Sepsis and its sequel multiple organ dysfunction syndrome are the leading cause of morbidity and mortality in the intensive care unit. The disorder affects approximately 750,000 North Americans annually, with more than 200,000 deaths attributed to it (1). Despite advances in the understanding of immune response, the role of neutrophils in sepsis is not well understood. An increase in white blood cell count is traditionally accepted as a protective physiological host response in sepsis. However, despite leukocytosis, a high mortality rate has questioned the protective role of neutrophils in sepsis (2, 3).
Neutrophils are known to eradicate invading microbes through the process of phagocytosis. Severe sepsis has been reported to affect phagocytosis as well; however, the reported results are quite conflicting. Phagocytosis by neutrophils may be enhanced (4-7), suppressed (8-12), or unaltered (13, 14) in response to proinflammatory stimuli. Different stimuli and experimental methods used in these studies can be the potential explanation for the contradictory results (15); however, the exact reasons for the observed discrepancy are poorly understood.
A compensatory increase in circulating immature neutrophils (>10% band forms) is a criterion to define the systemic inflammatory response syndrome (SIRS) (16). However, the functional competence of immature neutrophils and their impact on overall host response have never been evaluated in sepsis. Preliminary evidence in nonsepsis scenarios has pointed toward reduced functional responses of immature neutrophils (17, 18). Because none of the previous studies evaluating phagocytosis in sepsis commented on the effect of immature neutrophils, we argued the possibility of a similar reduced competence of immature neutrophils in sepsis resulting in a reduced host phagocytic capacity to eradicate infection.
To test our hypothesis, we conducted a pilot study in patients with severe sepsis to identify the proportion of immature neutrophils in them and to compare their phagocytic properties with phagocytosis by mature neutrophils in sepsis and in healthy volunteers.
MATERIALS AND METHODS
We studied 16 patients with severe sepsis admitted to the intensive care unit at the University Hospital of Wales, Cardiff. Patients older than 18 years with a suspected infection, who based on the clinical and microbiological grounds fulfilled the SIRS criteria (16), were enrolled in the study. Patients with neutropenia, metastatic malignant disease, human immunodeficiency disease, systemic corticosteroids, or immunosuppressive medications were excluded. The study protocol was approved by the local research ethics board, and written consent was obtained for all the patients and volunteers before enrollment.
Blood (approximately 5 mL) was drawn through indwelling arterial catheters from the patients and intravenously from the healthy volunteers. Samples were transported immediately to the laboratory for neutrophil isolation in heparinized tubes. Acute physiology and chronic health evaluation scores, multiple organ dysfunction scores, and total and differential leukocyte counts on the day of sampling were recorded. Intensive care unit lengths of stay and outcome data were collected retrospectively.
Source of reagents
Fura2-AM molecular probes (Eugene, Ore), purified human complement receptor type 3 (C3bi; Calbiochem, Nottingham, UK), antibodies to CD49d (DakoCytomation, Ely, UK), and the standard reagents (Sigma-Aldrich, Poole, UK) were used for the study purpose.
Circulating neutrophils were isolated from the patients and the healthy volunteers as described previously (19). Briefly, leukocytes were isolated from the citrated peripheral blood after dextran sedimentation of erythrocytes, platelet separation by centrifugation at 200 g for 10 min, and subsequent two washings with cold sterile phosphate-buffered saline (PBS) at pH 7.3. Neutrophils and mononuclear cells were separated on Ficoll-Paque by centrifugation at 400 g for 35 min at 20°C. Further treatment with hypotonic saline was done to lyse contaminating erythrocytes. After washing in PBS, neutrophils were suspended and counted in a Coulter counter (Coulter Electronics, Ltd, Beds, UK). Subsequently, iso-osmocity was restored with the addition of 2 mL of cold 3.5% (wt/vol) NaCl, and the cells were again centrifuged at 720 g for 5 min, followed by two washings in Ca2+-free Krebs 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) buffer containing 120 mM NaCl, 4.8 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 25 mM HEPES, and 0.1% bovine serum albumin. Finally, the cells were resuspended in Krebs HEPES buffer containing 1.3 mM CaCl2, and the medium was adjusted to a pH of 7.4 with NaOH. This suspension contained 98% neutrophils, as seen by phase-contrast microscopy. Neutrophils were diluted to 1 × 106 cells/mL and kept on ice until use. Immature neutrophils were differentiated from the mature neutrophils based on the characteristics of nuclei, number of nucleoli, and cytoplasmic granularity, as described previously (20). A separate immature cell counting by an observer, blinded to the study, served as the control.
