Neutrophil (polymorphonuclear leukocytes [PMN]) infiltration plays a central role in inflammation and is also a major cause of tissue damage. Thus, PMN infiltration must be tightly controlled. Using zymosan-induced peritonitis as an in vivo PMN infiltration model, we show in this study that PMN response and infiltration were significantly enhanced in mice experiencing various types of systemic inflammation, including colitis and diabetes. Adoptive transfer of leukocytes from mice with inflammation into healthy recipients or from healthy into inflammatory recipients followed by inducing peritonitis demonstrated that both circulating PMN and tissue macrophages were altered under inflammatory conditions and that they collectively contributed to enhanced PMN infiltration. Detailed analyses of dextran sulfate sodium-elicited colitis revealed that enhancement of PMN infiltration and macrophage function occurred only at the postacute/chronic phase of inflammation and was associated with markedly increased IL-17A in serum. In vitro and ex vivo treatment of isolated PMN and macrophages confirmed that IL-17A directly modulates these cells and significantly enhances their inflammatory responses. Neutralization of IL-17A eliminated the enhancement of PMN infiltration and IL-6 production and also prevented severe tissue damage in dextran sulfate sodium-treated mice. Thus, IL-17A produced at the chronic stage of colitis serves as an essential feedback signal that enhances PMN infiltration and promotes inflammation.

Polymorphonuclear leukocyte (PMN) infiltration into tissues during inflammation plays a central role in innate immunity. Upon pathogen infection or irritant infliction, local macrophages and other cells sense the insult and produce a panel of inflammatory mediators such as cytokines and chemokines that stimulate the nearby microvasculature and attract large numbers of PMN to migrate across the vascular wall and infiltrate into tissues. After arrival at the inflammatory site, PMN perform phagocytosis and also release powerful antipathogen and tissue-damaging reagents to kill pathogens and aberrant cells. Although these forceful PMN activities are extremely important for host defense, their adverse effects are also apparent, as demonstrated in inflammatory bowel diseases, arthritis, some cardiovascular conditions, inflammatory pulmonary and renal diseases, and viral/bacterial infection-associated damage. In these cases, the powerful but relatively low target specificity of PMN results in grave tissue/organ injuries, often leading to a rapid progression of the pathophysiological condition. Thus, effective control of PMN infiltration would much alleviate the inflammatory condition.

The mechanisms that control PMN response and regulate their infiltration involve multiple cell types and various inflammatory factors, and studies in this area are still inadequate. Some aspects of PMN function, such as chemotaxis and degranulation, have been studied in vitro by refined assays (13). However, these assays do not reveal the entire design of PMN inflammatory response and infiltration during inflammation and how these processes are dynamically regulated in vivo. In the study of PMN function in diabetes, we have constantly observed that PMN isolated from diabetic patients showed impaired transmigration in the in vitro chemotaxis assays (Y. Liu and K. Zen, unpublished observations). These observations, which are consistent with other reports (46), indicated an important fact: that PMN from diabetic patients were different from those of healthy donors; however, they could not ascertain whether the activities of PMN of diabetes were truly abated or, on the contrary, were primed (activated to an extent). In the second possibility, the primed PMN were hypersensitive and may have become activated during the in vitro isolating procedure, resulting in their inefficient chemotaxis in the later transmigration assays (3).

To further study PMN response and infiltration, we designed a two-layered in vivo inflammatory system in which a relatively chronic inflammatory condition, such as colitis, type 1 diabetes, or other inflammatory condition, was first created in the experimental animals. After the condition was stabilized, a typical acute inflammatory response, such as zymosan-induced peritonitis, was induced, and the immediate PMN infiltration into the newly established inflammatory site was assayed within a few hours. Using this design, we found that active inflammation at the postacute/chronic phase in particular induces systemic alterations in both PMN and cells within tissue microenvironments, including macrophages. Instead of suppressing inflammation, these changes result in an enhancement of PMN infiltration, hence exaggerating the inflammatory reaction.

C57BL/6 mice 6–10 wk old (18–22 g) purchased from The Jackson Laboratory (Bar Harbor, ME) were housed with free access to water and food in a specific pathogen-free facility. All experiments using animals and procedures of animal care and handling were carried out following protocols approved by the Institutional Animal Care and Use Committee of Georgia State University. To establish dextran sulfate sodium (DSS)-induced colitis (7, 8), 2% DSS (MP Biomedicals) prepared in pure water was given to the mice as the drinking water. Mice were inspected daily for distress and colitis symptoms such as wet stool, diarrhea, bloody diarrhea, and weight loss. To induce hyperglycemia/type 1 diabetes (9, 10), the mice were fasted for 4 h and then injected with streptozotocin (STZ; Sigma-Aldrich) at a dose of 50 mg/kg body weight. STZ was freshly prepared in sodium citrate buffer (pH 4.5) prior to the injections, which were performed daily for 5 consecutive d. The mice were then maintained for an additional 2 wk to allow the diabetic condition to be stabilized. Serum glucose levels were tested with the One-Touch glucose monitoring system (LifeScan), and in general, >90% of injected mice developed hyperglycemia. To induce bronchitis/lung inflammation, the mice were given LPS (Escherichia coli serotype 026:B6; Sigma-Aldrich), 0.3 mg/kg body weight, in 70 μl PBS through the nares every other day for 15 d.

This model was used to assay the acute PMN response and infiltration in vivo (11). Briefly, mice were injected i.p. with 0.5 mg zymosan A (Sigma-Aldrich) that was prepared in 0.5 ml PBS. Mice were euthanized at varying time points after the injection. PMN that transmigrated into the peritoneum were collected by peritoneal lavage with 3 ml cold HBSS without Ca2+ and Mg2+ followed by analysis with myeloperoxidase (MPO) assay, cell counting, and immunofluorescence labeling using a PE-Cy5–conjugated anti-mouse Ly-6G (Gr-1) Ab (eBioscience). Macrophages were analyzed using an Alexa Fluor 488-conjugated anti-mouse F4/80 Ab (clone BM8; BioLegend). For leukocyte adoptive transfer experiments, bone marrow cells were obtained by flushing femur bone cavities followed by labeling with the fluorescent cell tracer CFSE or 1,1'-dihexadecyl-3,3,3′,3′-tetramethylindocarbocyanine iodide [DiIC16(3)] (both from Invitrogen). In some experiments, the bone marrow cells were further separated by a Percoll (GE Healthcare) density gradient to obtain the population of mature PMN (12). In general, 5 × 106 labeled cells (∼2.5 × 106 were Gr-1+) were transfused into a recipient mouse via i.v. injection before inducing peritonitis using zymosan. To determine the role of IL-17A in PMN infiltration, ex vivo experiments were performed. In these experiments, freshly isolated bone marrow cells were labeled with CFSE followed by treatment with a recombinant mouse IL-17A (rIL-17A) (2 ng/ml; BioLegend), with or without the presence of a neutralization anti–IL-17A Ab (low endotoxin, azide-free–purified anti-mouse IL-17A Ab [BioLegend]), or 50 μl serum collected from nontreated mice or mice treated with DSS for 12 d. After incubation for 2 h (37°C), the cells (∼2.5 × 106 PMN) were combined with equal amount of DiIC16(3)-labeled bone marrow PMN before transfer into healthy recipient mice. Zymosan-induced peritonitis was then performed to test the transferred PMN infiltration into the peritoneum.

Whole blood from mice was collected without anticoagulation additives, and the serum was isolated. For ELISA, mouse serum of different dilutions was incubated in anti-cytokine Ab-coated wells in 96-well plates for 2 h (25°C). After washing, the wells were incubated with biotin-conjugated detecting Abs and HRP-conjugated streptavidin followed by detection with o-phenylenediamine dihydrochloride (Sigma-Aldrich) substrate. Both capture and detecting Abs against different cytokines including IL-1β, IL-4, IL-6, IL-12 p40/70, IL-17A, IFN-γ, TNF-α, MCP-1, IL-10, and GM-CSF were purchased from BD Biosciences and BioLegend. Purified recombinant murine cytokines used as the standards were purchased from PeproTech. To detect cytokines produced by in vitro-treated macrophages, peritoneal macrophages were obtained by lavage of the peritoneal space and collection by centrifugation. The macrophages were then stimulated with zymosan or rIL-17A (2 ng/ml) with or without the presence of the anti–IL-17A Ab (2 mg/ml) for various time periods up to 48 h in DMEM with 8% FBS (37°C). After the treatment, the supernatants were collected, and levels of IL-6 and other cytokines were detected by ELISA.

