12 in the gut of patients with Crohn's disease. Am J Clin Pathol 1997; 150:823–832.
3. Farrar JD, Asnagli H, Murphy KM. T-helper subset development: role of instruction, selection, and transcription. J Clin Invest 2002; 109:431–435.
4. Kidd P. Th1/Th2 balance: the hypothesis, its limitations, and implications for health and disease. Altern Med Rev 2003; 8:223–246.
5. Brown KA, Back SJ, Ruchelli ED, et al. Lamina propria and circulating interleukin-6 in newly diagnosed pediatric inflammatory bowel disease patients. Am J Gastroenterol 2002; 97:2603–2608.
6. Mahida YR, Kurlac L, Gallagher A, et al. High circulating concentrations of interleukin-6 in active Crohn's disease but not ulcerative colitis. Gut 1991; 32:1531–1534.
7. Niessner M, Volk BA. Altered Th1/Th2 cytokine profiles in the intestinal mucosa of patients with inflammatory bowel disease as assessed by quantitative reversed transcribed polymerase chain reaction (RT-PCR). Clin Exp Immunol 1995; 101:428–435.
8. Brandonisio O, Panaro MA, Sisto M, et al. Nitric oxide production by Leishmania-infected macrophages and modulation by cytokines and prostaglandins. Parassitologia 2001; 43 (suppl):1–6.
9. Camoglio L, Te Velde AA, Tigges AJ, et al. Altered expression of interferon-γ and interleukin-4 in inflammatory bowel disease. Inflamm Bowel Dis 1998; 4:285–290.
10. West GA, Matsuura T, Levine AD, et al. Interleukin 4 in inflammatory bowel disease and mucosal immune reactivity. Gastroenterology 1996; 110:1683–1695.
11. Mariani P, Bachetoni A, D’Alessandro M, et al. Effector Th-1 cells with cytotoxic function in the intestinal lamina propria of patients with Crohn's disease. Dig Dis Sci 2000; 45:2029–2035.
12. Karttunnen R, Breese EJ, Walker-Smith JA, et al. Decreased mucosal interleukin-4 (IL-4) production in gut inflammation. J Clin Pathol 1994; 47:1015–1018.
13. Reimund JM, Wittersheim C, Dumont S, et al. Increased production of tumour necrosis factor-a, interleukin-1B, and interleukin-6 by morphologically normal intestinal biopsies from patients with Crohn's disease. Gut 1996; 39:684–689.
14. MacDonald TT, Hutchings P, Choy MY, et al. Tumour necrosis factor-alpha and interferon-gamma production measured at the single cell level in normal and inflamed human intestine. Clin Exp Immunol 1990; 81:301–305.
15. Breese EJ, Michie CA, Nicholls SW, et al. Tumor necrosis factor-alpha producing cells in the intestinal mucosa of children with inflammatory bowel disease. Gastroenterology 1994; 106:1455–1466.
16. Murch SH, Lamkin VA, Savage MO, et al. Serum concentrations of tumour necrosis factor a in childhood chronic inflammatory bowel disease. Gut 1991; 32:913–917.
17. Romagnani P, Annunziato F, Baccari MC, et al. T cells and cytokines in Crohn's disease. Immunology 1997; 9:793–799.
18. Diamanti A, Basso MS, Gambarara M, et al. Positive impact of blocking tumor necrosis factor alpha on the nutritional status in pediatric Crohn's disease patients. Int J Colorectal Dis 2009; 24:19–25.
19. De Tena JG, Manzano L, Leal JC, et al. Distinctive pattern of cytokine production and adhesion molecule expression in peripheral blood memory CD4+ T cells from patients with active Crohn's disease. J Clin Immunol 2006; 26:233–242.
20. Roman LI, Manzano L, De La Hera A, et al. Expanded CD4+CD45RO+ phenotype and defective proliferative response in T lymphocytes from patients with Crohn's disease. Gastroenterology 1996; 110:1008–1019.
21. De Tena JG, Manzano L, Leal JC, et al. Active Crohn's disease patients show a distinctive expansion of circulating memory CD4+CD45RO+CD28null T cells. J Clin Immunol 2004; 24:185–196.
22. Wysocka J, Hassmann E, Lipska A, et al. Naïve and memory T cells in hypertrophied adenoids in children according to age. Int J Pediatr Otorhinolaryngol 2003; 67:237–241.
23. Hartel C, Adam N, Strunk T, et al. Cytokine responses correlate differentially with age in infancy and early childhood. Clin Exp Immunol 2005; 142:446–453.
24. Holland N, Dong J, Garnett EA, et al. Reduced intracellular T-helper 1 interferon gamma in blood of newly diagnosed children with Crohn's disease and age-related changes in TH1/TH2 cytokine profiles. Pediatr Res 2008; 63:257–262.
25. Mack DR, Beedle S, Warren J, et al. Peripheral blood intracellular cytokine analysis in children newly diagnosed with inflammatory bowel disease. Pediatr Res 2002; 51:328–332.
26. Autschbach F, Giese T, Gassler N, et al. Cytokine/chemokine messenger-RNA expression profiles in ulcerative colitis and Crohn's disease. Virchows Arch 2002; 441:500–513.
27. Szkaradkiewicz A, Marciniak R, Chudzicka-Strugala I, et al. Proinflammatory cytokines and IL-10 in inflammatory bowel disease and colorectal cancer patients. Arch Immunol Ther Exp 2009; 57:291–294.
28. Kawamoto N, Kaneko H, Takemura M, et al. Age-related changes in intracellular cytokine profiles and Th2 dominance in allergic children. Pediatr Allergy Immunol 2006; 17:125–133.
29. Duramad P, McMahon CW, Hubbard A, et al. Flow cytometric detection of intracellular TH1/Th2 cytokines using whole blood: validation of immunologic biomarker for use in epidemiologic studies. Cancer Epidemiol Biomarkers Prev 2004; 13:1452–1458.
30. Falcao RP, De-Santis GC. Age-associated changes of memory (CD45RO+) and naïve (CD45R+) T cells. Braz J Biol Res 1991; 24:275–279.
31. Aldhous MC, Raab GM, Doherty KV, et al. Age-related ranges of memory, activation, and cytotoxic markers on CD4 and CD8 cells in children. J Clin Immunol 1994; 14:289–299.
32. Heyman MB, Kirschner BS, Gold BD, et al. Children with early-onset inflammatory bowel disease (IBD): analysis of a pediatric IBD consortium registry. J Pediatr 2005; 146:35–40.