Abstract
Introduction
Trauma alters the immune response in numerous ways, affecting both the innate and adaptive responses. Macrophages play an important role in inflammation and wound healing following injury. We hypothesize that macrophages mobilize from the circulation to the site of injury and secondary sites after trauma, with a transition from proinflammatory (M1) shortly after trauma to anti-inflammatory (M2) at later time points.
Methods
C57Bl6 mice (n=6/group) underwent a polytrauma model using cardiac puncture/hemorrhage, pseudo-femoral fracture and liver crush injury. The animals were sacrificed at several time points: uninjured, 24 hours and 7 days. Peripheral blood mononuclear cells, spleen, liver non parenchymal cells and lung were harvested, processed and stained for flow cytometry. Macrophages were identified as CD68+; M1 macrophages were identified as iNOS+; M2 macrophages as Arginase1+.
Results
We saw a slight presence of M1 macrophages at baseline in peripheral blood mononuclear cells (6.6%), with no significant change at 24 hours and 7 days after polytrauma. In contrast, the spleen has a larger population of M1 macrophages at baseline (27.7%), with levels decreasing at 24 hours and 7 days after trauma (20.6% and 12.6% respectively). A similar trend is seen in the lung where at baseline, 14.9% of CD68+ macrophages are M1, with subsequent continual decrease reaching 8.7% at 24 hours and 4.4% at 7 days after polytrauma. M1 macrophages in the liver represent 14.3% of CD68+ population in liver non parenchymal cells at baseline. This percentage increases to 20.8% after trauma and decreases at 7 days after polytrauma (13.4%).
There are few M2 macrophages in circulating PBMCs and in spleen at baseline and after trauma. The percentage of M2 macrophages in lungs remains constant after trauma (7.2% at 24 hours and 9.2% at 7 days). In contrast, a large proportion of M2 macrophages are seen in the liver at baseline (36.0%). This percentage trends upwards and reaches 45.6% acutely after trauma and drops to 21.4% at 7 days.
The phenotypic changes in macrophages seen in lungs did not correlate with a functional change in the ability of the macrophages to perform oxidative burst, with an increase from 2.0% at baseline to 22.1% at 7 days after polytrauma (p=0.0258).
Conclusion
Macrophage phenotypic changes after polytrauma are noted, especially with a decrease in the lung M1 phenotype and a short-term increase in the M2 phenotype in the liver. However, macrophage function as measured by oxidative burst increased over the time course of trauma, which may signify a change in subset polarization after injury not captured by the typical macrophage phenotypes.
Keywords: Macrophage polarization, inflammation, polytrauma, M1 macrophages, M2 macrophages
Introduction
Trauma is one of the leading causes of death worldwide1. While death within the first 24 hours of injury is typically related to hemorrhagic shock and the injury itself, in the subsequent days and weeks, the main cause of death after traumatic injury becomes infectious complications, affecting around 10% of trauma patients1. The incidence of sepsis increases with the severity of the trauma and is related to the dysregulation of the immune response caused by injury2. Numerous changes occur to both the innate and adaptive arms of the immune response, leading to the clinical presentation of the systemic inflammatory response syndrome (SIRS) as well as a compensatory anti-inflammatory response syndrome (CARS)1. These changes can lead to the findings that Moore and colleagues described as a two-hit response in survivors of trauma, where the initial insult of traumatic injury causes a maladaptive immune response to a secondary insult such as an infection3. This maladaptive response may present as a dangerous hyperinflammation leading to multiple organ failure or, alternatively, a diminished response to microbial threat leading to overwhelming infection. Better delineation of the specific effects that trauma has on immune cell phenotypes may provide insight into which subgroup of patients will have impaired responses to subsequent infection, and may also provide targets for therapies to avoid infection after trauma.