An in vitro transmigration setup using transwell devices with collagen-coated transfilters (0.33 cm2, 5 μm pore size) established previously (13) was used to assess PMN chemotactic capability. Briefly, bone marrow leukocytes were isolated from femur bones of nontreated, healthy mice and mice with DSS-induced colitis or STZ-induced diabetes by flushing the bone cavities using Hank’s buffer devoid of Ca2+ and Mg2+. After lysis of RBC, percentages of PMN in cell populations were determined by MPO assay and immunofluorescence staining using PE-Cy5–conjugated anti-mouse Ly-6G (Gr-1) Ab. Aliquots of cell samples from each type of mice were also subject to further separation to obtain the population of mature PMN according to a protocol described by Dong and Wu (12). After normalization of the amount of mature PMN, the cells (containing 2.5 × 106 PMN) were added into the upper chamber of the chemotaxis setup in 150 μl HBSS followed by inducing transmigration with 10 μM chemoattractant fMLF added into the lower chamber in 500 μl HBSS. The setup was then incubated at 37°C, and the PMN transmigrated across the filter into the lower chamber were then quantified at different time points by MPO assay. To test the effect of IL-17A on PMN, the freshly isolated cells loaded in the upper transmigration chambers were treated with rIL-17A with or without neutralization Ab or vehicle control buffer for 2 h before inducing transmigration by fMLF.

For histochemical staining, mouse tissues including colon and lung were fixed in 4% paraformaldehyde, embedded in paraffin, and cut into 5–10-μm-thin sections. Sections were stained with H&E. For staining of infiltrated leukocytes and PMN, freshly sectioned tissues were blocked with 5% BSA followed by staining using a PE-Cy5–conjugated anti-mouse Ly-6G (Gr-1) Ab. Images were then taken using an Olympus BX21 fluorescence microscope (Olympus).

To directly observe IL-17A binding to PMN and macrophages, recombinant murine IL-17A was labeled with FITC using an EZ-Label FITC protein labeling kit (Pierce). The FITC-labeled IL-17A was then incubated with freshly isolated mouse bone marrow leukocytes and peritoneal macrophages for 30 min (25°C) followed by washing and analysis by fluorescence microscopy. To detect IL-17A–induced protein phosphorylation changes in macrophages, isolated murine peritoneal macrophages were treated with rIL-17A at concentrations of 0.5 and 2.0 ng/ml, or vehicle, for 48 h (37°C). The cells were then lysed in a buffer containing 100 mM Tris (pH 7.5), 150 mM NaCl, 1% Triton X-100, 1:100 dilution of a protease inhibitor mixture (product code P8340; Sigma-Aldrich), and 3 mM PMSF followed by SDS-PAGE and Western blot detection using the anti-phosphotyrosine Ab 4G10 (Millipore), an anti-phosphoserine Ab (Cell Signaling Technology), and an anti-actin Ab (Sigma-Aldrich).

In this study, we used zymosan-induced peritonitis (11), a commonly used self-resolving acute inflammation model, to assay the PMN immediate response and infiltration in vivo. For these experiments, healthy C57BL/6 mice that were maintained in a pathogen-free animal facility were injected i.p. with zymosan A, followed by analysis for PMN recruitment into the peritoneum after various time periods. As shown in Fig. 1A and Table I, before zymosan injection, ∼2.6 × 106 cells were present in the peritoneum, and among these cells, ∼30% were F4/80+ macrophages, and the remainder contained few (∼5%) Gr-1+, which presumably were tissue granulocytes and other tissue cells. Soon after zymosan challenge, PMN began to infiltrate into the peritoneum, resulting in drastic changes in the cell composition and numbers. As shown, at 2 h, small numbers of PMN (∼1 × 106) transmigrated across the endothelium and arrived at the peritoneum. Meanwhile, the majority of macrophages were seemingly leaving the area, resulting in a significant decrease of the F4/80+ cells (Table I). Massive PMN infiltration into the peritoneum occurred after 2 h, resulting in ∼107 PMN and >1.2 × 107 PMN recruited to the location at 4 and 6 h, respectively. By 8 h, PMN infiltration slowed rapidly or stopped, resulting in a decrease in the number of PMN in the area. After the peak of PMN recruitment, monocyte infiltration began to increase at 6 h and continued to increase thereafter, suggesting that the acute inflammation proceeded to a resolving stage. The infiltrating monocytes and their mediated clearance likely accounted for the decrease of PMN numbers in the peritoneum at later time points. Consistent with other reports (13, 14), we detected a robust but transient increase in IL-6 in the peritoneal fluid and in the blood serum (Fig. 1B, Table II) after zymosan challenge. As observed, the time course of IL-6 change tightly correlated with the dynamics of PMN infiltration, suggesting a potential role of this cytokine in controlling the episode of zymosan-induced peritonitis. Moderate increases in IL-1, IL-17A, and TNF-α were also detected. The dynamics of IL-1 resembled that of IL-6, but IL-17A was only increased at later time periods, when the massive PMN transmigration was nearly completed (Fig. 1B, Table II).

FIGURE 1.

Inflammatory response/peritonitis after zymosan challenge. Mice receiving injection (i.p.) of zymosan A were euthanized at the indicated time points. A, Macrophages and PMN lavaged from the peritoneum were stained with anti-F4/80 (green) and anti–Gr-1 (red) Abs, respectively, and visualized under a fluorescent microscope. B, Serum concentrations of inflammatory cytokines during peritonitis were evaluated by ELISA. Data are expressed as means ± SEM; n = 6 mice/group. **p < 0.01 versus respective serum cytokine levels at 0 h.

FIGURE 1.

Inflammatory response/peritonitis after zymosan challenge. Mice receiving injection (i.p.) of zymosan A were euthanized at the indicated time points. A, Macrophages and PMN lavaged from the peritoneum were stained with anti-F4/80 (green) and anti–Gr-1 (red) Abs, respectively, and visualized under a fluorescent microscope. B, Serum concentrations of inflammatory cytokines during peritonitis were evaluated by ELISA. Data are expressed as means ± SEM; n = 6 mice/group. **p < 0.01 versus respective serum cytokine levels at 0 h.

Close modal
Table I.
Leukocytes in the peritoneum and their dynamic changes during zymosan-induced peritonitis
Cell Number (Mean ± SEM)
Time (h)F4/80+ (× 105)Gr-1+ (× 105)Total (× 105)
10.4 ± 1.4 2.0 ± 0.7 26.4 ± 3.7 
1.5 ± 0.2 10.7 ± 1.5 15.3 ± 2.2 
0.9 ± 0.1 96.2 ± 6.8 120.5 ± 8.5 
1.5 ± 0.2 124.3 ± 10.4 146.2 ± 12.2 
8.2 ± 1.1 115.1 ± 6.6 154.3 ± 8.8 
Cell Number (Mean ± SEM)
Time (h)F4/80+ (× 105)Gr-1+ (× 105)Total (× 105)
10.4 ± 1.4 2.0 ± 0.7 26.4 ± 3.7 
1.5 ± 0.2 10.7 ± 1.5 15.3 ± 2.2 
0.9 ± 0.1 96.2 ± 6.8 120.5 ± 8.5 
1.5 ± 0.2 124.3 ± 10.4 146.2 ± 12.2 
8.2 ± 1.1 115.1 ± 6.6 154.3 ± 8.8 

Mice injected with zymosan A were euthanized at various time points followed by collection cells from peritonea and staining for macrophages and PMN/granulocytes using an anti-F4/80 Ab (green) and an anti–Gr-1(red) Ab, respectively. Control cell staining used normal rat serum. The cells of different staining were counted and analyzed by FACS. The 0 h time point was determined without zymosan injection. The table shows the mean cell number ± SEM, n = 6 mice/group.

