1. Introduction
Blast exposure-induced injury is the most commonly encountered wounds in modern warfare. Traditionally associated with the battlefield environment, blast injuries are being increasingly observed among noncombatants because of increasing terrorist incidents, as well as gas and underground explosion events [ 1 ]. Blast injury is characterized by complex injuries, high shock rate, and high mortality and accounts for over 75% of all combat casualties in the United States forces [ 2 ]. An epidemiological analysis of the injuries in Operation Iraqi Freedom/Operation Enduring Freedom demonstrated that 81% of all injuries were associated with explosions [ 3 ]. A recent study showed that bomb blasts accounted for 82% of all injuries caused by terrorists and this number continues to rise [ 4 ]. The ears, lungs, and gastrointestinal tract are the most susceptible organs to blast injury, and lung injury is a major cause of high mortality. Sufficient energy exposure causes disruption of the capillary-alveolar interface, which leads to parenchymal hemorrhage and destruction of the alveolar walls. Interstitial changes in blast lungs gradually develop into acute respiratory distress syndrome and seriously affect the quality of life or prognosis. Therefore, it is of great importance to investigate the mechanism of blast exposure-induced lung injury for the treatment of wounded individuals.
Inflammatory responses are thought to play an important role in the development of blast exposure-induced lung injury, because increased lung leukocyte infiltration and elevated systemic and proinflammatory cytokine levels in the lungs are frequently associated with blast injury of the lung; furthermore, several factors including tumor necrosis factor- α , interleukin-6, and interleukin-1 β accelerate lung tissue fibrosis and respiratory dysfunction [ 5 ]. In addition to inflammation, blast exposure also induced oxidative stress [ 6 , 7 ] and apoptosis [ 8 ] in the lungs of experimental animals. Recent studies reported that T cell activation had a pivotal role in the development of inflammation [ 9 , 10 ]. At least two signals are required for the full activation of T cells. Signal one requires the engagement of a T cell receptor by antigen-major histocompatibility complex proteins present on the surface of antigen-presenting cells [ 11 ], whereas signal two requires the engagement of CD28, a potent T cell costimulator expressed on antigen-presenting cells [ 12 ]. A blockade of CD28 signaling with antibodies or genetic ablation of CD28 in mice attenuated T cell activation. Barnett-Vanes et al. [ 13 ] found that blast exposure induced pulmonary barotrauma and inflammation, which were associated with increases of interleukin-6 (IL-6) and tumor necrosis factor- α (TNF- α ). Bricker-Anthony et al. [ 14 ] demonstrated that blast exposure caused corneal edema and neovascularization, as well as immune cell infiltrate throughout the eyes. Struebing et al. [ 15 ] showed that ocular blast injury was related to the activation of the innate and acquired immune systems and infiltration of lymphocyte cells. Although it was demonstrated that CD28 signaling is associated with the expressions of inflammatory mediators, apoptosis-related proteins, immunosuppression, and tumorigenesis [ 16 – 19 ], the effects of CD28 deficiency on blast exposure-induced lung injury and activated T cell accumulation have not been investigated.
To demonstrate the mechanism of CD28 deficiency in blast exposure-induced lung injury, we tested the hypothesis that blast exposure-induced lung inflammation, oxidative stress, and apoptosis are attenuated in CD28 knockout (CD28 −/− ) mice. Our results suggest that CD28 deficiency significantly ameliorates blast exposure-induced lung inflammation, oxidative stress, and apoptosis and that CD28 deficiency is also associated with a reduction of T cell accumulation in blast exposure-induced lungs, which might be associated with the PI3K/Akt/FoxO1 signaling pathway. These findings suggest that CD28 deficiency is a promising therapeutic approach to mitigate blast exposure-induced lung injury.
2. Materials and Methods
2.1. Animals and Experimental Protocols
Thirty male CD28 knockout (CD28 −/− ) mice were obtained from the Jackson Laboratory, and thirty male C57BL/6 wild-type (WT) mice were provided by the Experimental Animal Department of the General Hospital of Northern Theater Command. All mice were kept in a room, maintained at a temperature of 20 ± 2°C and humidity of 55%–65%, and allowed free access to food and water in their cages. After acclimation, C57BL/6 WT and CD28 −/− mice were randomly divided into control or/and blast injury groups (model group). The control group was subdivided into two subgroups: C57BL/6 WT and CD28 −/− control groups. The model group was randomly divided into four groups: C57BL/6 24 h and 48 h and CD28 −/− 24 h and 48 h groups. Animal welfare and experimental design were approved by the Ethics Committee of the General Hospital of Northern Theater Command.
2.2. Establishment of Blast Injury Models
A model that mimics blast exposure-induced acute lung injury was used as previously described [ 20 ]. Briefly, mice were anesthetized by the abdominal injection of 2% pentobarbital sodium (1.5 ml/kg). The anesthetized mice were placed in a protective cover with their chest exposed, and aluminum foil was placed in the middle layer. After fixing the screws, the mouse was placed on the wire mesh atop the device. A pressure pump was used to increase the air pressure in the lower part until the aluminum foil burst. The compressed air rapidly expanded from the blasting port at high speed, forming shock waves that impacted the chest of the mouse. The pressure detected by a pressure sensor was transmitted through a data cable and recorded by a computer. Pressure waveform was obtained by the following formula: pressure (PSI) = voltage value∗1000/50.08. The instantaneous shock wave overpressure was 321 ± 24 PSI in this experiment.