Macrophages play an important role in innate immunity4. Macrophages exist in a spectrum of phenotypes, and can change their polarization depending on a number of factors such as their microenvironment5. The M1 or “classical” pro-inflammatory phenotype is stimulated by microbial products or pro-inflammatory cytokines such as TNF-α and IFN-γ5. M1 macrophages typically produce nitrous oxide and reactive oxygen intermediates as well as pro-inflammatory cytokines, promote a Th1 response, and have microbicidal and tumoricidal activity5. The M2 anti-inflammatory phenotype is observed in healing tissue and is induced by anti-inflammatory cytokines such as IL-4, −10 and −135. The M2 phenotype is involved in the promotion of a Th2 response, tissue remodeling, immune tolerance and tumor progression5. These phenotypes are malleable, existing on a spectrum, and it is known that macrophages can “re-polarize”6.
In previous work, we found that monocytes harvested from the peripheral blood of human trauma patients had increased ability for innate functions such as oxidative burst, but had decreased ability to produce pro-inflammatory cytokines after stimulation with heat-killed bacteria7. This work led to the hypothesis that macrophage polarization may be affected by traumatic injury, leading to this mixed phenotype. Macrophage polarization has not been widely studied after trauma. The bulk of this literature is focused on traumatic brain injury (TBI), with studies showing the presence of a chronic and persistent M1 polarization lasting months after TBI within the lesion microenvironment8. In this study, we attempted to fill this knowledge gap by comprehensively phenotyping macrophage polarization in blood, spleen, liver, and lung over several time points in a mouse polytrauma model. We then set out to examine the functional changes that macrophages corresponding to these time points by evaluating oxidative burst in end organs. We hypothesized that in a process of inflammation and subsequent wound healing, we would observe an increase in M1 inflammatory macrophages early after trauma with a conversion to an M2 anti-inflammatory phenotype at later time points, and that the functional change would match any phenotypic change seen.
Materials and methods
1. Animals
Wild type adult male mice (C57BL; age: 8–12 weeks; weight: 26–28g) were obtained from Jackson Laboratory. All animals were housed in a controlled environment and provided with standard rodent nutrition and water ad libitum, under a 12-hour light-dark cycle. All surgical procedures were carried out in accordance with protocols approved by the Institutional Animal Care and Use Committee at the University of Pittsburgh Medical Center.
2. Polytrauma model
Mice were weighed and anesthetized with isofluorane (3% on induction and 2% for maintenance). Mice were placed on heated pads with temperature maintained at 37°C. A blind cardiac puncture was performed where 25% of the total blood volume (determined by the weight of the mice) was taken. A bilateral pseudofemoral fracture was then performed during which 0.15cc of ground bone marrow matrix from a matched age donor uninjured mouse was injected around the femur of each leg9. A laparotomy was then performed, and ten liver crush injuries were completed by placing the lobes of the liver between the tips of a forceps. The abdomen was then closed with 4–0 PDS, anesthesia was terminated, and the mice were placed into a cage with access to a heated pad. They were given buprenorphine at a dose of 0.03 mg/mL every 12 hours by subcutaneous injection for pain management post operatively.
3. Harvest of organs
Animals were sacrificed either after no injury (out of box control mice), at 24 hours after polytrauma (PT), or 7 days after PT (n=6 per group). Blood was collected into a citrated tube and peripheral blood mononuclear cells (PBMC) were purified using a Ficoll gradient (Millipore Sigma). Harvest of spleen, liver and lung was also performed. The spleens were macerated, and immune cells were extracted. Lung tissues were subjected to tissue digestion using liberase TL (50 μg/mL final concentration; Millipore Sigma) and DNase I (1 μg/mL final concentration; Millipore Sigma)10. The liver was first perfused with a Per 1 Ready solution and tissue was digested using a Per II ready solution. A detailed list of components for both solutions can be found in Appendix 1. For PBMC and spleen, experiments for every timepoint were split into two with the processing of tissues from 3 mice at one time. For liver and lung processing, each group of 6 from each timepoint was processed at once. Repeat experiments were done if mice from a group did not survive the polytrauma, to reach a total of 6 mice per group.
We chose PBMC and spleen to characterize macrophages in the circulation. We then evaluated liver as a site of injury in this model, and the lung because of the link between the systemic immune response to trauma and the development of localized organ damage as seen in Acute Respiratory Distress Syndrome11.