Table II.
Time-course detection of serum cytokines during zymosan A-induced peritonitis
Time Post-Zymosan Injection (h)
Cytokine (pg/ml)02468
TNF-α 49.2 ± 10.3 118.1 ± 20.6* 111.8 ± 16.0* 96.4 ± 12.8* 100.7 ± 16.5* 
IL-1β 31.0 ± 6.6 159.3 ± 26.5* 498.0 ± 58.9** 113.7 ± 32.5* 64.3 ± 22.7 
IL-6 65.6 ± 22.8 2180.0 ± 387.9** 4713.1 ± 1333.4** 1023.7 ± 322.4** 555.0 ± 165.6** 
IL-17A 109.6 ± 10.8 126.9 ± 12.7 370.3 ± 94.5** 562.0 ± 77.4** 517.3 ± 64.6** 
Time Post-Zymosan Injection (h)
Cytokine (pg/ml)02468
TNF-α 49.2 ± 10.3 118.1 ± 20.6* 111.8 ± 16.0* 96.4 ± 12.8* 100.7 ± 16.5* 
IL-1β 31.0 ± 6.6 159.3 ± 26.5* 498.0 ± 58.9** 113.7 ± 32.5* 64.3 ± 22.7 
IL-6 65.6 ± 22.8 2180.0 ± 387.9** 4713.1 ± 1333.4** 1023.7 ± 322.4** 555.0 ± 165.6** 
IL-17A 109.6 ± 10.8 126.9 ± 12.7 370.3 ± 94.5** 562.0 ± 77.4** 517.3 ± 64.6** 

Concentrations of inflammatory cytokines in serum during peritonitis were evaluated by ELISA. Data are expressed as means ± SEM; n = 6 mice/group.

*p < 0.05, **p < 0.01 versus respective serum cytokine levels at 0 h.

To investigate whether chronic inflammation affects PMN infiltration, we induced various inflammatory conditions in C57BL/6 mice before performing the zymosan-peritonitis assay. In these experiments, mice were treated with 2% DSS to induce colitis (7, 8), STZ to induce type 1 diabetes (9, 10), or LPS to induce bronchitis/lung inflammation (15). A group of mice were also treated with one dose of emulsified CFA (50% in PBS) to induce chronic inflammation. As shown in Fig. 2A, progressive body weight loss, which was associated with diarrhea and colitis, was observed in DSS-treated mice. Staining of colon tissues after treating the mice with DSS demonstrated significant intestinal inflammation that was associated with aggressive leukocyte infiltration, severe mucosal damage, and loss of epithelial structures (Fig. 2A). Treating mice with STZ and LPS also successfully induced diabetic and lung inflammatory conditions. As shown in Fig. 2B, all of the STZ-treated mice developed hyperglycemia, with the mean blood glucose level being increased to 431.7 ± 56.1 mg/dl, whereas the control group treated with vehicle (PBS) maintained a blood glucose level of 141.2 ± 14.0 mg/dl. Meanwhile, treating mice with low-dose LPS via the nares for 15 d induced evident upper respiratory inflammation that was characterized by noticeable leukocyte infiltration around bronchi, bronchioles, and in some area of the lung parenchyma (Fig. 2C).

FIGURE 2.

Mice with various chronic inflammatory conditions display enhanced PMN infiltration during zymosan-induced peritonitis. A, DSS-induced colitis. C57BL/6J mice were given 2% DSS in drinking water continuously, and development of colitis was monitored by body weight loss, diarrhea, etc., and confirmed by dissection of the colonic tissues. B, STZ-induced type 1 diabetes. Mice were consecutively injected with STZ or the vehicle (CTL) for 5 d. Serum glucose levels were assayed 3 wk after the injections. C, Inducing upper respiratory inflammation by LPS. LPS or vehicle was given to mice via the nares every other day for 15 d. The figure shows the histology images (H&E staining) of lung tissue sections after vehicle (Ctl.) or LPS treatment. D and E, Enhanced PMN infiltration in mice with systemic inflammation. Control healthy mice and mice with colitis (DSS for 12 d), hyperglycemia (3 wk post-STZ treatment), or lung inflammation (LPS) were tested for PMN infiltration by zymosan A-induced peritonitis. Mice that received one dose of emulsified CFA (50% in PBS) were also tested in the assay. D and E show the numbers of PMN infiltrated into the peritoneum at 2 h and the kinetics of PMN infiltration into the peritoneum, respectively. The data represent one of eight independent experiments and are expressed as means ± SEM; n = mice number/group. **p < 0.01, ***p < 0.001 versus respective controls.

FIGURE 2.

Mice with various chronic inflammatory conditions display enhanced PMN infiltration during zymosan-induced peritonitis. A, DSS-induced colitis. C57BL/6J mice were given 2% DSS in drinking water continuously, and development of colitis was monitored by body weight loss, diarrhea, etc., and confirmed by dissection of the colonic tissues. B, STZ-induced type 1 diabetes. Mice were consecutively injected with STZ or the vehicle (CTL) for 5 d. Serum glucose levels were assayed 3 wk after the injections. C, Inducing upper respiratory inflammation by LPS. LPS or vehicle was given to mice via the nares every other day for 15 d. The figure shows the histology images (H&E staining) of lung tissue sections after vehicle (Ctl.) or LPS treatment. D and E, Enhanced PMN infiltration in mice with systemic inflammation. Control healthy mice and mice with colitis (DSS for 12 d), hyperglycemia (3 wk post-STZ treatment), or lung inflammation (LPS) were tested for PMN infiltration by zymosan A-induced peritonitis. Mice that received one dose of emulsified CFA (50% in PBS) were also tested in the assay. D and E show the numbers of PMN infiltrated into the peritoneum at 2 h and the kinetics of PMN infiltration into the peritoneum, respectively. The data represent one of eight independent experiments and are expressed as means ± SEM; n = mice number/group. **p < 0.01, ***p < 0.001 versus respective controls.

Close modal

We then examined PMN function in these mice with various inflammatory conditions by performing zymosan-induced peritonitis. For these experiments, we used the mice that had been treated with DSS for 12 d and manifested apparent colitic condition. As shown in Fig. 2D, compared with the control healthy mice that PMN just began to infiltrate into the peritoneum at 2 h and only small numbers (∼1 × 106 PMN) were detected, significantly increased numbers of PMN (>6 × 106) were recruited to the area in colitic mice at the same time point. Similar early promoted PMN infiltration into the peritoneum in response to zymosan challenge was also observed in STZ-induced diabetic mice and LPS-treated mice. These mice were used to test for PMN infiltration by peritonitis after the STZ- and LPS-induced pathological conditions were stabilized. As shown in Fig. 2D, at 2 h, 5.6 × 106 PMN and ∼3 × 106 PMN were recruited to peritonea in STZ- and LPS-treated mice, respectively. We also induced peritonitis in the mice that were treated with CFA 1 mo ago. As shown in Fig. 2D, this form of chronic inflammation also significantly enhanced PMN response to zymosan, resulting in >7 × 106 PMN being recruited to the peritoneum by 2 h.

Kinetic analysis of PMN infiltration (Fig. 2E) in colitic mice by plotting the total numbers of PMN transmigrated as a function of time showed that the robust PMN infiltration occurred earlier than that in the control mice. Indeed, >70% of the infiltrated PMN in mice with DSS treatment for 12 d had arrived at the peritoneum by 2 h, whereas in the controls, the majority of PMN were recruited by 4 h. Significant enhanced PMN infiltration into the peritoneum at an early time point (2 h) was also observed in mice with DSS treatment for 9 d (Fig. 2E). The kinetic results also indicated that the early enhancement of PMN infiltration in colitic mice was not due to inflammation-induced increases in PMN production but to accelerated PMN response and transmigration. In fact, the accumulative PMN numbers that infiltrated into the peritoneum in colitic mice at 4 h were not higher, but tended to be lower, than those in the controls. Parallel analyses of PMN numbers in the peripheral circulation and in bone marrow indicated that despite the increase of GM-CSF in the serum (Table III), continual DSS treatment (≥9 d) and colitis exhausted the entire system, and also consumed PMN availability in the circulation (data not shown).