2.3. Sample Collection and Processing
At 24 h and 48 h postblast, lung tissue samples were collected. Briefly, after 12 hours of fasting and 4 hours of water deprivation preoperatively, mice were intraperitoneally anesthetized with 2% sodium pentobarbital (1.5 ml/kg) and fixed in a prone position on the operating table. The abdominal cavity was opened, and blood was harvested through the abdominal aorta. The left lung was immersed in 10% formalin buffer for histological analysis, and the upper lobe of the right lung was dried with filter paper and weighed on an analytical balance to obtain the wet weight. After drying in an oven at 60°C for 72 hours, the dry weight of the right lung was recorded. The calculated ratio of dry weight to wet weight was an indicator of edema formation. The remaining lung was placed in a nitrogen canister for protein determination.
2.4. Histological Analysis
The left lung was fixed in 10% formaldehyde at room temperature and embedded in paraffin blocks using a Leica Microsystem tissue processor (ASP 300S, Germany) for histological analysis. For histological staining, sections of 3 μ m thickness were sliced using a Leica Microsystem microtome (Model RM 2265, Germany) and stained with hematoxylin and eosin (H&E).
2.5. Immunohistochemistry Staining
Sections were treated with 1% H 2 O 2 to block endogenous peroxidase and then heated to 97°C in antigen retrieval solution for 15 min. The sample was incubated at room temperature for 60 min with mouse anti-rabbit CD44 (1 : 400, ab157107, Abcam, UK) and anti-rabbit CD163 antibodies (1 : 500, ab182422, Abcam, UK) and rinsed with phosphate buffer solution (PBS). After incubation with primary antibodies, the sections were washed with PBS and incubated with a secondary antibody (Sigma, USA), and then immunohistochemistry visualization was completed using 3,3-diaminobenzidine (DAB) (Sigma, USA) and counterstained with hematoxylin solution. A negative control consisted of the same immunohistochemical method with the omission of the primary antibodies. Cells with yellow and brown particles in their cytoplasm and/or nuclei were identified as positive cells.
2.6. Reactive Oxygen Species (ROS) Detection
Frozen sections of the right lobe of the lungs were immersed in PBS twice for 5 minutes each time, incubated with 10 μ M dihydroethidium in the dark for 20 minutes, immersed again in PBS twice for 5 minutes each time, and observed and photographed by a fluorescence microscope.
2.7. TUNEL Staining
Lung tissue apoptosis was detected in accordance with the instructions of the TUNEL kit (Cloud-Clone Company, USA). The nuclei of apoptotic cells were stained brown. Five different fields were randomly selected under a high-power microscope (×400) to calculate the apoptotic index.
2.8. Immunofluorescence Staining
Immunofluorescence staining was performed to localize p-PI3K (1 : 200, #4228S, Cell Signaling Technology, USA), p-Akt (1 : 400, #4051S, Cell Signaling Technology, USA), and p-FoxO1 (1 : 250, ab131339, Abcam, UK). Lung tissues were dewaxed with xylene, hydrated with a graded alcohol series, incubated with 0.1% Triton X-100 for 30 minutes, and washed three times with PBS for 5 minutes each time. Samples were blocked with 5% bovine serum albumin and 10% goat serum, each for 30 minutes, incubated with a primary antibody overnight at 4°C in a wet box, and stained with a fluorescent secondary antibody. Finally, samples were observed and photographed with a microscope.
2.9. Western Blotting
Western blotting was performed as described previously [ 21 ]. Whole protein was extracted from the lung tissues using a tissue protein extraction kit (Cat No: FD0889 (year 2016), Hangzhou Fude Biological Technology Company, China), and the protein concentration was determined using a BCA protein assay kit (Cat No: FD2001 (year 2017), Hangzhou Fude Biological Technology Company, China). Equal amounts of soluble protein were separated on 10% polyacrylamide gels, transferred onto a nitrocellulose membrane, and followed by routine western blot analysis. The primary antibody is composed of the following: TNF- α (1 : 2000, ab8348, Abcam, UK), IL-10 (1 : 2000, ab9969, Abcam, UK), IL-6 (1 : 1000, ab83053, Abcam, UK), IRE α (1 : 2000, #3294, Abcam, UK), and MDA5 (1 : 200, ab69983, Abcam, UK). IL-1 β (1 : 2000, sc-7884, Santa Cruz Biotechnology Inc., USA), IL-4 (1 : 1000, sc-73318, Santa Cruz Biotechnology Inc., USA), SOD-1 (1 : 1000, sc-11407, Santa Cruz Biotechnology Inc., USA), Bax (1 : 2000, sc-526, Santa Cruz Biotechnology Inc., USA), Caspase-3 (1 : 500, sc-7148, Santa Cruz Biotechnology Inc., USA), Bcl-2 (1 : 1000, sc-7382, Santa Cruz Biotechnology Inc., USA), active Caspase-8 (1 : 100, sc-5263, Santa Cruz Biotechnology Inc., USA), PI3K (1 : 1000, ab186612, Abcam, UK), p-PI3K (1 : 1000, #4228S, Cell Signaling Technology, USA), Act (1 : 2000, ab32505, Abcam, USA), p-Act (1 : 2000, #4051S, Cell Signaling Technology, USA), Forkhead box O1 (FoxO1) (1 : 1000, #2880, Cell Signaling Technology, USA), p-FoxO1 (1 : 1000, ab131339, Abcam, UK), and GAPDH (1 : 5000, sc-32233, Santa Cruz Biotechnology Inc., USA) were obtained. The secondary antibody was goat anti-mouse secondary antibody (HRP) (1 : 4000 mouse IgG, ab6789, Abcam, UK), goat anti-rabbit secondary antibody (HRP) (1 : 4000, ab6721, Abcam, UK), and goat anti-rat secondary antibody (HRP) (1 : 2000, ab7097, Abcam, UK). Proteins were visualized using a Clarity™ Western ECL Substrate (#170-5061; Bio-Rad Laboratories Inc., USA) and a Tanon 5200 Full automatic chemiluminescence image analysis system (Tanon Science and Technology Co. Ltd., Shanghai, China).