4. Flow cytometry staining methods
The cells were counted and equally split into different FACS sample tubes and washed by centrifugation. Mouse Fc receptor blocking agent (BD Biosciences) was added to all tubes containing cells for 5 minutes at 4°C. Cells were then stained for viability (Live/Dead stain Thermofisher) and for macrophage surface markers CD68 (Biolegend) and F4/80 (BD bioscience). Samples were kept at 4°C for 30 minutes. Samples were then washed by centrifugation and cells were fixed with 2% paraformaldehyde (Thermo Scientific). The appropriate cells were permeabilized with BD Cytofix/Cytoperm (BD bioscience) for 30 minutes. After washing the cells with the permeabilization buffer and centrifugation, antibodies that stained intracellular markers were added for 60 minutes at 4°C. Those included iNOS (Thermofisher) as a marker for the M1 phenotype and Arg1 (R&D) as markers for the M2 phenotype12–14. The staining panel is included in Appendix 2. The samples were then analyzed with a BD LSR II flow cytometer, and results were analyzed with FlowJo version 10.7.1. Gating strategy is shown in Figure 1.
Figure 1: Gating strategy.
Gate 1: live cell determination. Gate 2: determination of single cell population. Gate 3: determination of macrophage CD68+ population. Gate 4.a (upper right): determining M1 macrophages (CD68+; iNOS+ cells). Gate 4.b (lower right): determining M2 macrophages (CD68+; Arg1+ cells).
5. Oxidative burst assay
Lung and liver non parenchymal cells were harvested from out of box control and polytrauma mice. After an initial wash, cells were incubated at 37°C for 5 minutes with or without DHR123 (50 μM - Thermofischer). Mouse Fc receptor blocking agent (BD biosciences) was added to stained tubes for 10 minutes at room temperature. The cells were stained for viability (Live/Dead stain kit – Thermofisher), PE/Cy7-labeled CD68 antibody (Biolegend) and BUV395-labeled F4/80 antibody (BD Biosciences). The cells were fixed with 2% paraformaldehyde (Thermo Scientific). The assay was verified using a positive and negative control. Our positive control consisted of cells stimulated with PMA (4 μM) or LPS (100 ng/mL), and our negative control consisted of cells coming from out of box mice, not treated with any stimulant. The samples were then analyzed with a BD LSR II flow cytometer immediately after staining and results were analyzed with FlowJo version 10.7.1.
6. Statistical analysis
All data was analyzed using Prism 9 version 9.0.2. All data is reported as mean ± standard deviation. Statistical outliers were identified through the ROUT method (robust regression and outlier removal) and removed before analysis. A Shapiro-Wilk test was run to determine normality. Based on the results, a one-way ANOVA with Tukey post-hoc testing or a Kruskal-Wallis test was performed for the examination of the dynamic change in phenotype or of function over time. An independent t-test or Mann-Whitney test was used to compare macrophages before and after trauma. Statistical significance was set at a p-value of 0.05.
7. Sample size
This study is novel in that it evaluates dynamic changes in macrophage populations over time. Since this was an exploratory study with no previously characterized or defined change between time points, we set out to use a sample size of 6 animals per group.
Results
1. Macrophage phenotyping
Macrophages in PBMC, spleen, lung and liver were isolated at three timepoints: before trauma, 24 hours after injury and late after trauma (7 days). We specifically looked at circulating macrophages (CD68+ cells) and characterized them as M1 (iNOS+) or M2 (Arginase1+). Values represented are percentages of all CD68+ cells within the organ of interest and are represented as mean +/− standard deviation.
a. M1 macrophages in different tissues over time (Figure 2):
Figure 2: M1 macrophages in tissues over time.
Data presented corresponds to mean +/− standard deviation. A: graph showing the presence of M1 macrophages in PBMC at different timepoints after polytrauma. B: graph showing the presence of M1 macrophages in spleen at different timepoints after polytrauma. Significant results through t-test analysis and one way ANOVA. C: graph showing the presence of M1 macrophages in lung at different timepoints after polytrauma. Significant results through t-test analysis. D: graph showing the presence of M1 macrophages in liver at different timepoints.