Table III.
Serum cytokine levels during DSS-induced colitis
DSS Treatment
Cytokine (pg/ml)Day 0Day 3Day 6Day 9Day 12
TNF-α 52.8 ± 5.6 71.0 ± 5.8* 69.7 ± 8.1 57.8 ± 11.6 52.1 ± 9.3 
IL-1β 35.4 ± 8.6 28.5 ± 6.0 39.4 ± 11.2 227.5 ± 22.7** 144.6 ± 16.4** 
IL-6 70.2 ± 17.6 78.7 ± 10.4 132.2 ± 42.4* 430.8 ± 109.1** 173.4 ± 24.0** 
IL-17A 113.6 ± 10.6 108.9 ± 10.9 249.8 ± 24.1* 714.1 ± 93.6** 2023.9 ± 172.1*** 
MCP-1 30.3 ± 8.6 37.2 ± 9.4 156.8 ± 16.2** 79.4 ± 18.2* 22.5 ± 4.3 
IFN-γ 51.0 ± 6.7 44.2 ± 7.8 51.1 ± 9.3 40.4 ± 18.0 41.2 ± 6.4 
IL-12/23(p40) 55.6 ± 7.3 57.8 ± 6.5 77.4 ± 11.9* 67.0 ± 6.1 54.6 ± 7.2 
IL-4 39.5 ± 4.2 43.6 ± 7.3 ND ND ND 
IL-10 59.2 ± 20.9 133.6 ± 16.4* 35.5 ± 12.8 30.4 ± 11.3 27.2 ± 13.6 
GM-CSF ND 65.0 ± 16.3* 29.4 ± 9.2 21.0 ± 8.4 ND 
DSS Treatment
Cytokine (pg/ml)Day 0Day 3Day 6Day 9Day 12
TNF-α 52.8 ± 5.6 71.0 ± 5.8* 69.7 ± 8.1 57.8 ± 11.6 52.1 ± 9.3 
IL-1β 35.4 ± 8.6 28.5 ± 6.0 39.4 ± 11.2 227.5 ± 22.7** 144.6 ± 16.4** 
IL-6 70.2 ± 17.6 78.7 ± 10.4 132.2 ± 42.4* 430.8 ± 109.1** 173.4 ± 24.0** 
IL-17A 113.6 ± 10.6 108.9 ± 10.9 249.8 ± 24.1* 714.1 ± 93.6** 2023.9 ± 172.1*** 
MCP-1 30.3 ± 8.6 37.2 ± 9.4 156.8 ± 16.2** 79.4 ± 18.2* 22.5 ± 4.3 
IFN-γ 51.0 ± 6.7 44.2 ± 7.8 51.1 ± 9.3 40.4 ± 18.0 41.2 ± 6.4 
IL-12/23(p40) 55.6 ± 7.3 57.8 ± 6.5 77.4 ± 11.9* 67.0 ± 6.1 54.6 ± 7.2 
IL-4 39.5 ± 4.2 43.6 ± 7.3 ND ND ND 
IL-10 59.2 ± 20.9 133.6 ± 16.4* 35.5 ± 12.8 30.4 ± 11.3 27.2 ± 13.6 
GM-CSF ND 65.0 ± 16.3* 29.4 ± 9.2 21.0 ± 8.4 ND 

Inflammatory cytokine levels in serum during DSS-induced colitis were evaluated by ELISA. Data are expressed as means ± SEM; n = 6-8 mice/group. ND, undetectable.

*p < 0.05, **p < 0.01, ***p < 0.001 versus respective controls.

The results of enhanced PMN infiltration in mice under various inflammatory conditions suggest a likelihood that alterations occurred in either PMN, which directly promote PMN transmigration, or in cells within the peritoneal vicinity, which led to an earlier local response to zymosan and/or contributed to the facilitation when PMN transmigrated through. To test these possibilities, we designed two directional, adoptive leukocyte transfer and peritonitis experiments. In these, donor PMN derived from mice with different inflammatory conditions were transfused into healthy, noninflammatory recipient mice and tested for response and infiltration in zymosan-induced peritonitis; conversely, donor PMN from healthy mice were transfused into recipient mice with ongoing inflammation and then tested for infiltration in response to zymosan challenging. The former experiments determine whether chronic inflammation converts PMN to be faster migrating cells (even in a new host environment). The latter experiments examine the peritoneal tissue microenvironment and determine whether chronic inflammation could induce changes that promote PMN infiltration during peritonitis. Control experiments were performed by transfusion of PMN from healthy mice into other healthy mice followed by testing the donor PMN infiltration.

Fig. 3A shows the results of the transfusion of PMN from inflammatory mice into healthy recipients and the infiltration of donor PMN under zymosan challenge. As shown, although the host mice had no prior inflammation, donor PMN from colitic and diabetic mice (Fig. 3A, labeled with CFSE, green) still displayed much faster infiltration, resulting in more cells transmigrated into the peritoneum at 2 h than control donor PMN from healthy mice. In parallel, we also performed in vitro chemotaxis assays and compared the transmigration of PMN derived from DSS-treated mice and from healthy mice. To avoid activation of PMN, especially those from mice with inflammation, we directly applied the total bone marrow leukocytes without further isolation. As shown in Fig. 3B, greater numbers of PMN transmigrating across matrix-coated filters toward the chemoattractant fMLF were observed in samples derived from DSS-treated mice than those from healthy mice. In Fig. 3C, we further labeled donor PMN or total bone marrow cells derived from DSS-treated mice and from healthy mice with different dyes (green and red) and simultaneously transfused into the same recipient mice (at a 1:1 ratio of the PMN number). As shown, zymosan induced more infiltration of PMN from colitic mice (green) than those from noncolitic, healthy mice (red) at 2 h, confirming that PMN in chronic inflammatory mice had an enhanced ability to respond and migrate toward inflammatory stimuli.

FIGURE 3.

Systemic inflammation alters both PMN and tissue microenvironments. A, Testing donor PMN from inflammatory mice response in healthy recipients by zymosan-peritonitis. Bone marrow cells (2.5 × 106 mature PMN) derived from colitic (DSS, 12 d), diabetic (STZ), or control healthy mice were labeled with CFSE and transfused into healthy recipients, respectively. Figure shows the microscopic analysis of infiltrated donor PMN in peritoneal lavage after inducing peritonitis for 2 h and the total numbers of donor PMN infiltrated at 2 and 4 h. The result represents one of four independent experiments. B, PMN chemotaxis in vitro. Bone marrow leukocytes from healthy control and colitic (DSS, 12 d) mice were normalized for equal PMN contents by MPO assays before being applied into the upper chambers of the transmigration setups. Chemotaxis induced by fMLF was measured at 45 min. C, Cotransfer of leukocytes from healthy and inflammatory mice and testing by peritonitis. A mixture of bone marrow leukocytes derived from healthy mice (red, DiIC16 labeling) and colitic mice (DSS, 12 d) (green, CFSE labeling) (2.5 × 106 PMN each) were transferred into the same healthy recipients. The images show zymosan-induced PMN infiltration into the peritoneum at 2 h in the peritoneal lavage. The result represents one of five independent experiments. D, Transfer of PMN from healthy mice into inflammatory recipients followed by testing PMN infiltration during peritonitis. Bone marrow leukocytes from five healthy mice were combined and labeled, followed by transfusion into different types of inflammatory recipient mice (three mice per type, 2.5 × 106 PMN/mouse). E, Peritoneal macrophages obtained by lavaging peritonea were stimulated in vitro using zymosan (1 mg/ml) in DMEM (containing 5% FBS) at 37°C. IL-6 secreted into the medium at different time points was assayed by ELISA. The data are expressed as means ± SEM. *p < 0.05, **p < 0.01 versus respective controls.

FIGURE 3.

Systemic inflammation alters both PMN and tissue microenvironments. A, Testing donor PMN from inflammatory mice response in healthy recipients by zymosan-peritonitis. Bone marrow cells (2.5 × 106 mature PMN) derived from colitic (DSS, 12 d), diabetic (STZ), or control healthy mice were labeled with CFSE and transfused into healthy recipients, respectively. Figure shows the microscopic analysis of infiltrated donor PMN in peritoneal lavage after inducing peritonitis for 2 h and the total numbers of donor PMN infiltrated at 2 and 4 h. The result represents one of four independent experiments. B, PMN chemotaxis in vitro. Bone marrow leukocytes from healthy control and colitic (DSS, 12 d) mice were normalized for equal PMN contents by MPO assays before being applied into the upper chambers of the transmigration setups. Chemotaxis induced by fMLF was measured at 45 min. C, Cotransfer of leukocytes from healthy and inflammatory mice and testing by peritonitis. A mixture of bone marrow leukocytes derived from healthy mice (red, DiIC16 labeling) and colitic mice (DSS, 12 d) (green, CFSE labeling) (2.5 × 106 PMN each) were transferred into the same healthy recipients. The images show zymosan-induced PMN infiltration into the peritoneum at 2 h in the peritoneal lavage. The result represents one of five independent experiments. D, Transfer of PMN from healthy mice into inflammatory recipients followed by testing PMN infiltration during peritonitis. Bone marrow leukocytes from five healthy mice were combined and labeled, followed by transfusion into different types of inflammatory recipient mice (three mice per type, 2.5 × 106 PMN/mouse). E, Peritoneal macrophages obtained by lavaging peritonea were stimulated in vitro using zymosan (1 mg/ml) in DMEM (containing 5% FBS) at 37°C. IL-6 secreted into the medium at different time points was assayed by ELISA. The data are expressed as means ± SEM. *p < 0.05, **p < 0.01 versus respective controls.

