Sepsis has been recently qualified as the quintessential medical disorder of the 21st century (1). It is the most frequent cause of death in intensive care unit (ICU) patients while, to date, no therapeutic intervention has specifically been approved for its treatment. While sepsis has been usually described as inducing a tremendous systemic inflammatory response, current data indicate that it initiates a more complex immunologic response that evolves over time, with the simultaneous occurrence of both pro- and anti-inflammatory mechanisms (1, 2). As a result, after a short unbridled pro-inflammatory phase, septic patients present with profound immunosuppression that would prevent efficient clearing of the primary infection despite adequate antibiotics (3–5). This immune defect is also illustrated by reactivation of latent viruses (CMV or HSV) or increased nosocomial infections due to pathogens, including fungi, which are normally weakly virulent or opportunistic (3). Although secondary infections may contribute only modestly to overall mortality (6), globally, these alterations are believed to be responsible for worsening outcome in patients who survived initial resuscitation since the magnitude and persistence over time of immunosuppression is known to be associated with increased mortality. In accordance, it is reported that a majority of deaths occur after the first 3 days of the disorder (7, 8). Consequently, new therapeutic options based on adjunctive immunostimulation (IFN-γ, GM-CSF, IL-7, anti-PD1/L1 antibodies) are emerging (9). Nevertheless, as there is no clinical sign of immune dysfunctions, such intervention relies on biomarkers for identifying the patients who could benefit from immunostimulation (i.e., the most immunosuppressed). Among potential biomarkers, immunologically speaking, functional testing remains the gold standard because it directly measures the capacity of a cell population to respond to an immune challenge (10, 11). However, these protocols although providing excellent insights regarding pathophysiology remain barely usable on a routine clinical monitoring basis (long incubation time, lengthy cell purification procedures, low standardization). Thus, one current major challenge is to abolish the gap between functional testing for comprehensive studies and functional testing usable on routine clinical daily practice (12).
Although observed in whole blood including a mixture of various cell types, the decreased capacity to release pro-inflammatory cytokines (mainly tumor necrosis factor [TNF]) in response to endotoxin challenge is considered a pivotal feature of altered monocyte functionality during sepsis-induced immunosuppression (13, 14). As intracellular flow cytometry allows determination of both cell function and phenotype at the same time, the detection of cytokines directly inside cells producing them is one of the most elegant ways to assess immune reactivity of a given cell subset (15). We took advantages from recent improvements in intracellular staining for flow cytometry to design a rapid and simple protocol to measure in vitro TNF production by monocytes in response to lipopolysaccharide (LPS) (no cell purification, no wash, no centrifuge procedure). The aim of this study was thus to evaluate feasibility in clinic and suitability to routine workload of this novel whole blood technique for the measurement of intracellular TNF production in monocytes. This was performed in the clinical context of septic shock. In addition, considering the large body of literature describing the potential of monocyte HLA-DR (mHLA-DR) in monitoring septic patients (11), we also assessed the interest and feasibility of including HLA-DR staining within a single tube that would concomitantly explore TNF and HLA-DR expression in monocyte.
PATIENTS AND METHODS
The study group consisted of 16 septic shock patients (diagnostic criteria of the International Guidelines for Management of Severe Sepsis and Septic Shock) (16), admitted to the surgical ICU (details provided in supplemental digital content, http://links.lww.com/SHK/A452).
We used unitized 12 × 75 mm tubes containing a dry coating of LPS (from E coli O127:B8, final concentration: 200 ng/mL, Sigma [Saint-Quentin-Fallavier, France]), and brefeldin A to block cytokine secretion via the Golgi apparatus and unitized 12 × 75 mm tubes containing a dry coating of conjugated antibodies formulated for this assay: Krome Orange (KrO)-labeled anti-CD14 (clone RMO52), Alexa fluor-700 (AF700)-labeled anti-TNF (clone IPM2), and phycoerythrin (PE)-labeled anti- HLA-DR (clone Immu-357). These research reagents are custom-made reagents, optimized for this research evaluation by Beckman Coulter Immunotech, and are not available as commercial products.
Intracellular staining procedure
Heparin anticoagulated blood was collected from patients during the first 24 h after diagnosis of septic shock. Fifty microliters of undiluted whole blood was directly added to the stimulation tube (LPS-BrefA), or to an empty control tube, then incubated 2 h at 37°C. Samples were then treated according to the regular PerFix-no centrifuge (nc) (Beckman Coulter, Brea, Calif) procedure: in brief, fixation of the sample was conducted for 15 min at room temperature, applying 5 μL of PerFix-nc Fixative Reagent, followed by 15 min permeabilization and concomitant staining at room temperature in the dark (dried antibodies were resuspended extemporaneously with 300 μL of the Permeabilizing Reagent, this mixture being added to the primary reaction tube). Finally, reaction was terminated by adding 2.5 mL of PerFix-nc final reagent. After vortexing, cells were ready to be acquired on flow cytometer. Overall, it is a no wash, no centrifuge procedure completed in precisely 2.5 h. A schematic protocol representation is presented in Figure S1 (supplemental digital content, http://links.lww.com/SHK/A452).