PBMC: peripheral blood mononuclear cells; Sham: out of box controls; PT: polytrauma.
In PBMC, at baseline, 6.6% (+/− 3.9%) CD68+ cells are of the M1 phenotype. That number slightly increases acutely after trauma, reaching 8.8% (+/− 11.0%) at 24 hours, and decreases again at 7 days post polytrauma (4.9% +/− 4.2%). These differences in average did not reach significance.
The spleen has a larger population of M1 macrophages at baseline (27.7% +/− 7.5%), with that level decreasing after trauma to 20.6% (+/− 13.2%) at 24 hours and 12.6% (+/− 5.6%, p = 0.0027 compared to out of box control) at 7 days.
A similar trend is seen in the lung. At baseline, 14.9% (+/− 8.7%) of CD68+ macrophages are M1, with a subsequent continual decrease reaching 8.7% (+/− 7.4%) at 24 hours and 4.4% (+/− 0.8%, p = 0.0265 compared to out of box control) 7 days after polytrauma.
M1 macrophages in the liver are 14.3% (+/− 7.5%) of the CD68+ population in liver non parenchymal cells at baseline, increase at 24 hours after trauma (20.8% +/− 9.7%), then decrease at 7 days post-polytrauma (13.4% +/− 5.7%). These changes were non-statistically significant.
b. M2 macrophages in different tissues over time (Figure 3):
Figure 3: M2 macrophages in tissues over time.
Data presented corresponds to mean +/− standard deviation.
A: graph showing the presence of M2 macrophages in PBMC at different timepoints after polytrauma. B: graph showing the presence of M2 macrophages in spleen at different timepoints after polytrauma. C: graph showing the presence of M2 macrophages in lung at different timepoints after polytrauma. D: graph showing the presence of M2 macrophages in liver at different timepoints. Results significant through one way ANOVA.
PBMC: peripheral blood mononuclear cells; Sham: out of box controls; PT: polytrauma.
There is minimal presence of M2 macrophages in circulating PBMCs and in spleen at baseline (<5%) and there is minimal variation after trauma. The M2 macrophages are present in larger proportion in the lung at baseline (7.2% +/− 3.9%). The percentage of M2 macrophages in lung remains relatively constant after trauma (7.2% +/− 6.3% at 24 hours and 9.2% +/− 8.6% at 7 days). A large proportion of M2 macrophages is also seen in the liver (36.0% +/− 12.7%). This percentage trends upwards and reaches 45.6% (+/− 11.5%) acutely after trauma and drops to 21.4 % (+/− 12.8%) at 7 days. These changes were non-statistically significant.
c. Ratio of M2/M1 macrophages in lung and liver over time (Figure 4):
Figure 4: Ratio of M2 to M1 percentages in lung and liver.
A: graph showing ratios of average M2 to average M1 in lung over time after polytrauma. B: graph showing ratios of average M2 to average M1 in liver over time after polytrauma.
Sham: out of box controls; PT: polytrauma.
The baseline ratio of the average lung M2/M1 is 0.48 demonstrating M1 macrophage predominance at baseline. This changes 24 hours after trauma with the ratio increasing to 0.84, as lung macrophages shift to an M2 phenotype. The ratio increases to 2.07 at 7 days after polytrauma, showing an even larger M2 predominance.
In the liver, the baseline ratio of the average lung M2/M1 is 2.53 demonstrating M2 macrophage predominance at baseline. This remains relatively constant 24 hours polytrauma, with a ratio of 2.19 and trending downwards to 1.88 7 days after polytrauma, indicating a trend towards an increase in M1 response.
2. Oxidative burst
We evaluated oxidative burst in liver and lung macrophages at baseline, 24 hours and 7 days after polytrauma. Results are shown in Figure 5.
Figure 5: Macrophage oxidative burst in lung and liver.
A: graph showing oxidative burst in lung macrophages over time after polytrauma. Results are significant through one way ANOVA. B: graph showing oxidative burst in liver macrophages over time after polytrauma.
Sham: out of box controls; PT: polytrauma.