Close modal

To investigate whether the tissue microenvironment also contributed to the enhanced PMN infiltration under inflammatory conditions, we transferred the PMN of healthy mice into DSS- and STZ-treated chronic inflammatory mice (recipients). Inducing peritonitis subsequently showed significantly faster infiltration of donor PMN in mice with ongoing colitis or diabetes than in control healthy recipients (Fig. 3D). These results suggest that considerable alterations had occurred in the peritoneal tissue environment in inflammatory mice and that these changes promoted PMN infiltration. Because peritoneal macrophages play an important role in the development of peritonitis, we collected these macrophages and directly tested their responses to zymosan in vitro. As shown in Fig. 3E, in the cell-culture dishes, zymosan induced increased IL-6 production in macrophages collected from DSS- and STZ-treated mice than those isolated from noninflammatory controls. Interestingly, we observed significantly increased numbers of macrophages in the peritoneum of STZ-induced diabetic mice (data not shown). Thus, these results confirmed tissue environment changes under chronic inflammation. Increases in cytokine production from the local macrophages certainly contributed to enhanced PMN infiltration.

The findings that circulating PMN and peritoneal macrophages had altered responsiveness under colitis and diabetes were striking given that both cells were distal to the original inflammatory sites in those conditions. We thus hypothesized that critical inflammatory factors were released into the circulation, through which it reached and regulated cells at a distance. To identify these important factors, especially those that might contribute to enhance PMN infiltration, we analyzed inflammatory cytokines associated with DSS-induced colitis in the serum. As shown in Table III, whereas most of cytokines tested were reported in colonic tissues (7, 1619), these cytokines stayed at low or moderate levels in the serum throughout the course of the DSS treatment. The only cytokine that we found significantly elevated in the serum was IL-17A (Fig. 4A), which started to increase at days 6–8 of DSS treatment and continued increasing, eventually reaching >2 ng/ml. We found that not only was its elevation in the serum extraordinary, but also that the dynamics of IL-17A tightly correlated with the enhancement of PMN infiltration observed in peritonitis. As shown in Fig. 4B, inducing peritonitis in mice that were treated with DSS for different days revealed that significant promotion of PMN infiltration occurred only after DSS treatment for 6 d and continued to be enhanced thereafter. Supporting this observation, assay of peritonitis-associated IL-6 release also demonstrated that enhancement of IL-6 release upon zymosan challenging occurred in mice with DSS treatment for >6 d (Fig. 4C). Interestingly, significant body weight decrease in DSS-treated mice also started at approximately the same time (Figs. 2A, 5C). These results suggest that, although DSS-induced colitis started much earlier than day 6, as elevations of inflammatory cytokines (Table III) and leukocyte infiltration into the colonic tissues were apparent on days 2–5 (Fig. 4D), the quick collapse of the pathological condition at the later stage was associated with the enhancement of PMN infiltration and the consequent severe tissue damage.

FIGURE 4.

Enhancement of PMN infiltration in DSS-induced colitis is correlated with IL-17A production. A, Time-course analysis of serum cytokines during DSS-induced colitis (also see Table III). B, Analysis of PMN infiltration by zymosan-induced peritonitis in mice treated with DSS for different days. Note that significantly enhanced PMN infiltration occurs after 6 d of DSS treatment and in parallel with increases in IL-17A. C, Analysis of peritonitis-associated IL-6 production in mice treated with DSS for different days. D, Tissue section of colon intestines and labeling for infiltrated PMN using an anti–Gr-1 Ab. The data represent one of at least three independent experiments and are expressed as means ± SEM with at least five mice per group. *p < 0.05, **p < 0.01, ***p < 0.001 versus respective controls.

FIGURE 4.

Enhancement of PMN infiltration in DSS-induced colitis is correlated with IL-17A production. A, Time-course analysis of serum cytokines during DSS-induced colitis (also see Table III). B, Analysis of PMN infiltration by zymosan-induced peritonitis in mice treated with DSS for different days. Note that significantly enhanced PMN infiltration occurs after 6 d of DSS treatment and in parallel with increases in IL-17A. C, Analysis of peritonitis-associated IL-6 production in mice treated with DSS for different days. D, Tissue section of colon intestines and labeling for infiltrated PMN using an anti–Gr-1 Ab. The data represent one of at least three independent experiments and are expressed as means ± SEM with at least five mice per group. *p < 0.05, **p < 0.01, ***p < 0.001 versus respective controls.

Close modal
FIGURE 5.

Suppression of IL-17A function by a neutralization Ab eliminates the augmentation of inflammation and PMN infiltration at postacute stage of DSS-induced colitis. Mice under DSS treatment were administered (i.v.) 30 μg low endotoxin, azide-free–purified anti-mouse IL-17A Ab on days 4, 6, 8, and 10. Control injections were performed by administration of an isotype-matched Ab (rat IgG1) (BioLegend). On day 12, mice with and without Ab administration were tested by zymosan-induced peritonitis. A, PMN infiltration into the peritoneum at 2 and 4 h. B, Serum IL-6 after zymosan challenge. C, The body weight of mice treated with free water or DSS plus Abs. The data are expressed as means ± SEM and represent one of at least three independent experiments with at least five mice per group. **p < 0.01, ***p < 0.001 versus respective controls.

FIGURE 5.

Suppression of IL-17A function by a neutralization Ab eliminates the augmentation of inflammation and PMN infiltration at postacute stage of DSS-induced colitis. Mice under DSS treatment were administered (i.v.) 30 μg low endotoxin, azide-free–purified anti-mouse IL-17A Ab on days 4, 6, 8, and 10. Control injections were performed by administration of an isotype-matched Ab (rat IgG1) (BioLegend). On day 12, mice with and without Ab administration were tested by zymosan-induced peritonitis. A, PMN infiltration into the peritoneum at 2 and 4 h. B, Serum IL-6 after zymosan challenge. C, The body weight of mice treated with free water or DSS plus Abs. The data are expressed as means ± SEM and represent one of at least three independent experiments with at least five mice per group. **p < 0.01, ***p < 0.001 versus respective controls.

