Sepsis is characterized by the coexistence of both hyperinflammatory and hypoinflammatory responses to pathogens (1). Recent clinical studies primarily sought to influence the hyperinflammatory pathway using different pharmacological approaches to decrease proinflammatory mediators (2). But suppression of the hyperinflammatory response reduces the host's ability to fight primary or secondary infections, and therefore this treatment regimen failed to significantly improve septic survival (3–5).
However, the concomitant release of anti-inflammatory mediators, as a constituent aspect of the hypoinflammatory response, provokes T cell apoptosis (6). This excessive T cell death produces immune paralysis, which is thought to be the main reason for the inability of the septic patient to fight against infections (7) that correlates with sepsis mortality (3,5,8,9).
Peroxisome proliferator-activated receptor gamma (PPARγ) causes one of these anti-inflammatory responses (10–12) and acts as a key regulator in apoptosis of activated T cells during sepsis (6). The transcription factor heterodimerizes with the retinoid X receptor protein and activates target genes by binding to PPAR response elements (3,10). With regard to sepsis, it is of interest that PPARγ binds to other transcription factors or coactivators, thus preventing their binding to their responsive site(s) in promoter/enhancer regions of mainly proinflammatory target genes. Consequently, in a mouse polymicrobial sepsis model, genetic deletion of PPARγ in T cells or its pharmacological antagonism improves survival by preventing T cell apoptosis, consequently reducing immune paralysis (3). As a potential mechanism causing T cell apoptosis during sepsis, PPARγ-dependent induction of its target gene PTEN emerged. The phosphatase PTEN is known to counteract prosurvival PI3K and Akt signaling, thereby provoking a proapoptotic pathway (3,13). In addition, PPARγ inhibits interleukin (IL)-2 expression and downstream antiapoptotic Bcl-2 signaling by scavenging NFAT, a transcription factor essential for IL-2 expression and concomitant T cell survival (3,14). In line with these in vivo animal data, T cells derived from septic patients were found to be more prone to PPARγ-dependent apoptosis following exposure to PPARγ agonists ex vivo compared with T cells isolated from the blood of healthy donors (6). Moreover, an initial study determining PPARγ expression in T cells from septic patients revealed higher expression of PPARγ in these cells compared with T cells from healthy volunteers (6,12).
The pilot study described here was performed to elucidate whether T cell PPARγ expression is regulated during sepsis progression and whether it can be used as a prognostic factor in human septic patients. To answer these questions, we determined the T cell count and T cell-specific PPARγ expression in T cells derived from blood of septic patients. Sampling started on the day when sepsis was diagnosed (= day 0) and then, consecutively, up to day 14.
MATERIALS AND METHODS
We received 9 mL blood in EDTA tubes from each patient with septic symptoms (n = 18) treated on the ICU of the University Hospital Frankfurt. As inclusion criteria for healthy donors, people aged between 18 and 75 years were included and for sepsis samples intensive care-treated patients aged between 18 and 75 years were involved. Exclusion criteria were not defined. After diagnosing a septic syndrome, as defined in (15), blood was harvested from each patient by the medical staff at days 0, 1, 2, 4, 6, and 14. Blood samples from healthy donors (n = 11) were used as controls. The study was approved by the ethics committee of the University Hospital Frankfurt, Germany (F199/13).
T cell enrichment
Following erythrolysis, 100 μL human CD3 microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany) were added and incubated as described by the manufacturer. CD3+, CD4+, and CD8+ T cells were isolated using automatic magnetic activated cell sorting (AutoMACS) technology (Miltenyi Biotec). Purity of each subpopulation was above 98% and verified by flow cytometric analysis (Supplemental Figure 1, Supplemental Digital Content 1, http://links.lww.com/SHK/A360).