Flow cytometric data acquisition and analysis
Data acquisition was performed on a Navios Flow Cytometer using Kaluza software for data analysis (Beckman Coulter, Brea, Calif). There was no compensation applied, since only flurochromes without spectral overlap were selected to eliminate the need for compensation insuring a better robustness to the assay. Monocytes were first gated out from other cells on the basis of labeling with CD14. Intracellular TNF (iTNF) expression was then measured on this selected population (CD14 high). All results were expressed either as percentages of TNF-positive monocytes (% positive cells) among the total monocyte population (positivity threshold was defined with non-stimulated values from healthy donors) or as means of fluorescence intensity (MFI) of the entire monocyte population. According to same gating strategy of monocytes, we also determined HLA-DR expression on their surface and expressed results as MFI thanks to antibodies against HLA-DR (clone Immu-357) included in the same tube. Thereafter, this mHLA-DR value will be called experimental HLA-DR (expHLA-DR) to avoid any confusion with mHLA-DR expressed as ABC and obtained through standardized protocol.
Details on mHLA-DR measurement (standardized protocol) and statistical analysis are provided in supplemental digital content, http://links.lww.com/SHK/A452.
Sixteen septic shock patients and eight healthy controls were included. Clinical and biological characteristics are listed in Table S1 (supplemental digital content, http://links.lww.com/SHK/A452). Briefly, median values for age, SAPS II, and SOFA scores at onset of shock were 66 (interquartile range [IQR] 61–72), 67 [IQR 51–81], and 10 [IQR 9–12] respectively. Five patients died within 28 days (mortality = 31%) and secondary nosocomial infections were diagnosed in three patients (19%). At D3-4, septic shock patients presented with usual characteristics of injury-induced immunosuppression with a reduced mHLA-DR expression, low CD4+ lymphocyte count, and an increased percentage of circulating Treg cells in comparison with normal values (supplemental digital content , Table S1, http://links.lww.com/SHK/A452).
Intracellular TNF results
As expected, LPS challenge induced a tremendous expression of iTNF (Fig. 1). This increase was statistically more pronounced in controls than in patients. This was observed when results were expressed as MFI (16.1 [IQR 14.5–19.1] vs. 5 [IQR 4.0–8.0], P = 0.0001) or as percentage of positive cells (99.7 [IQR 99.6–99.8] vs. 85 [IQR 74–97], P = 0.0001, Fig. 2). When considering controls and patients values as a whole, iTNF expression was correlated to mHLA-DR expression (standardized measurement as ABC) at day 3 (r: 0.79, P <0.0001). This correlation was less obvious when solely focusing on values from septic patients (Figure S2, supplemental digital content, http://links.lww.com/SHK/A452). It may indicate that for a given mHLA-DR level (herein most values <10, 000 ABC), various degrees of functionality may be measured.
expHLA-DR expression in stimulated samples (LPS-BrefA)
In addition to mHLA-DR results obtained by the standardized protocol in a separate tube, we also assessed expHLA-DR expression in the iTNF tube. One potential issue was the necessity to incubate cells at +37°C as it is well known that monocyte HLA-DR artificially rises up if staining is not rapidly processed after sampling (17). Actually, this enhancement was not seen in patients’ samples but only in healthy controls (Figure S3, supplemental digital content, http://links.lww.com/SHK/A452). We obtained good correlations between mHLA-DR and expHLA-DR (+4°C) or expHLA-DR (+37°C), correlation coefficients were 0.68 (P: 0.004) and 0.70 (P: 0.003) respectively. We observed an excellent correlation between iTNF and expHLA-DR results (Figure S4, supplemental digital content, http://links.lww.com/SHK/A452), even when focusing only on septic patients. This indicates that a tricolor protocol (CD14-iTNF-HLA-DR) would be feasible (in a single tube with 50 μL whole blood) and provide both crucial information: mHLA-DR expression and monocyte capacity to respond to LPS challenge (i.e., functional testing).
The weight of evidence now supports the view that the immune-suppressive phase is the predominant immunological response in most septic patients (18). As there is no clinical sign of immune dysfunctions, it is crucial to identify the best biological tools for patients’ stratification according to their immune status. Since it directly measures ex vivo capacity of a cell population to respond to an immune challenge, functional testing ideally represents the best method to establish the occurrence of immunosuppression. Regarding monocytes, many groups have described the altered capacity of septic patients’ cells to release pro-inflammatory cytokines in response to stimulation with LPS, various TLR agonists, or whole bacteria in vitro(13, 14, 19, 20). These tests, based on cytokine measurements in whole blood supernatant, represent reliable methods to assess the phenomenon of endotoxin tolerance, i.e., refractory state to subsequent endotoxin challenge (13). That said, they are not really suitable to routine monitoring as they are time consuming (incubation time + centrifugation + cytokine Elisa assay) and not standardized yet (19, 21). Intracellular cytokine staining protocols, commonly used in experimental studies, would be an attractive alternative (15). To our knowledge, nothing has been conducted in significant clinical cohorts of septic patients whereas preliminary results provided convincing results. Indeed, Fumeaux et al. (22) observed that septic monocytes were characterized by an altered IL-10/TNF intracellular ratio. Again, it is likely due to lengthy protocols including many permeabilization/staining/wash/centrifugation/incubation steps (15, 23). Recently, progress has been made regarding intracellular protocol for flow cytometry (e.g., Foxp3) (24). This is also true for functional testing based on intracellular parameter determination. A typical example is provided by whole blood phospho-STAT5 measurement in response to IL-7 stimulation (25). The entire protocol takes 90 min to be completed in one step. It provided promising results in septic shock patients in which it could differentiate survivors from non-survivors depending on their STAT5 response upon IL-7 challenge (26).