Close modal

To further explore the role of IL-17A in the enhancement of PMN infiltration, we treated the DSS mice with a functional neutralization Ab against IL-17A (20, 21) (i.v.). As shown in Fig. 5, suppression of IL-17A function by the Ab in DSS-treated mice nearly completely eliminated the enhanced PMN infiltration at 2 h during zymosan-induced peritonitis (Fig. 5A) and such event-associated increases in IL-6 (Fig. 5B). The colitic symptoms in these DSS-treated mice, such as body weight loss, diarrhea, and intestinal tissue damage, were also largely alleviated (Fig. 5C). However, IL-17A neutralization had no effect on the basal level of PMN infiltration, as large numbers of PMN were still recruited into the peritoneum at 4 h during peritonitis even with the Ab administration (Fig. 5A). PMN infiltration into intestinal mucosa was also present in DSS-treated mice with anti–IL-17A administration, but remained at a low level (data not shown). We further performed ex vivo experiments and tested whether exposure to IL-17A directly modulates PMN function. In these experiments, bone marrow leukocytes isolated from healthy mice were ex vivo treated with a functional rIL-17A and then transferred into other healthy mice (recipients) followed by testing by zymosan-induced peritonitis. As shown in Fig. 6A, PMN treated with rIL-17A demonstrated markedly accelerated infiltration into the peritoneum during peritonitis compared with nontreated control PMN that were cotransferred into the same recipients. As expected, the presence of the anti–IL-17A Ab in the treatment completely eliminated the enhancement of PMN infiltration. Similarly, PMN that were treated with the serum from colitic mice (DSS treatment for 12 d) also manifested accelerated infiltration in recipient mice during peritonitis. Consistent with these results, assaying PMN chemotaxis in vitro (Fig. 6B) observed significantly augmented chemotactic transmigration for PMN treated with rIL-17A, whereas neutralization of IL-17A function erased such augmentation. Because tissue macrophage responses were also altered under inflammatory conditions, we tested whether IL-17A also directly modulates macrophage function. As shown in Fig. 6C, treatment of peritoneal macrophages isolated from healthy mice with rIL-17A markedly elevated IL-6 production upon the subsequent stimulation with zymosan. Western blot analyses of IL-17A–treated macrophages indicated significantly increased protein tyrosine phosphorylations (Fig. 6D), suggesting that active signaling events were elicited by IL-17A. In addition, we also observed direct binding of rIL-17A to the cell surfaces of PMN and macrophages (not shown), presumably to the receptors. In conclusion, these results strongly suggest that IL-17A, which was produced at a large amount in the postacute/chronic stage of colitis, serves as a critical feedback factor and contributes to reinforcement of the inflammatory reaction through systemically modulating leukocytes in the circulation and tissues.

FIGURE 6.

IL-17A directly enhances PMN and macrophage inflammatory functions. A, PMN ex vivo exposure to IL-17A exhibits an accelerated infiltration during zymosan-induced peritonitis. Bone marrow leukocytes from healthy mice were treated for 2 h with an rIL-17A (2 ng/ml) or serum from mice with DSS-induced colitis (DSS, 12 d) in the presence or absence of the neutralization anti–IL-17A Ab. After treatment, the cells were labeled with CSFE (green) and cotransferred with control nontreated bone marrow leukocytes (red) into healthy recipient mice followed by testing for PMN infiltration during zymosan-induced peritonitis. B, IL-17A directly enhances PMN chemotactic transmigration. Bone marrow PMN with and without exposure to IL-17A were assayed in vitro for chemotaxis toward fMLF for 90 min. C, IL-17A promotes IL-6 production in macrophage upon stimulation. Peritoneal macrophages obtained from healthy mice were treated with rIL-17A (2 ng/ml) with or without the presence of anti–IL-17A Ab for 48 h (37°C). IL-6 production in cells with or without zymosan stimulation (2 h) was assayed by ELISA. D, Western blot analyses of protein phosphorylation of macrophages treated with IL-17A. **p < 0.01.

FIGURE 6.

IL-17A directly enhances PMN and macrophage inflammatory functions. A, PMN ex vivo exposure to IL-17A exhibits an accelerated infiltration during zymosan-induced peritonitis. Bone marrow leukocytes from healthy mice were treated for 2 h with an rIL-17A (2 ng/ml) or serum from mice with DSS-induced colitis (DSS, 12 d) in the presence or absence of the neutralization anti–IL-17A Ab. After treatment, the cells were labeled with CSFE (green) and cotransferred with control nontreated bone marrow leukocytes (red) into healthy recipient mice followed by testing for PMN infiltration during zymosan-induced peritonitis. B, IL-17A directly enhances PMN chemotactic transmigration. Bone marrow PMN with and without exposure to IL-17A were assayed in vitro for chemotaxis toward fMLF for 90 min. C, IL-17A promotes IL-6 production in macrophage upon stimulation. Peritoneal macrophages obtained from healthy mice were treated with rIL-17A (2 ng/ml) with or without the presence of anti–IL-17A Ab for 48 h (37°C). IL-6 production in cells with or without zymosan stimulation (2 h) was assayed by ELISA. D, Western blot analyses of protein phosphorylation of macrophages treated with IL-17A. **p < 0.01.

Close modal

PMN, which comprise more than two-thirds of peripheral leukocytes, are the first-tier responsive cells during inflammation. As shown in zymosan-induced peritonitis, when a stimulatory signal is effectively produced, PMN rapidly transmigrate through the vasculature and tissues and arrive at the inflammatory site within a few hours. After a short period and vigorous activity, PMN infiltration is promptly ceased, provided that the inflammatory causes are largely cleared, to confine and limit tissue damage. However, when the inflammatory causes linger, as those in DSS-induced colitis, PMN infiltration continues, and the inflammatory reaction prolongs and becomes chronic. In this study, we employed inflammation mouse models and studied PMN infiltration under various chronic conditions. We found that animals that were preconditioned with nonresolving colitis, type 1 diabetes, or other inflammatory conditions had significantly enhanced PMN infiltration during zymosan-induced peritonitis. Cross-transfer of leukocytes from mice with ongoing inflammation to healthy recipients, or from healthy mice to inflammatory recipients, further revealed that both circulating PMN and tissue microenvironments were altered under chronic inflammatory conditions and that these changes collectively contributed to the promotion of PMN infiltration during peritonitis.

Detailed analysis of DSS-induced colitis found that PMN infiltration into intestinal mucosa occurred shortly after the starting of DSS treatment (days 2 to 3); concomitantly, a panel of inflammatory cytokines associated with colitis, including low levels of IL-17A, which are presumably released from the local γδ T cells and infiltrated PMN (22), were also produced (Table III). However, when the DSS treatment continued over 6–8 d, we observed extraordinarily high levels of IL-17A in the blood serum. Our studies suggest that this late-produced strong wave of IL-17A, which was likely through the activation of the second-tier inflammatory cells such as Th17 cells, is capable of reaching and regulating cells distant from the original inflammatory site through circulation, leading to systemic modulations of the entire inflammation and homeostasis. Indeed, our data showed that both macrophages and PMN in distant tissues are regulated by such released, circulating IL-17A. In fact, PMN appeared to be altered even before their release from the bone marrow, consistent with the microarray study by Theilgaard-Monch et al. (23, 24) in which various cytokine receptors including IL-17R were identified in bone marrow myelocytes, especially mature PMN. Strikingly, the time course of elevation of IL-17A in serum during DSS treatment tightly correlated with the significant enhancement of PMN infiltration observed in zymosan-induced peritonitis. Subsequent in vitro and ex vivo experiments confirmed that IL-17A directly modulates PMN and significantly accelerated PMN chemotactic transmigration toward inflammatory stimuli. Not only modulating PMN, IL-17A also enhances the responses and sensitivities of macrophages around the body. In vitro exposure of macrophages to IL-17A prior to stimulation with zymosan resulted in much increased production of the proinflammatory cytokine IL-6. Peritoneal macrophages isolated from late-stage colitic mice also demonstrated enhanced responses upon inflammatory challenge. All of these results together indicate that when inflammation prolongs, feedback signals are produced and trigger a systemic regulation of the entire inflammatory reaction. In the case of DSS-induced colitis, large amounts of IL-17A produced at the postacute stage prime circulating PMN and tissue macrophages and enhance their inflammatory functions. These regulations by IL-17A, which are presumably designed to accelerate the removal of the original inflammatory cause, nevertheless lead to exaggerated inflammation-associated tissue damage. Suppression of IL-17A–mediated propagation of inflammation mitigates the disease condition.

Priming PMN under inflammatory conditions and by inflammatory factors has been observed in research through the years and has been considered to be a major factor responsible for the pathogenesis of organ failure occurred under postinjury conditions such as trauma, burn, and surgery and other conditions such as adult respiratory distress syndrome, sepsis, cystic fibrosis, blood transfusion, etc. (2532). Reported agents that prime PMN include platelet-activating factor (33, 34), G-CSF (35, 36), GM-CSF (37), IL-1β (38), IL-6 (39), IL-8 (35), etc. In this study, our data suggest that IL-17A is likely to be a critical priming agent for PMN and macrophages under inflammatory conditions, yet the mechanism underlying its effect and the phenotypic characteristics of PMN (40) after exposure to IL-17A need to be further determined. Although many studies have shown that IL-17A production is regulated by IL-6 and IL-23 (and other cytokines) (41, 42), our study in DSS-induced colitis only detected slight increases of these cytokines in the serum, implicating that IL-6– and IL-23–mediated activation of Th17 cells may be mainly restricted in the local intestinal tissues. In addition, previous studies by Schwarzenberger et al. (43, 44) and others (45, 46) have shown that increases of IL-17A promote promyelocyte proliferation and PMN maturation. Because IL-17A was significantly increased in DSS-treated mice, it is thus likely that DSS-induced colitis was associated with increased granulopoiesis and PMN production, which are important for the ongoing inflammation and continual PMN infiltration into intestines. However, increases of PMN production under colitis were not the reason for the enhanced PMN infiltration observed during peritonitis, which instead was rather due to PMN priming and accelerated recruitment. Indeed, we detected only minimal PMN in the peripheral circulation and no increase of mature PMN in bone marrow in mice treated with DSS for ≥9 d, suggesting that incessant colitis has consumed majority of PMN and largely depleted the reservoirs.