C3bi opsonization and presentation of zymosan
Zymosan particles (10 mg/mL) were opsonized either by incubation with human serum (50% diluted; 30 min; 37°C) or with purified human C3bi (1 mg/mL; 30 min; 4°C). The particles were then washed by centrifugation and resuspension to remove unfixed C3bi and used immediately or stored at −20°C.
Phagocytosis was first quantified in neutrophil suspensions (21), with subsequent confirmation by the micromanipulation technique after individual cell analysis for cellular maturity (22). For the initial quantitation in the neutrophil suspensions, opsonized zymosan particles were added to the neutrophils and suspended in Krebs medium in a ratio of 10:1 particles per cell (106/mL). Neutrophils were subsequently incubated at 37°C for 1 h. Thereafter, the neutrophils were fixed on glass slides at room temperature by immersing in methanol for 15 min, followed by a May-Grunwald stain (diluted 1:1 with PBS at pH 6.8) for another 15 min. Four hundred neutrophils were counted in each sample for internalized zymosan particles under a microscope (×40-100).
For micromanipulation studies (22), neutrophils were allowed to adhere to glass coverslips for 10 min at 37°C before the addition of C3bi-opsonized zymosan particles. Zymosan particles (×2 mm in diameter) were allowed to sediment among the cells. A micropipette (tip diameter, 1-1.5 μm; WPI, Stevenage, UK), with slight negative pressure applied, was used to pick and hold a single zymosan particle. The particle was brought to a target neutrophil to facilitate the contact between the two and to allow a subsequent phagocytosis to proceed. A phagocytic event was defined when a neutrophil had completely internalized a zymosan particle. For this purpose, 50 cells were counted in each neutrophil subpopulation.
Phagocytosis index (PI) was calculated (PI = cells containing zymosan particle / total cells counted × 100) (23). Moreover, the extent of phagocytic work (particulate burden within each cell) was calculated by counting the number of zymosan particles within each neutrophil (24).
Simultaneous cytosolic-free Ca2+ and phagocytosis imaging
For Ca2+ imaging studies, cells were loaded with the Ca2+ probe fura-2. Individual neutrophils, both the mature and immature ones, were challenged with C3bi-opsonized zymosan particles by micromanipulation technique. Subsequent free Ca2+ responses were recorded. The ratiometric (Ca2+) images were pseudocolored according to the previously described scale. A single or a twin peak of a rapid and transient intracellular Ca2+ increase (t1/2 <50 s) suggested normal phagocytosis (22, 25).
Results are expressed as mean values ± SD for continuous variables and as percentages for nominal variables. Continuous variables between the two groups were compared by unpaired t test or Mann Whitney U test as applicable. Correlation was done using Spearman correlation analysis, and statistical differences were considered significant at P < 0.05 (SPSS v 12.0, Chicago, Ill).
Sixteen patients and five healthy controls were enrolled. In the patient group, the average age was 60.5 ± 15.8 years; 63% of them were women. At admission, mean acute physiology and chronic health evaluation II scores and multiple organ dysfunction scores were 26 ± 2.4 and 8.3 ± 0.6, respectively. Mortality at 28 days was 37.5%. Healthy volunteers (three women, two men) had an average age of 41 ± 4.5 years.
Immature neutrophils in sepsis
Of the 16 patients, 12 had elevated leukocyte counts (normal, 4-12 × 109 cells/L). Circulating immature neutrophils (Fig. 1) were present in 9 of the 12 patients with leukocytosis and 3 of the 4 patients with normal leukocyte counts (39.3% ± 20.7%; normal ≤5%; Table 1). The counting of the immature neutrophils correlated well with the reported results by the blinded observer (35% ± 16.7%; r = 0.98). In the control group, all the volunteers had less than 5% immature neutrophils.