RNA extraction and reverse transcriptase quantitative real-time PCR
After isolation of T cell subpopulations (CD3+, CD4+, CD8+), total RNA was isolated using the RNeasy Mini Kit (Qiagen, Hilden, Germany) according to the distributor's manual. Reverse transcription of mRNA was performed with the Maxima First Standard cDNA synthesis kit (Thermo Fischer Scientifics, Schwerte, Germany). The reverse transcriptase quantitative real-time (RT-qPCR) was performed using the CFX real-time PCR system (Bio-Rad Laboratories GmbH, Munich, Germany) and the Absolute iQ SYBR Green Supermix (Bio-Rad Laboratories GmbH) according to the manufacturer's instructions. Validated primer sets obtained from Qiagen were used to amplify human PPARγ (NM_138712), human IL-2 (NM_000586), and human PTEN (NM_000314). A primer pair to assess 18S rRNA (NR_003286; forward 5[Combining Acute Accent]-GTA ACC CGT TGA ACC CCA TT-3[Combining Acute Accent], reverse 5[Combining Acute Accent]-CCA TCC AAT CGG TAG TAG CG-3[Combining Acute Accent]) as a housekeeping gene for normalization was provided by biomers.net GmbH (Ulm, Germany). PPARγ mRNA expression was quantified by a PPARγ plasmid standard (pSEW-hPPARγ1 WT-DsRed Monomer C1, map is provided in Supplemental Figure 2, Supplemental Digital Content 1, http://links.lww.com/SHK/A360) in each of the performed RT-qPCR reactions using the PPARγ-specific primer pair as described above. With this standard, the absolute rate of PPARγ copies was determined. A standard curve is provided in Supplemental Figure 3, Supplemental Digital Content 1, http://links.lww.com/SHK/A360.
Analysis of T cells
T cell counts were directly determined with a BD LSR Fortessa Cell Analyzer (Becton Dickinson, Heidelberg, Germany). Fc receptor binding was blocked by CD16/CD32 antimouse antibody for 15 min on ice followed by 20 min incubations with anti-CD3e-PE-CF594, anti-CD4-AmCyan, and anti-CD8-APC-Cy7 on ice. Afterward, erythrocytes were lysed, and the number of stained cells was examined by flow cytometry. The gating strategy is exemplified for CD3+T cells in Supplemental Figure 4, Supplemental Digital Content 1, http://links.lww.com/SHK/A360.
We used two-tailed statistical analysis to assess the data. For statistical analysis of intergroup comparisons, we used the unpaired Student t test. Differences were considered significant when: *P < 0.05; **P < 0.01; ***P < 0.001. To estimate a significant difference in PPARγ mRNA expression in T cells of healthy donors versus sepsis patients, we performed a ROC curve analysis (Supplemental Figure 5, Supplemental Digital Content 1, http://links.lww.com/SHK/A360).
T lymphocyte count in septic patients
Our study protocol is provided in Figure 1. As a first approach and in extension of previous studies, we analyzed blood of 17 patients at day 0 (= day of sepsis diagnosis), and then on the consecutive days 1 (18 patients), 2 (17 patients), 4 (12 patients), 6 (14 patients), and 14 (7 patients), following the initial diagnosis, to determine the number of CD3+ T cells. Patient data are provided in Table 1. As shown in Figure 2A and Table 2 (second row), T cell counts were significantly decreased on each evaluation day compared with healthy donors. The lowest CD3+ T cell number was observed on day 1 after the diagnosis of sepsis (day 0: 277 ± 65 vs. day 1: 218 ± 32 CD3+ T cells/μL), followed by a continuous rise up to day 14. A statistical difference was only found when analyzing day 1 in comparison with day 14 (day 1: 218 ± 32 vs. day 14: 525 ± 248 CD3+ T cells/μL). Thus, our data suggest that by approximately day 14 the number of CD3+ T cells recovered. However, the T cell number at this time point was still significantly reduced compared with healthy donors (1,803 ± 134 vs. 524 ± 248 CD3+ T cells/μL). To assess whether the transcription factor PPARγ correlates with CD3+ T cell demise, we analyzed PPARγ mRNA expression in enriched T cells.