In the present work, we took advantage from recent progress for intracellular staining flow cytometry to design and evaluate a new protocol dedicated to cytokine detection in monocytes. Within the context of sepsis-induced immunosuppression, measurement of iTNF would constitute a real improvement since simple, rapid, and robust functional testing is regularly desired in order to better define immune status of patients. Overall, the setup of the protocol was easy in our hands and we did not report major issue. It takes only twice 50 μL of whole blood (unstimulated and LPS-stimulated conditions). It is a no-wash, no-centrifuge procedure and everything is standardized in tubes thanks to the dry coating of reagents (what further reduces the pipetting steps). Of note, these dried reagents are stable at room temperature. Functional results are obtained in 2.5 h and flow histograms are easy to interpret, without any compensation needed. In addition, results are specific for a given cell type (i.e., monocyte) whereas usual TNF release test measures cytokine production from all mixed cells contained in whole blood (21). As depicted in Figure S1 (http://links.lww.com/SHK/A452), manual interventions are very limited meaning that a technician could easily manage several samples in parallel with other routine tasks (this was not the case with former protocols). Thus, technically speaking, we made the proof of concept that this iTNF protocol could be an appropriate tool for routine work. In addition, we would like to emphasize that, beside the availability of flow cytometers in routine hospital labs that could be a limiting factor, no strong flow cytometry skills are required for users as this is a ready-to-use kit performed in whole blood (no need for cell purification) that does not need any fluorescence compensation (limitation in most FACS protocols and for standardization in multicentric studies). As far as intracellular flow cytometry measurements, it appears difficult to design a simpler functional protocol. Thus, we strongly believe that the adaptability of this test to general clinical setting can be very good and that, upon validation in larger cohorts of patients, this protocol may become a real tool for patients’ monitoring. In addition, upon validation in a larger cohort of patients, one may likely skip the nonactivated tube as we did not detect any iTNF both in controls and septic patients (i.e., no baseline production). The present preliminary patients’ results look very promising as there was a major fall in TNF production in monocytes from septic patients. In addition, iTNF values were nicely correlated to mHLA-DR values at day 1 to 2 and day 3 to 4 (gold standard in the field). Although it should be explored in larger cohorts of patients, monocyte reactivity might be even more informative than mHLA-DR regarding association with deleterious outcome. Moreover, we also reported that both intracellular and cell surface staining are concomitantly doable suggesting the possibility of a kit including both iTNF and mHLA-DR values. Overall, preliminary results are encouraging and this approach might become a standard tool to stratify patient for trial enrolment or to manage drug efficacy. Indeed, whole-blood TNF release has been once used to stratify patients in a small clinical trial evaluating GM-CSF as an immuno-adjuvant therapy in pediatrics (27), whereas administration of GM-CSF to adult septic patients induced a restoration of ex vivo TNF-α production capacity (28).
Of note, one potential drawback of this approach remains the use of flow cytometry which is not, to date, available everywhere and often linked to specialized laboratories. In addition, our study presents with limitations mostly inherent to the fact we conducted a proof of concept study. Indeed, this work was not designed to improve our understanding of septic shock pathophysiology but rather to evaluate in clinical conditions the feasibility of such an innovative functional assay. Results are thus preliminary as we only included 16 patients. We did not provide comparison with TNF whole-blood release or with alternate iTNF intracellular staining (based on usual protocol in purified cells). However such comparisons should now be performed in a further study. Due to ethical reason, we only got a single time point of blood sampling (day 1). So, we cannot speculate on later time points in patients’ evolution. In addition, healthy volunteers were not strictly age-matched with septic shock patients so we cannot definitely rule out an effect of age on our results. That said, it has been recently reported that the loss of in vitro TNF release in septic patients was not due to age (14). Lastly, we obviously could not investigate association with clinical outcomes as our study was not powered for such purposes.
Overall, these preliminary results indicate the reliability of this novel iTNF protocol that deserves now to be widely assessed and validated in various ICU conditions. In sepsis, innovative immuno-monitoring strategies that characterize the host immune response and thus permit personalized medicine approaches appear as a reasonable perspective. This would allow to decrease heterogeneity of septic population and to better take into account the rapidly changing immune response overtime after initial infection, from exacerbated inflammation to severe immunosuppression. For such purposes, the possibility of obtaining functional testing on routine manner could be a major step.
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