2.10. Flow Cytometry
Flow cytometric analysis was performed as described previously [ 22 ]. Briefly, individual lungs were excised, cut into small pieces, and enzymatically digested in 5 ml of digestion buffer (HBSS without Ca+/Mg+ (Life Technologies) and 1 mg/ml collagenase (Roche)) at 37°C for 30 min with agitation and then pressed against the bottom of a 100 μ m strainer with the plunger of a 3 ml syringe. Single cells from tissues were washed through the strainer with 10 ml cold buffer (PBS + 0.5%BSA + 2 mM EDTA). After erythrocyte lysis using Red Blood Cell Lysing Buffer (Sigma), cells were counted using a hemocytometer. Single cell suspensions were preincubated with anti-mouse CD3 or CD28 antibodies (BD Biosciences, Franklin Lakes, NJ, USA) to prevent nonspecific binding of antibodies to FcR γ , followed by multistaining with fluorescence conjugated primary antibodies. Dead cells were stained with propidium iodide staining solution. Samples were subjected to FACS Aria II analysis (BD Biosciences). Data were analyzed by FlowJo software.
2.11. Statistical Analysis
Data are expressed as the mean ± standard deviation and were analyzed using SPSS 20.0 statistical software. All experiments were repeated at least 3 times. Significance was determined when P values < 0.05 were obtained by two-way ANOVA with the Bonferroni post hoc test.
3. Results
3.1. CD28 Deficiency Attenuates Blast Exposure-Induced Acute Lung Injury
Compared with the control group, the level of CD28 protein was significantly elevated in the model group, whereas CD28 deficiency reduced blast exposure-induced levels of CD28 compared with the C57BL/6 model group (Figures 1(a) and 1(b) , P < 0.05). Blast exposure significantly increased the ratio of lung weight/body weight and the ratio of wet weight/dry weight compared with the control group, whereas CD28 deficiency significantly inhibited blast exposure-induced increases of the ratios compared with the C57BL/6 model group (Figures 1(c) and 1(d) , P < 0.05). In addition, serious lung hemorrhage, edema, and obvious inflammatory cell infiltration were observed in the C57BL/6 model group, whereas these were attenuated in the CD28 −/− model group ( Figure 1(e) ). These data demonstrate that CD28 deficiency was effective at ameliorating blast exposure-induced acute lung injury.
3.2. CD28 Deficiency Inhibits Blast Exposure-Induced Lung Proinflammatory Cytokine Expression
Both CD28 deficiency and blast affected the expression of IL-1 β , IL-6, TNF- α , NF- κ B, CD44, CD163, and IL-10. At 24 h and 48 h postblast, higher expressions of IL-1 β , IL-6, TNF- α , NF- κ B, CD44, and CD163 and a lower expression of IL-10 were observed in the control group compared with the CD28 −/− model group ( Figure 2(a) – 2(h) , P < 0.05). Moreover, immunohistochemical data also demonstrated that the expression levels of CD44- and CD163-positive inflammatory cells in the CD28 −/− model group were significantly lower than those in the C57BL/6 model group ( Figure 2(i) – 2(l) , P < 0.05). These data demonstrate that CD28 deficiency was effective at ameliorating blast exposure-induced lung proinflammatory cytokine expression.
3.3. CD28 Deficiency Inhibits Blast Exposure-Induced Expression of ROS and Oxidative Stress
Compared with the control group, the expression of ROS was significantly higher in the model group, while CD28 deficiency significantly decreased blast-induced ROS expression compared with the C57BL/6 model group ( Figure 3(a) ). Western blot results showed that blast exposure increased the levels of MDA5 and IRE α proteins and decreased SOD-1 protein compared with the control group, whereas blast-induced increases of MDA5 and IRE α protein and decreases of SOD-1 protein were significantly attenuated by CD28 deficiency compared with the C57BL/6 model group ( Figure 3(b) – 3(e) , P < 0.05). This demonstrated that CD28 deficiency is effective at ameliorating blast exposure-induced lung reactive oxygen species and oxidative stress.