PPARγ expression in T cells derived from septic patients
To follow the time course of PPARγ expression in CD3+ T cells derived from blood of septic patients, we enriched CD3+ T cells from blood on day 0 (= day of sepsis diagnosis) and then on successive days 1, 2, 4, 6, and 14. Analyzing enriched CD3+ T cells for PPARγ mRNA expression by qPCR, we observed the highest PPARγ expression on the day of sepsis diagnosis (i.e., day 0), with a continuously slight decrease on each of the subsequent days, but this was still significantly higher on day 14 compared with healthy controls (Fig. 2B and Table 2, first row). Because PPARγ is known to inhibit IL-2 expression by blocking NFAT binding to the IL-2 promoter, we examined IL-2 expression in CD3+ T cells from septic patients and healthy donors. Indeed, high PPARγ expression mirrored the low IL-2 expression (Fig. 2C).
Inverse correlation of CD3+ T cell count and PPARγ expression
Combining the results of the CD3+ T cell count with the corresponding expression of PPARγ, we found that both parameters are inversely correlated (Fig. 2D). Whereas the ratio of PPARγ mRNA expression, depicted as number of PPARγ copies/25 ng mRNA vs. CD3+ T cell count, shown as number of cells/μL blood, in healthy donors was roughly 1 (Fig. 2D, controls), this relationship was significantly altered in all samples from septic patients, showing very high PPARγ expression in less T cells. As shown in Supplemental Figure 6, Supplemental Digital Content 1, http://links.lww.com/SHK/A360, CD4+ and CD8+ T cells of sepsis patients are equally affected (Supplemental Figure 6A, Supplemental Digital Content 1, http://links.lww.com/SHK/A360) and express similarly high PPARγ mRNA level (Supplemental Figure 6B, Supplemental Digital Content 1, http://links.lww.com/SHK/A360). Moreover, the induction of PTEN mRNA was followed as an established PPARγ target gene (16) in T cells derived from sepsis patients as shown in Supplemental Figure 6C, Supplemental Digital Content 1, http://links.lww.com/SHK/A360. These results suggest that the level of PPARγ expression might be useful as a prognostic factor for sepsis progression.
PPARγ as a marker for the outcome of septic patients
To determine, whether this inverse correlation can indeed be used as a prognostic marker for sepsis progression and survival, we arbitrarily set two limits: first, we included only patients with a PPARγ expression in T cells higher than 7,000 copies/25 ng mRNA (Fig. 3 A), of whom five of six patients died during the ICU stay (Fig. 3C). Second, we selected patients with a T cell count below 100 T cells/μL blood (Fig. 3B), of whom five of eight patients died (Fig. 3C). Among all 18 sepsis patients, four fulfilled the criteria for both arbitrary groups, namely high PPARγ expression and low T cell count, and all four of these patients subsequently died (Fig. 3C), confirming the relevance of the markers for outcome.
On the basis of this result, we reanalyzed the PPARγ data of all samples obtained from sepsis patients whether they survived or died during their ICU stay compared with the samples of healthy donors (1,264 ± 272 copies PPARγ/25 ng mRNA). As shown in Figure 4A and in Table 2, PPARγ expression was significantly higher in patients who died (5,391 ± 461 copies PPARγ/25 ng mRNA) compared with those patients who survived (3,086 ± 482 copies PPARγ/25 ng mRNA). This finding further supports our proposal to use PPARγ, in combination with the T cell count, as a prognostic marker for a poor sepsis outcome. Comparing PPARγ T cell mRNA expression (Fig. 2A) with expression in patients’ sera of IL-6 (Table 2 and Supplemantal Figure 6, Supplemental Digital Content 1, http://links.lww.com/SHK/A360), a putative sepsis marker routinely determined by the university hospital laboratory, no difference was seen in IL-6 expression in patients who died or survived in the ICU (Fig. 4A vs. B), in contrast to PPARgamma expression. Therefore, in our study, IL-6 was not a prognostic marker for sepsis outcome. Therefore, in our study IL-6 was no progonstic marker for sepsis outcome. Considering that a gain of time in sepsis diagnosis will improve effective sepsis therapy options (17,18), we reanalyzed IL-6 data of all patients by defining the day with highest IL-6 expression as day 0. All further samples were rearranged accordingly (Fig. 4C). Breaking down PPARγ mRNA data based on this IL-6 analysis scheme, we identified highest PPARγ mRNA expression in T cells 1 day before (= day -1) highest IL-6 expression (= day 0) as shown in Figure 4D.