Thus, through this study, we have learned of important mechanisms that regulate the dynamics of inflammation. Our results imply that when an inflammatory condition becomes unrelenting, such as in inflammatory bowel disease, diabetes, and rheumatoid arthritis, significantly enhanced inflammatory responses and PMN infiltration can occur and lead to severe tissue injury and deteriorated pathophysiological condition. In addition, systemically enhanced inflammatory status also renders the chronic patients highly vulnerable when they face a new inflammatory challenge, such as viral/bacterial infection or cancer, during which exaggerated reactions can prevail and cause overstated consequences.

This work was supported in part by National Institutes of Health Grant DK62894 (to Y.L.) and a Research Scholar grant from the American Cancer Society (to Y.L.).

Abbreviations used in this article:

DiIC16(3)

1,1'-dihexadecyl-3,3,3′,3′-tetramethylindocarbocyanine iodide

DSS

dextran sulfate sodium

MPO

myeloperoxidase

PMN

polymorphonuclear leukocyte

rIL-17A

recombinant mouse IL-17A

STZ

streptozotocin.

1
Chen
C. X.
,
Soto
I.
,
Guo
Y. L.
,
Liu
Y.
.
2011
.
Control of secondary granule release in neutrophils by Ral GTPase.
J. Biol. Chem.
286
:
11724
11733
.
2
Liu
Y.
,
Merlin
D.
,
Burst
S. L.
,
Pochet
M.
,
Madara
J. L.
,
Parkos
C. A.
.
2001
.
The role of CD47 in neutrophil transmigration. Increased rate of migration correlates with increased cell surface expression of CD47.
J. Biol. Chem.
276
:
40156
40166
.
3
Zen
K.
,
Reaves
T. A.
,
Soto
I.
,
Liu
Y.
.
2006
.
Response to genistein: assaying the activation status and chemotaxis efficacy of isolated neutrophils.
J. Immunol. Methods
309
:
86
98
.
4
Chanchamroen
S.
,
Kewcharoenwong
C.
,
Susaengrat
W.
,
Ato
M.
,
Lertmemongkolchai
G.
.
2009
.
Human polymorphonuclear neutrophil responses to Burkholderia pseudomallei in healthy and diabetic subjects.
Infect. Immun.
77
:
456
463
.
5
Mowat
A.
,
Baum
J.
.
1971
.
Chemotaxis of polymorphonuclear leukocytes from patients with diabetes mellitus.
N. Engl. J. Med.
284
:
621
627
.
6
Pereira
M. A.
,
Sannomiya
P.
,
Leme
J. G.
.
1987
.
Inhibition of leukocyte chemotaxis by factor in alloxan-induced diabetic rat plasma.
Diabetes
36
:
1307
1314
.
7
Alex
P.
,
Zachos
N. C.
,
Nguyen
T.
,
Gonzales
L.
,
Chen
T. E.
,
Conklin
L. S.
,
Centola
M.
,
Li
X.
.
2009
.
Distinct cytokine patterns identified from multiplex profiles of murine DSS and TNBS-induced colitis.
Inflamm. Bowel Dis.
15
:
341
352
.
8
Okayasu
I.
,
Hatakeyama
S.
,
Yamada
M.
,
Ohkusa
T.
,
Inagaki
Y.
,
Nakaya
R.
.
1990
.
A novel method in the induction of reliable experimental acute and chronic ulcerative colitis in mice.
Gastroenterology
98
:
694
702
.
9
Salahuddin
M.
,
Jalalpure
S. S.
,
Gadge
N. B.
.
2010
.
Antidiabetic activity of aqueous bark extract of Cassia glauca in streptozotocin-induced diabetic rats.
Can. J. Physiol. Pharmacol.
88
:
153
160
.
10
Wysocki
J.
,
Ye
M.
,
Soler
M. J.
,
Gurley
S. B.
,
Xiao
H. D.
,
Bernstein
K. E.
,
Coffman
T. M.
,
Chen
S.
,
Batlle
D.
.
2006
.
ACE and ACE2 activity in diabetic mice.
Diabetes
55
:
2132
2139
.
11
Cash
J. L.
,
White
G. E.
,
Greaves
D. R.
.
2009
.
Chapter 17. Zymosan-induced peritonitis as a simple experimental system for the study of inflammation.
Methods Enzymol.
461
:
379
396
.
12
Dong
X.
,
Wu
D.
.
2006
.
Methods for studying neutrophil chemotaxis.
Methods Enzymol.
406
:
605
613
.
13
Soria-Castro
I.
,
Krzyzanowska
A.
,
Pelaéz
M. L.
,
Regadera
J.
,
Ferrer
G.
,
Montoliu
L.
,
Rodríguez-Ramos
R.
,
Fernández
M.
,
Alemany
S.
.
2010
.
Cot/tpl2 (MAP3K8) mediates myeloperoxidase activity and hypernociception following peripheral inflammation.
J. Biol. Chem.
285
:
33805
33815
.
14
McLoughlin
R. M.
,
Witowski
J.
,
Robson
R. L.
,
Wilkinson
T. S.
,
Hurst
S. M.
,
Williams
A. S.
,
Williams
J. D.
,
Rose-John
S.
,
Jones
S. A.
,
Topley
N.
.
2003
.
Interplay between IFN-γ and IL-6 signaling governs neutrophil trafficking and apoptosis during acute inflammation.
J. Clin. Invest.
112
:
598
607
.
15
Szarka
R. J.
,
Wang
N.
,
Gordon
L.
,
Nation
P. N.
,
Smith
R. H.
.
1997
.
A murine model of pulmonary damage induced by lipopolysaccharide via intranasal instillation.
J. Immunol. Methods
202
:
49
57
.
16
Yan
Y.
,
Kolachala
V.
,
Dalmasso
G.
,
Nguyen
H.
,
Laroui
H.
,
Sitaraman
S. V.
,
Merlin
D.
.
2009
.
Temporal and spatial analysis of clinical and molecular parameters in dextran sodium sulfate induced colitis.
PLoS ONE
4
:
e6073
.
17
Alex
P.
,
Ye
M.
,
Zachos
N. C.
,
Sipes
J.
,
Nguyen
T.
,
Suhodrev
M.
,
Gonzales
L.
,
Arora
Z.
,
Zhang
T.
,
Centola
M.
,
Guggino
S. E.
,
Li
X.
.
2010
.
Clcn5 knockout mice exhibit novel immunomodulatory effects and are more susceptible to dextran sulfate sodium-induced colitis.
J. Immunol.
184
:
3988
3996
.
18
Saubermann
L. J.
,
Nakajima
A.
,
Wada
K.
,
Zhao
S.
,
Terauchi
Y.
,
Kadowaki
T.
,
Aburatani
H.
,
Matsuhashi
N.
,
Nagai
R.
,
Blumberg
R. S.
.
2002
.
Peroxisome proliferator-activated receptor gamma agonist ligands stimulate a Th2 cytokine response and prevent acute colitis.
Inflamm. Bowel Dis.
8
:
330
339
.
19
Togawa
J.-I.
,
Nagase
H.
,
Tanaka
K.
,
Inamori
M.
,
Nakajima
A.
,
Ueno
N.
,
Saito
T.
,
Sekihara
H.
.
2002
.
Oral administration of lactoferrin reduces colitis in rats via modulation of the immune system and correction of cytokine imbalance.
J. Gastroenterol. Hepatol.
17
:
1291
1298
.
20
Smith
E.
,
Stark
M. A.
,
Zarbock
A.
,
Burcin
T. L.
,
Bruce
A. C.
,
Vaswani
D.
,
Foley
P.
,
Ley
K.
.
2008
.
IL-17A inhibits the expansion of IL-17A-producing T cells in mice through “short-loop” inhibition via IL-17 receptor.