3.4. CD28 Deficiency Ameliorates Blast Exposure-Induced Lung Tissue Apoptosis
TUNEL results showed that the number of apoptotic cells in the model group was much higher than that in the control group and that the number of apoptotic cells induced by blast injury was significantly decreased by CD28 deficiency compared with the C57BL/6 model group (Figures 4(a) and 4(b) , P < 0.05). These results demonstrated that blast exposure increased the levels of the proapoptotic proteins Bax, Caspase-3, and active Caspase-8 and decreased the levels of the antiapoptotic proteins Bcl-2 compared with the control group, whereas CD28 deficiency evidently attenuated blast exposure-induced increases of Bax, Caspase-3, and active Caspase-8 and decreased Bcl-2 compared with the C57BL/6 model group ( Figure 4(b) – 4(g) , P < 0.05). These data suggest that CD28 deficiency is effective at ameliorating blast exposure-induced lung tissue apoptosis.
3.5. CD28 Deficiency Inhibits CD3 + T Cell Accumulation in Blast Exposure-Induced Lung Tissues
At 24 h and 48 h postblast, the percentage of total CD3 + T cells in lungs was remarkably increased compared with the control group, while it was significantly ameliorated by CD28 deficiency, which decreased the accumulation of total CD3 + T cells and inhibited early activated T cells compared to the C57BL/6 model group ( Figure 5 , P < 0.05). These results show that CD28 deficiency is effective in ameliorating blast exposure-induced CD3 + T cell accumulation in lungs.
3.6. CD28 Deficiency Attenuates Blast Exposure-Induced Lung Injury through the PI3K/Akt/FoxO1 Signaling Pathway
The expression levels of p-PI3K, p-Akt, and p-FoxO1 proteins were not altered in CD28 −/− mice under control conditions. At 24 h and 48 h postblast, western blot results showed that the expressions of p-PI3K and p-Akt were significantly increased and p-FoxO1 levels were significantly decreased compared with the control group, whereas these factors were significantly attenuated by CD28 deficiency as compared with the C57BL/6 model group ( Figure 6(a) – 6(d) , P < 0.05). Moreover, immunofluorescence results also confirmed that blast exposure induced the higher expressions of p-PI3K and p-Akt and lower expression of p-FoxO1 in the CD28 −/− model group compared with the C57BL/6 model group ( Figure 6(e) – 6(j) ). These data demonstrate that CD28 deficiency is effective at ameliorating blast exposure-induced lung injury via the PI3K/Akt/FoxO1 signaling pathway.
4. Discussion
Blast exposure induces injury in several organs including the lungs [ 23 ], brain [ 24 , 25 ], heart [ 26 ], ears [ 27 ], jejunum [ 28 ], and kidney [ 29 ]. A number of studies demonstrated that blast exposure caused inflammation, oxidative stress, and cell apoptosis in the lungs of experimental animals and humans [ 20 , 30 , 31 ]. In this study, we showed that CD28 deficiency ameliorated blast exposure-induced acute lung injury, reduced proinflammatory cytokine expression, decreased oxidative stress and cell apoptosis, and also inhibited CD3 + T cell accumulation in lung tissues. Additionally, CD28 deficiency significantly ameliorated blast exposure-induced increases in p-PI3K and p-Akt expressions and ameliorated decreases in p-FoxO1 expression. Therefore, we propose that the attenuation of blast exposure-induced lung injury by CD28 deficiency might be associated with the activation of the PI3K/Akt/FoxO1 signaling pathway.
In this study, our results demonstrated that CD28 deficiency ameliorated blast exposure-induced acute lung injury and inhibited proinflammatory cytokine expression. Several studies demonstrated that CD28-mediated signaling prevented the spontaneous development of autoimmune diabetes [ 32 ] and decreased experimental autoimmune neuritis [ 33 ] and CD28 deficiency was highly resistant to collagen-induced arthritis in mice [ 34 ]. Similar to our results, Macal and Tam [ 35 ] reported that the levels of the proinflammatory cytokines TNF- α and IL-6 were significantly decreased in CD28-deficient pDCs compared with WT pDCs. Moreover, in vivo CD28 knockdown decreased the levels of COX-2, IL-10, and IL-17A mRNAs at day 24 in a chronic UVB exposure study [ 16 ]. Whereas the CD28-B7 blockade by CTLA4-Ig inhibited IFN- γ production in C57BL/6 recipients, it had little effect on the production of IL-6 [ 36 ]. Similarly, the administration of a CD28 antagonist peptide decreased irradiation-induced increase in IL-6 and lowered COX-2 expression and the numbers of macrophages in the small intestine of irradiated mice [ 37 ]. The blockade of CD28 signals also exacerbated left ventricular remodeling and increased cardiac rupture after myocardial infarction by prolonging the inflammatory period, which caused a reduction in collagen fibers in infarct scars [ 38 ]. These results suggest that CD28 deficiency is effective at ameliorating blast exposure-induced lung inflammation.