Considering that sepsis is the one of the most frequent causes of death in ICUs, the identification of biomarkers, which can be used to characterize the prevailing disease status, is important. We focused our interest on the nuclear receptor PPARγ, which is known to be expressed in activated but not in resting T cells (6,12,19). Accordingly, we previously reported PPARγ expression in T cells derived from blood of septic patients (6). However, in this earlier study, the expression of PPARγ was not related to patient survival. In another report, on PPARγ expression in peripheral blood mononuclear cells (PBMC) derived from the blood of children with sepsis or septic shock, reduced expression of PPARγ was observed in total leukocyte lysates (20). The authors did not analyze PPARγ expression in enriched T cells, but assumed that the decrease of PPARγ monocyte expression was related to the septic conditions. Moreover, in this study, blood samples from children were used, whereas in our study the blood was taken from older people. Whether the age of the patients affects PPARγ expression in PBMCs seems unlikely, but has not been investigated.
With regard to other putative sepsis markers, such as the release of IL-6 (21,22), C-reactive protein (23) or procalcitonin (21,23), or the neutrophil surface expression of CD64 (24), we also compared PPARγ mRNA with that of IL-6 protein expression. IL-6 was determined in patients’ sera. Notably, maximal PPARγ expression was found 1 day before highest IL-6 protein expression (Fig. 4A vs. B), thereby offering a potential gain of time, which is important for the effective treatment of the septic patient at an early time point. Importantly, IL-6 expression did not differ significantly between patients who survived and patients who died during their ICU stay (Fig. 4B).
Induction of PPARγ expression in T cells during polymicrobial sepsis has been shown to contribute to T cell depletion, thus accounting for immune paralysis (3). This immune system failure is one of the main causes of poor prognosis and ICU mortality (8). Although PPARγ-dependent immune paralysis has only been shown in a mouse model (3), our present data support the assumption that the PPARγ-mediated mechanism observed in mice, might be translatable to the septic patient, too. Mechanistically in the mouse, PPARγ-dependent transrepression of the nuclear factor of active T cells (NFAT) occurred, thus blocking prosurvival IL-2 expression (3). To corroborate that this effect also occurs in the septic patient, we performed qPCR for IL-2 expression in CD3+ T cells. In line with the mouse data, in CD3+ T cells derived from septic patients, IL-2 expression was significantly reduced compared with T cells from healthy donors. Two things have to be taken into consideration here. First, T cells from healthy donors were not expected to be activated, thus IL-2 expression is low. Secondly, during a systemic inflammatory response, at an early time point, we would expect high IL-2 expression due to the presence of activated T cells fighting against the infection. Increased PPARγ expression probably prevents this IL-2 expression by repressing NFAT and blocking its binding to the responsive element in the IL-2 promoter (3,25,26), thereby decreasing IL-2 expression below the level of that in healthy donors. Although not directly addressed in this study, these results suggest that PPARγ antagonism may be a therapeutic concept to prevent T cell depletion in the septic patient, analogous to the effective use of the irreversible PPARγ antagonist GW9662 to inhibit the mouse sepsis model (3).
In conclusion, our pilot preliminary study, which admittedly did not include a large patient collective, suggests that PPARγ mRNA expression and T cell count in combination may be used as prognostic markers for a poor sepsis outcome. These data should be considered as providing a basic concept for a further study, which would include significantly more patients.
The authors thank Nadja Wallner for excellent technical assistance.
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Blood; ICU; IL-6; immune paralysis; patient survival; T cell depletion
Supplemental Digital Content
© 2016 by the Shock Society