J. Immunol.
181
:
1357
1364
.
21
Wu
S.
,
Rhee
K. J.
,
Albesiano
E.
,
Rabizadeh
S.
,
Wu
X.
,
Yen
H. R.
,
Huso
D. L.
,
Brancati
F. L.
,
Wick
E.
,
McAllister
F.
, et al
.
2009
.
A human colonic commensal promotes colon tumorigenesis via activation of T helper type 17 T cell responses.
Nat. Med.
15
:
1016
1022
.
22
Ley
K.
,
Smith
E.
,
Stark
M. A.
.
2006
.
IL-17A-producing neutrophil-regulatory Tn lymphocytes.
Immunol. Res.
34
:
229
242
.
23
Theilgaard-Mönch
K.
,
Jacobsen
L. C.
,
Borup
R.
,
Rasmussen
T.
,
Bjerregaard
M. D.
,
Nielsen
F. C.
,
Cowland
J. B.
,
Borregaard
N.
.
2005
.
The transcriptional program of terminal granulocytic differentiation.
Blood
105
:
1785
1796
.
24
Theilgaard-Mönch
K.
,
Porse
B. T.
,
Borregaard
N.
.
2006
.
Systems biology of neutrophil differentiation and immune response.
Curr. Opin. Immunol.
18
:
54
60
.
25
Botha
A. J.
,
Moore
F. A.
,
Moore
E. E.
,
Fontes
B.
,
Banerjee
A.
,
Peterson
V. M.
.
1995
.
Postinjury neutrophil priming and activation states: therapeutic challenges.
Shock
3
:
157
166
.
26
Brown
K. A.
,
Brain
S. D.
,
Pearson
J. D.
,
Edgeworth
J. D.
,
Lewis
S. M.
,
Treacher
D. F.
.
2006
.
Neutrophils in development of multiple organ failure in sepsis.
Lancet
368
:
157
169
.
27
Nieman
G.
,
Searles
B.
,
Carney
D.
,
McCann
U.
,
Schiller
H.
,
Lutz
C.
,
Finck
C.
,
Gatto
L. A.
,
Hodell
M.
,
Picone
A.
.
1999
.
Systemic inflammation induced by cardiopulmonary bypass: a review of pathogenesis and treatment.
J. Extra Corpor. Technol.
31
:
202
210
.
28
Looney
M. R.
,
Gilliss
B. M.
,
Matthay
M. A.
.
2010
.
Pathophysiology of transfusion-related acute lung injury.
Curr. Opin. Hematol.
17
:
418
423
.
29
Juffermans
N. P.
2010
.
Transfusion-related acute lung injury: emerging importance of host factors and implications for management.
Expert Rev. Hematol.
3
:
459
467
.
30
Vlaar
A. P.
,
Schultz
M. J.
,
Juffermans
N. P.
.
2009
.
Transfusion-related acute lung injury: a change of perspective.
Neth. J. Med.
67
:
320
326
.
31
Pillay
J.
,
Hietbrink
F.
,
Koenderman
L.
,
Leenen
L. P.
.
2007
.
The systemic inflammatory response induced by trauma is reflected by multiple phenotypes of blood neutrophils.
Injury
38
:
1365
1372
.
32
Ramaiah
S. K.
,
Jaeschke
H.
.
2007
.
Role of neutrophils in the pathogenesis of acute inflammatory liver injury.
Toxicol. Pathol.
35
:
757
766
.
33
Biffl
W. L.
,
Moore
E. E.
,
Moore
F. A.
.
1995
.
Gut-derived mediators of multiple organ failure: platelet-activating factor and interleukin-6.
Br. J. Hosp. Med.
54
:
134
138
.
34
Gay
J. C.
1993
.
Mechanism and regulation of neutrophil priming by platelet-activating factor.
J. Cell. Physiol.
156
:
189
197
.
35
Suzuki
K.
,
Yamada
M.
,
Kurakake
S.
,
Okamura
N.
,
Yamaya
K.
,
Liu
Q.
,
Kudoh
S.
,
Kowatari
K.
,
Nakaji
S.
,
Sugawara
K.
.
2000
.
Circulating cytokines and hormones with immunosuppressive but neutrophil-priming potentials rise after endurance exercise in humans.
Eur. J. Appl. Physiol.
81
:
281
287
.
36
Mitchell
G. B.
,
Albright
B. N.
,
Caswell
J. L.
.
2003
.
Effect of interleukin-8 and granulocyte colony-stimulating factor on priming and activation of bovine neutrophils.
Infect. Immun.
71
:
1643
1649
.
37
Dang
P. M.
,
Dewas
C.
,
Gaudry
M.
,
Fay
M.
,
Pedruzzi
E.
,
Gougerot-Pocidalo
M. A.
,
El Benna
J.
.
1999
.
Priming of human neutrophil respiratory burst by granulocyte/macrophage colony-stimulating factor (GM-CSF) involves partial phosphorylation of p47(phox).
J. Biol. Chem.
274
:
20704
20708
.
38
Yagisawa
M.
,
Yuo
A.
,
Kitagawa
S.
,
Yazaki
Y.
,
Togawa
A.
,
Takaku
F.
.
1995
.
Stimulation and priming of human neutrophils by IL-1 alpha and IL-1 beta: complete inhibition by IL-1 receptor antagonist and no interaction with other cytokines.
Exp. Hematol.
23
:
603
608
.
39
Biffl
W. L.
,
Moore
E. E.
,
Moore
F. A.
,
Carl
V. S.
,
Kim
F. J.
,
Franciose
R. J.
.
1994
.
Interleukin-6 potentiates neutrophil priming with platelet-activating factor.
Arch. Surg.
129
:
1131
1136
.
40
Yamashiro
S.
,
Kamohara
H.
,
Wang
J. M.
,
Yang
D.
,
Gong
W. H.
,
Yoshimura
T.
.
2001
.
Phenotypic and functional change of cytokine-activated neutrophils: inflammatory neutrophils are heterogeneous and enhance adaptive immune responses.
J. Leukoc. Biol.
69
:
698
704
.
41
Torchinsky
M. B.
,
Blander
J. M.
.
2010
.
T helper 17 cells: discovery, function, and physiological trigger.
Cell. Mol. Life Sci.
67
:
1407
1421
.
42
Costa
V. S.
,
Mattana
T. C.
,
da Silva
M. E.
.
2010
.
Unregulated IL-23/IL-17 immune response in autoimmune diseases.
Diabetes Res. Clin. Pract.
88
:
222
226
.
43
Schwarzenberger
P.
,
Huang
W.
,
Ye
P.
,
Oliver
P.
,
Manuel
M.
,
Zhang
Z.
,
Bagby
G.
,
Nelson
S.
,
Kolls
J. K.
.
2000
.
Requirement of endogenous stem cell factor and granulocyte-colony-stimulating factor for IL-17-mediated granulopoiesis.
J. Immunol.
164
:
4783
4789
.
44
Schwarzenberger
P.
,
La Russa
V.
,
Miller
A.
,
Ye
P.
,
Huang
W.
,
Zieske
A.
,
Nelson
S.
,
Bagby
G. J.
,
Stoltz
D.
,
Mynatt
R. L.
, et al
.
1998
.
IL-17 stimulates granulopoiesis in mice: use of an alternate, novel gene therapy-derived method for in vivo evaluation of cytokines.
J. Immunol.
161
:
6383
6389
.
45
Forlow
S. B.
,
Schurr
J. R.
,
Kolls
J. K.
,
Bagby
G. J.
,
Schwarzenberger
P. O.
,
Ley
K.
.
2001
.
Increased granulopoiesis through interleukin-17 and granulocyte colony-stimulating factor in leukocyte adhesion molecule-deficient mice.
Blood
98
:
3309
3314
.
46
Stark
M. A.
,
Huo
Y.
,
Burcin
T. L.
,
Morris
M. A.
,
Olson
T. S.
,
Ley
K.
.
2005
.
Phagocytosis of apoptotic neutrophils regulates granulopoiesis via IL-23 and IL-17.
Immunity
22
:
285
294
.

The authors have no financial conflicts of interest.