In addition to its anti-inflammatory effects, we also showed that CD28 deficiency decreased blast exposure-induced oxidative stress and cell apoptosis. It was reported that deltamethrin induced apoptosis through its interaction with CD28 receptors, leading to oxidative stress and activation of the mitochondrial caspase-dependent pathways, which finally affected the functions of the immune system [ 39 ]. In addition, the CTLA4-Ig blockade of synovial adherent cell signaling to CD28 T cells prevented ROS induction in T lymphocytes exposed to inflammatory cytokines [ 40 ]. Similar results were reported by Ma et al. [ 41 ] where the blockade of the CD80/CD28 signaling pathway promoted recovery of spatial memory, which was associated with mechanisms involving the CD80/CD28 pathway that regulates inflammatory and immune responses and downregulates programmed cell death through apoptosis and autophagy. Our results also showed that CD28 deficiency decreased blast exposure-induced T cell accumulation. Similarly, Suresh et al. [ 42 ] demonstrated that the CD28 signaling pathway regulated the generation and maintenance of the immune responses. Soluble CD28 promoted T cell responses, leading to T cell activation and proliferation [ 43 ]. Zaitsu et al. [ 44 ] reported that the blockade of selective CD28 promoted Treg functions and the regulation of alloimmune responses in vivo . Moreover, Wang et al. [ 22 ] showed that the CD28/B7 blockade by CTLA4-Ig treatment and CD28/B7 KO inhibited activated effector T cell accumulation in transverse aortic constriction-induced heart failure. These studies indicate that CD28 deficiency might have a protective role against blast exposure-induced lung injury by ameliorating oxidative stress, apoptosis, and activated T cell accumulation.
Our results showed that CD28 deficiency decreased blast exposure-induced lung injury, which might be associated with the PI3K/Akt/FoxO1 signaling pathway. It was recently reported that CD28 signaling in T cells is related closely to the phosphatidylinositol 3-kinase (PI3K)/Akt pathway [ 45 , 46 ]. Similar to T cells, CD28 activation in multiple myeloma cells activated PI3K/Akt signaling, which might regulate the apoptotic resistance of multiple myeloma [ 47 , 48 ]. FoxO1 knockdown partly prevented the blockade of CD28 and led to death, whereas the activation of CD28 could increase p-FoxO1 [ 49 ]. A recent study showed that the PI3K/Akt pathway was required for the CD28-induced activation of NF- κ B and the CD28-induced signaling pathway, which lead to significantly increased of T cell proliferation and cytokine production [ 50 ]. Furthermore, blocking CD28 inhibited chemotherapy-induced cell apoptosis, and the CD28 signaling pathway was associated with the downstream activation of PI3K/Akt and inactivation of the FoxO1 in multiple myeloma [ 51 ]. Based on the above results, we speculated on the putative molecular mechanism involved in the CD28 regulation of blast injury ( Figure 7 ). Blast exposure induces the phosphorylation of PI3K protein, which activates Akt proteins in C57BL/6 mice. Once Akt is activated, it induces p-FoxO1, a downstream target protein of Akt, which is then dephosphorylated to form FoxO1 and transferred to the nucleus. Therefore, we hypothesize that FoxO1 upregulates IL-1 β , IL-6, TNF- α , NF- κ B, CD44, active Caspase-8, CD163, MDA5, IRE α , Bax, and Caspase-3 expressions; downregulates the expressions of IL-10, SOD-1, and Bcl-2; and then regulates inflammation, oxidative stress, and apoptosis. CD28 KO may reduce PI3K/Akt phosphorylation and increase FoxO1 phosphorylation, which finally reverses blast exposure-induced lung injury. More experiments are required to further study and clarify the detailed mechanism of this phenomenon.
In summary, blast exposure leads to inflammation, oxidative stress, and apoptosis in mouse lungs. CD28 deficiency has a protective effect on blast exposure-induced lung injury, which might be associated with the PI3K/Akt/FoxO1 signaling pathway.
Acknowledgments
We thank J. Ludovic Croxford, PhD, from Liwen Bianji, Edanz Group China ( http://www.liwenbianji.cn/ac ), for editing the English text of a draft of this manuscript. We would like to acknowledge funding support by the grants of the Liaoning Province Key Scientific and Technological Project (No. 201602786 and No. 20170540947) and the PLA Foundation of China (No. CSY12J002, No. AWS14L008, and No. CSY13J003).
Data Availability
All authors declare that all data are fully available without restriction.
Conflicts of Interest
The authors declare that there are no conflicts of interest.
Authors' Contributions
Mingxiao Hou and Hongxu Jin designed the experiments. Yunen Liu and Yan Zhao wrote the manuscript. Changci Tong, Ying Liu, Lin Shi, and Peifang Cong completed the immunofluorescence, western blot, and quantitative real-time PCR. Xiuyun Shi, Ying Xu, and Guangyuan Bi completed the statistical analysis. Yunen Liu and Changci Tong contributed equally to this work.