Neutrophils and their phagocytic capacity in sepsis
On incubation of neutrophils with C3bi-opsonized zymosan particles in suspension, PI was only modestly increased in patients with sepsis as compared with the healthy controls (52% ± 41% vs. 43% ± 10%; P = 0.3; Fig. 2A). Notably, neutrophils from sepsis patients showed a large variation in their phagocytic ability as reflected by their standard deviation. For example, in a typical sepsis patient, 39% neutrophils phagocytosed three or less zymosan particles (vs. 3% neutrophils from healthy donors; Fig. 2B). In the same sample, a subpopulation of neutrophils (58%) also phagocytosed more than five zymosan particles. Consistent and accurate identification of neutrophil maturation status was difficult in the presence of multiple internalized zymosan particles; hence, individual cell analysis for neutrophil maturation stages was conducted and micromanipulation studies with zymosan were carried out under direct vision.
Prior identification of individual neutrophils for their maturity and subsequent micromanipulation with zymosan particles demonstrated a significant difference in the phagocytic capabilities between mature and immature neutrophils. Mature neutrophils from the patients with sepsis displayed a significantly greater PI as compared with the mature neutrophils from healthy donors (69% ± 8% vs. 42% ± 6%; P < 0.05). In contrast, immature neutrophils in these patients demonstrated a significantly decreased PI (25% ± 5%; P < 0.05) as compared with mature neutrophils from both patients and controls (Fig. 3). Moreover, mature neutrophils from sepsis also demonstrated an increased internal particulate burden as compared with the mature neutrophils from healthy donors (9.4 ± 1 and 5.9 ± 1.1 particles per cell; P < 0.05). Immature neutrophils in sepsis again phagocytosed significantly fewer particles (2 ± 0.5 zymosan particles per cell; P < 0.05) as compared with mature neutrophils from the sepsis patients and healthy volunteers.
Altogether, these findings suggested that the evaluation of neutrophil phagocytosis by the suspension method did not reveal a significant difference between sepsis patients and healthy donors, with sepsis patients merely demonstrating a marked variation in the phagocytic capacity. However, micromanipulation studies after individual cell identification provided the explanation by identifying two distinct subpopulations of circulating neutrophils in sepsis with significantly different phagocytic capabilities.
Calcium signaling and immature neutrophils
Normally, phagocytosis of C3bi-opsonized particles requires a global elevation in cytosolic-free Ca2+, which results from Ca2+ influx (22, 25). Mature neutrophils in healthy volunteers demonstrate a single peak (t1/2 = 50 s) or twin peaks, with the magnitude of Ca2+ change from approximately 100 to 550 nM. The first Ca2+ signal occurs during β2 integrin engagement, as the phagocytic cup forms, and the second global signal triggers oxidase activation within the phagosome. In our study sample, all mature neutrophils from patients and healthy donors demonstrated large Ca2+ signals with single or twin peaks, with the magnitude of the Ca2+ change being from 100 to 550 nM (Fig. 4). Immature neutrophils from both sepsis patients and healthy controls, however, elicited weak Ca2+ signals with multiple peaks, similar to those seen in HL 60 cells (26).
Neutrophils form a critical component of the host defense against infection. The bone marrow normally produces approximately 1011 neutrophils a day, and their production can increase by 10-fold in the presence of a triggering stimulus (27). Despite a large reserve, immature neutrophils can still spill over in the circulation during severe infection, and greater than 10% circulating band forms is used as a criterion to define SIRS (16). Although considered a physiological response, preliminary evidence in an animal model (28) and a small study in trauma patients (18) suggested a reduced functional competence of immature neutrophils. Immature neutrophils can even have a detrimental effect on the host as suggested by their preferential sequestration in the lung microvasculature (28). Because of lack of similar data in human sepsis, we identified immature neutrophils in critically ill patients with severe sepsis and attempted to evaluate their phagocytic function.