1 Covey D. C. Born C. T. Blast injuries: mechanics and wounding patterns Journal of Surgical Orthopaedic Advances 2010 19 8 12 20371000
2 Belmont P. J. Schoenfeld A. J. Goodman G. Epidemiology of combat wounds in Operation Iraqi Freedom and Operation Enduring Freedom: orthopaedic burden of disease Journal of Surgical Orthopaedic Advances 2010 19 1 2 7 20370999
3 Owens B. D. Kragh J. F. Jr. Wenke J. C. Macaitis J. Wade C. E. Holcomb J. B. Combat wounds in operation Iraqi Freedom and operation Enduring Freedom The Journal of Trauma 2008 64 2 295 299 10.1097/TA.0b013e318163b875 2-s2.0-43149122483 18301189
4 Kluger Y. Bomb explosions in acts of terrorism--detonation, wound ballistics, triage and medical concerns Israel Medical Association Journal 2003 5 235 240 14509125
5 Ning J. L. Mo L. W. Lu K. Z. Lai X. N. Wang Z. G. Ma D. Lung injury following lower extremity blast trauma in rats Journal of Trauma and Acute Care Surgery 2012 73 6 1537 1544 10.1097/TA.0b013e318266013a 2-s2.0-84871210662 23064609
6 Elsayed N. M. Armstrong K. L. William M. T. Cooper M. F. Antioxidant loading reduces oxidative stress induced by high-energy impulse noise (blast) exposure Toxicology 2000 155 1-3 91 99 10.1016/S0300-483X(00)00281-X 2-s2.0-0034735890 11154801
7 Ning J. Mo L. Yi B. Therapeutic whole-body hypothermia protects remote lung, liver, and kidney injuries after blast limb trauma in rats Anesthesiology 2016 124 6 1360 1371 10.1097/ALN.0000000000001106 2-s2.0-84961943229 27028466
8 Liener U. C. Knöferl M. W. Sträter J. Induction of apoptosis following blunt chest trauma Shock 2003 20 6 511 516 10.1097/01.shk.0000095057.62263.fb 2-s2.0-3042825657 14625474
9 Niemela V. Burman J. Blennow K. Zetterberg H. Larsson A. Sundblom J. Cerebrospinal fluid sCD27 levels indicate active T cell-mediated inflammation in premanifest Huntington’s disease PLoS One 2018 13 2, article e0193492 10.1371/journal.pone.0193492 2-s2.0-85042508910 29474427 PMC5825143
10 Wang D. Gu X. Liu X. Wei S. Wang B. Fang M. NK cells inhibit anti- Mycobacterium bovis BCG T cell responses and aggravate pulmonary inflammation in a direct lung infection mouse model Cellular Microbiology 2018 20 7 10.1111/cmi.12833 2-s2.0-85043310427 29447423
11 Fontana M. F. Vance R. E. Two signal models in innate immunity Immunological Reviews 2011 243 1 26 39 10.1111/j.1600-065X.2011.01037.x 2-s2.0-80052153210 21884165
12 Kuo C. L. Hood L. Antigen/major histocompatibility complex-specific activation of murine T cells transfected with functionally rearranged T-cell receptor genes Proceedings of the National Academy of Sciences of the United States of America 1987 84 21 7614 7618 10.1073/pnas.84.21.7614 2-s2.0-0023605098 2823268 PMC299350
13 Barnett-Vanes A. Sharrock A. Eftaxiopoulou T. CD43Lo classical monocytes participate in the cellular immune response to isolated primary blast lung injury Journal of Trauma and Acute Care Surgery 2016 81 3 500 511 10.1097/TA.0000000000001116 2-s2.0-84974817730 27306447
14 Bricker-Anthony C. Hines-Beard J. D'Surney L. Rex T. S. Exacerbation of blast-induced ocular trauma by an immune response Journal of Neuroinflammation 2014 11 1 p. 192 10.1186/s12974-014-0192-5 2-s2.0-84986183436 25472427 PMC4264554
15 Struebing F. L. King R. Li Y. Transcriptional changes in the mouse retina after ocular blast injury: a role for the immune system Journal of Neurotrauma 2018 35 1 118 129 10.1089/neu.2017.5104 2-s2.0-85040466330 28599600 PMC5757081
16 Singh T. P. Mayer G. Wolf P. In vivo siRNA targeting of CD28 reduces UV-induced DNA damage and inflammation The Journal of Investigative Dermatology 2014 134 3 861 864 10.1038/jid.2013.397 2-s2.0-84894101262 24042341
17 Escarra-Senmarti M. Bueno-Topete M. R. Jave-Suarez L. F. Loss of CD28 within CD4 + T cell subsets from cervical cancer patients is accompanied by the acquisition of intracellular perforin, and is further enhanced by NKG2D expression Immunology Letters 2017 182 30 38 10.1016/j.imlet.2017.01.006 2-s2.0-85009423939 28087292
18 Oda S. K. Daman A. W. Garcia N. M. A CD200R-CD28 fusion protein appropriates an inhibitory signal to enhance T-cell function and therapy of murine leukemia Blood 2017 130 22 2410 2419 10.1182/blood-2017-04-777052 2-s2.0-85036555262 29042364 PMC5709784
19 Zhou Y. Salker M. S. Walker B. Acid sphingomyelinase (ASM) is a negative regulator of regulatory T cell (Treg) development Cellular Physiology and Biochemistry 2016 39 3 985 995 10.1159/000447806 2-s2.0-84986888941 27512981
20 Liu Y. E. Tong C. C. Zhang Y. B. Chitosan oligosaccharide ameliorates acute lung injury induced by blast injury through the DDAH1/ADMA pathway PLoS One 2018 13 2, article e0192135 10.1371/journal.pone.0192135 2-s2.0-85041494161 PMC5802901 29415054
21 Ansari M. Y. Ahmad N. Haqqi T. M. Butein activates autophagy through AMPK/TSC2/ULK1/mTOR pathway to inhibit IL-6 expression in IL-1 β stimulated human chondrocytes Cellular Physiology and Biochemistry 2018 49 3 932 946 10.1159/000493225 2-s2.0-85053034932 30184535
22 Wang H. Kwak D. Fassett J. CD28/B7 deficiency attenuates systolic overload-induced congestive heart failure, myocardial and pulmonary inflammation, and activated T cell accumulation in the heart and lungs Hypertension 2016 68 3 688 696 10.1161/HYPERTENSIONAHA.116.07579 2-s2.0-84978698576 27432861 PMC5314944
23 Scott T. E. Kirkman E. Haque M. Gibb I. E. Mahoney P. Hardman J. G. Primary blast lung injury - a review British Journal of Anaesthesia 2017 118 3 311 316 10.1093/bja/aew385 2-s2.0-85016052741 28203741
24 Callahan M. L. Storzbach D. Sensory sensitivity and posttraumatic stress disorder in blast exposed veterans with mild traumatic brain injury Applied Neuropsychology: Adult 2019 26 4 365 373 10.1080/23279095.2018.1433179 2-s2.0-85042230347 29465307
25 Yamamoto S. DeWitt D. S. Prough D. S. Impact & blast traumatic brain injury: implications for therapy Molecules 2018 23 2 p. 245 10.3390/molecules23020245 2-s2.0-85041221489 PMC6017013 29373501
26 Han G. Wang Z. Wang J. Characteristics and mechanisms of cardiopulmonary injury caused by mine blasts in shoals: a randomized controlled study in a rabbit model PLoS One 2013 8 12, article e81310 10.1371/journal.pone.0081310 2-s2.0-84892934693 24358110 PMC3864783
27 Ballivet de Regloix S. Crambert A. Maurin O. Lisan Q. Marty S. Pons Y. Blast injury of the ear by massive explosion: a review of 41 cases Journal of the Royal Army Medical Corps 2017 163 5 333 338 10.1136/jramc-2016-000733 2-s2.0-85030763070 28209807
28 Vellar D. J. Pucius R. Vellar I. D. Explosion injury of the proximal jejunum caused by diathermy in a patient with obstructing sclerosing peritonitis The British Journal of Surgery 1986 73 2 157 158 10.1002/bjs.1800730229 2-s2.0-0022618947 3947910
29 Wong C. Ogan K. Pareek G. Controversial case in endourology. A 27-year-old man suffered a blast injury while serving in Iraq. A right perinephric hematoma and shrapnel near the hilum of the right kidney were noted on CT Journal of Endourology 2009 23 8 1223 1225 10.1089/end.2009.0331 2-s2.0-68949084844 19653870
30 Chavko M. Adeeb S. Ahlers S. T. McCarron R. M. Attenuation of pulmonary inflammation after exposure to blast overpressure by N-acetylcysteine amide Shock 2009 32 3 325 331 10.1097/SHK.0b013e31819c38f1 2-s2.0-69549088528 19174737
31 Gorbunov N. V. Elsayed N. M. Kisin E. R. Kozlov A. V. Kagan V. E. Air blast-induced pulmonary oxidative stress: interplay among hemoglobin, antioxidants, and lipid peroxidation American Journal of Physiology-Lung Cellular and Molecular Physiology 1997 272 2 L320 L334 10.1152/ajplung.1997.272.2.l320 9124384
32 Salomon B. Lenschow D. J. Rhee L. B7/CD28 costimulation is essential for the homeostasis of the CD4 + CD25 + immunoregulatory T cells that control autoimmune diabetes Immunity 2000 12 4 431 440 10.1016/S1074-7613(00)80195-8 2-s2.0-0033682056 10795741
33 Schmidt J. Elflein K. Stienekemeier M. Treatment and prevention of experimental autoimmune neuritis with superagonistic CD28-specific monoclonal antibodies Journal of Neuroimmunology 2003 140 1-2 143 152 10.1016/S0165-5728(03)00182-6 2-s2.0-0038643708 12864982
34 Tada Y. Nagasawa K. Ho A. CD28-deficient mice are highly resistant to collagen-induced arthritis The Journal of Immunology 1999 162 203 208 9886387
35 Macal M. Tam M. A. Hesser C. CD28 deficiency enhances type I IFN production by murine plasmacytoid dendritic cells The Journal of Immunology 2016 196 4 1900 1909 10.4049/jimmunol.1501658 2-s2.0-84958576534 26773151 PMC4744517