Immature neutrophils were demonstrated in 9 of the 12 patients with leukocytosis. Interestingly, three of the four patients with no leukocytosis also showed an increase in immature neutrophils. Sepsis patients had an average of 39% immature neutrophils as compared with less than 5% in healthy controls. Although detection of immature neutrophils in the peripheral blood was an expected finding; still, their large proportion, despite normal total leukocyte counts in a few patients, was an interesting observation.
We used cellular morphology to identify immature neutrophils. Other techniques such as density centrifugation and flow cytometry have been used to isolate immature neutrophils; however, they have also been reported to yield an incomplete segregation of immature neutrophils (29, 30). The immature neutrophils were identified in this study by previously described standard morphological criteria (20), and this technique enabled us to perform further micromanipulation studies on individual cells for phagocytosis experiments.
Quantification of phagocytosis in patients with sepsis revealed a difference in phagocytic function of neutrophils depending on their stage of maturity. Mature neutrophils in sepsis demonstrated significantly increased phagocytosis as compared with their immature precursors and mature neutrophils from healthy controls. Previous studies on phagocytic efficacy in sepsis did not evaluate the reduced phagocytic capability of neutrophils precursors, and it is possible that a different proportion of unaccounted neutrophil precursors can be a potential reason for the conflicting results in these studies (5-7, 9-14). Even in our study sample, initial evaluation by the conventional suspension method could not detect a difference in the phagocytic pattern of the mature and immature neutrophils, and we noticed an equivalent phagocytosis in the patients due to the counterbalancing of increased phagocytosis of the mature neutrophils by the reduced phagocytosis of immature neutrophils. Similarly, the previously reported high morbidity and mortality in the presence of leukocytosis (2, 31, 32), and a lack of improved outcome despite leukocytosis induced by colony-stimulating factors (33, 34), can also be the result of an increase in functionally less competent immature neutrophils. To our knowledge, this is the first study that highlights the impaired phagocytic function of immature neutrophils in severe sepsis.
Phagocytic activity is altered through a variety of the mechanisms, including reduced expression of CR3 (β2 integrin) (33), calcium-related signaling (34), altered membrane characteristics, and decreased cell deformability (35). We evaluated phagocytosis by a previously established technique of zymosan internalization and change in intracellular calcium signaling (22, 25). With this technique, we can demonstrate a relationship between the immature neutrophils and reduced phagocytosis; however, we could not comment on any variation in phagocytic capability secondary to different maturation stages of immature neutrophils.
The main objective of our pilot project was to evaluate the phagocytic capacity of immature neutrophils in sepsis patients. In addition, we also tried to explore a potential impact of immature neutrophils on overall phagocytic capability. Our results indicate that, despite increased numbers of immature neutrophils, overall phagocytic function in sepsis patients did not differ significantly as compared with the controls. We have previously reported a significant delay in neutrophil apoptosis in sepsis (36); however, the effect of sepsis on neutrophil maturation is not known. It is possible that the proportion of immature neutrophils in a patient with sepsis may be determined by the severity of triggering stimulus and the balance between neutrophil production, maturation, and elimination. Thus, the proportion of immature and mature neutrophils may vary during the course of sepsis in different individuals and also at different times in the same individual. It is possible that a cross-sectional design of our study could not adequately evaluate a possible longitudinal change in phagocytosis during the course of a sepsis episode. Considering that a physiological response to life-threatening infection should comprise an enhanced bacterial clearance by the host, future longitudinal studies are needed to clarify the clinical impact of immature neutrophils on overall phagocytic function over the time span of a sepsis episode.
We conclude that severe sepsis is associated with a significant increase in circulating immature neutrophils. These immature neutrophils have decreased phagocytosis and aberrant calcium signaling. Although our results did not reveal an effect of these cells on overall phagocytosis in sepsis patients, future longitudinal studies are needed to evaluate the clinical impact of our observations.
The authors thank the nursing and medical staff in the intensive care unit of the University Hospital of Wales for help in patient recruitment and sample collection.
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