36 Zhao X. Boenisch O. Yeung M. Critical role of proinflammatory cytokine IL-6 in allograft rejection and tolerance American Journal of Transplantation 2012 12 1 90 101 10.1111/j.1600-6143.2011.03770.x 2-s2.0-84855962311 21992708
37 Mirzoeva S. Paunesku T. Wanzer M. B. Single administration of p 2TA (AB103), a CD28 antagonist peptide, prevents inflammatory and thrombotic reactions and protects against gastrointestinal injury in total-body irradiated mice PLoS One 2014 9 7, article e101161 10.1371/journal.pone.0101161 2-s2.0-84904632453 PMC4108308 25054224
38 Kubota A. Hasegawa H. Tadokoro H. Deletion of CD28 co-stimulatory signals exacerbates left ventricular remodeling and increases cardiac rupture after myocardial infarction Circulation Journal 2016 80 9 1971 1979 10.1253/circj.CJ-16-0327 2-s2.0-84983250847 27396441
39 Kumar A. Sasmal D. Bhaskar A. Mukhopadhyay K. Thakur A. Sharma N. Deltamethrin-induced oxidative stress and mitochondrial caspase-dependent signaling pathways in murine splenocytes Environmental Toxicology 2016 31 7 808 819 10.1002/tox.22091 2-s2.0-84920019907 25534813
40 Remans P. H. J. Wijbrandts C. A. Sanders M. E. CTLA-4IG suppresses reactive oxygen species by preventing synovial adherent cell-induced inactivation of Rap1, a Ras family GTPASE mediator of oxidative stress in rheumatoid arthritis T cells Arthritis and Rheumatism 2006 54 10 3135 3143 10.1002/art.22139 2-s2.0-33750343810 17009234
41 Ma L. Shen X. Gao Y. Blocking B7-1/CD28 pathway diminished long-range brain damage by regulating the immune and inflammatory responses in a mouse model of intracerebral hemorrhage Neurochemical Research 2016 41 7 1673 1683 10.1007/s11064-016-1883-3 2-s2.0-84961183907 26980009
42 Suresh M. Whitmire J. K. Harrington L. E. Role of CD28-B7 interactions in generation and maintenance of CD8 T cell memory The Journal of Immunology 2001 167 10 5565 5573 10.4049/jimmunol.167.10.5565 2-s2.0-0035889949 11698427
43 Crepeau R. L. Ford M. L. Challenges and opportunities in targeting the CD28/CTLA-4 pathway in transplantation and autoimmunity Expert Opinion on Biological Therapy 2017 17 8 1001 1012 10.1080/14712598.2017.1333595 2-s2.0-85023621872 28525959 PMC5590720
44 Zaitsu M. Issa F. Hester J. Vanhove B. Wood K. J. Selective blockade of CD28 on human T cells facilitates regulation of alloimmune responses JCI Insight 2017 2 19 10.1172/jci.insight.89381 28978798 PMC5841880
45 Friend L. D. Shah D. D. Deppong C. A dose-dependent requirement for the proline motif of CD28 in cellular and humoral immunity revealed by a targeted knockin mutant The Journal of Experimental Medicine 2006 203 9 2121 2133 10.1084/jem.20052230 2-s2.0-33748468629 16908623 PMC2118406
46 Burr J. S. Savage N. D. L. Messah G. E. Cutting edge: distinct motifs within CD28 regulate T cell proliferation and induction of Bcl-XL The Journal of Immunology 2001 166 9 5331 5335 10.4049/jimmunol.166.9.5331 2-s2.0-0035340150 11313368
47 Bahlis N. J. King A. M. Kolonias D. CD28-mediated regulation of multiple myeloma cell proliferation and survival Blood 2007 109 11 5002 5010 10.1182/blood-2006-03-012542 2-s2.0-34249671217 17311991 PMC1885531
48 Wang G. Sun J. Interactive effects of Snps located within CD28/B7 pathway and environment on susceptibility to recurrent spontaneous abortion Cellular Physiology and Biochemistry 2017 43 6 2185 2199 10.1159/000484297 2-s2.0-85032383511 29069644
49 Lu H. Zhang L. H. Yang L. Tang P. F. The PI3K/Akt/FOXO3a pathway regulates regeneration following spinal cord injury in adult rats through TNF- α and p27kip1 expression International Journal of Molecular Medicine 2018 41 5 2832 2838 10.3892/ijmm.2018.3459 2-s2.0-85042788984 29436581
50 Skanland S. S. Moltu K. Berge T. Aandahl E. M. Tasken K. T-cell co-stimulation through the CD2 and CD28 co-receptors induces distinct signalling responses The Biochemical Journal 2014 460 3 399 410 10.1042/BJ20140040 2-s2.0-84901723757 24665965
51 Murray M. E. Gavile C. M. Nair J. R. CD28-mediated pro-survival signaling induces chemotherapeutic resistance in multiple myeloma Blood 2014 123 24 3770 3779 10.1182/blood-2013-10-530964 2-s2.0-84902602754 24782505 PMC4055924