At baseline, 2.0% (+/− 7.9%) of CD68+ lung cells performed oxidative burst. 24 hours after polytrauma, 6.9% (+/− 3.6%) of CD68+ lung macrophages were undergoing oxidative burst. At 7 days after polytrauma, 22.1% (+/− 11.5%) of lung macrophages were undergoing oxidative burst respectively (compared to baseline, 24 hours p=0.0028; day 7 p=0.0258).
In contrast, liver resident macrophages did not undergo a functional change after polytrauma. 31.8% (+/− 6.8%) of CD68+ macrophages underwent oxidative burst at baseline, 29.8% (+/− 5.3%) of macrophages underwent oxidative burst 24 hours after PT and 35.7% (+/− 10.7%) at 7 days after PT. The changes seen in the liver were non-statistically significant.
Discussion
Our present study characterized the distribution of macrophage phenotype and changes in polarization in specific organs in a murine model of polytrauma and hemorrhage. Myeloid cell contributors to systemic inflammatory response syndrome post trauma include mast cells15 and polymorphonuclear leukocytes16, but few studies have examined the association of macrophage polarization with post-traumatic immunopathology. We found a baseline presence of M1 macrophages in PBMCs, with the percentage only marginally changing over time after polytrauma. Very few M2 macrophages were found in the PBMC compartment. A similar trend is found in the spleen, where we discovered a high M1 presence that decreases over time after polytrauma, along with low M2 presence. The presence of M1 macrophages was consistent with our hypothesis, with the addition of a potential baseline priming effect secondary to spleen pathogen filtering. The white pulp of the spleen is where cells from the immune system will face foreign material and start responding to them17. The decrease in M1 macrophages in the spleen over time after trauma could be due to decrease in stimulation after the initial trauma or mobilization of these circulating cells to end organs. The low M2 macrophage presence in circulating PBMCs as well as the spleen was not unexpected, as these are not end organs requiring tissue repair or angiogenesis after injury.
The liver has the largest fixed macrophage population in the body, with tissue specific macrophages (Kupffer cells) accounting for 40 to 65% of the total liver non-parenchymal cells18. In this study, we evaluated the resident rather than the intravascular macrophages in the liver, because of the nature of the digestion of the tissue. We saw a baseline level of both M1 and M2 phenotype macrophages. This is expected as these cells are in contact with circulating blood from the portal vein and the hepatic artery and can therefore have a certain degree of activation, all while preserving a “tolerogenic” phenotype18. The liver also has a large regenerative capacity19, making the presence of the M2 phenotype at baseline reasonable.
We saw a baseline M1 macrophage phenotype present in the liver that is not significantly changed after polytrauma, and a baseline M2 phenotype that is stable in the first 24 hours post-trauma and decreases at 7 days post-trauma. This decrease caused the shift of the ratio of M2/M1 macrophages at 7 days after polytrauma towards the M1 phenotype. These observations point us to the fact that these changes might be specific to the crush injury as the response is different when compared to other models of liver injury such as the ischemia-reperfusion injury model. In the ischemia and reperfusion model, a predominance of M1 macrophages is seen in the early stages after the insult18, with a shift to an M2 predominant phenotype in later stages20. In contrast, the crush injury model showed an earlier infiltration of wound-healing macrophages and persistence of M1 macrophages for debris clearance. It is possible that there would be a change in the ratio towards the M2 phenotype, consistent with a “late stage” M2 predominance as seen in ischemia-reperfusion, if the mice were followed for a longer period of time.
Macrophage polarization in the lung after trauma is important to study, particularly in order to understand their role in the increased vulnerability of hospitalized trauma patients to nosocomial infections including pneumonia21. We found a baseline presence of both M1 and M2 macrophages in the lung. This is to be expected as the lung both clears respiratory pathogens and participates in tissue repair at baseline22. Interestingly, we saw a significant decrease in percentages of M1 macrophages after trauma in the lung. This was unexpected given the association of M1 macrophage polarization with systemic inflammation in the setting of insults such as pancreatitis15 and thermal injury16, and that macrophages isolated from post-traumatic patients exhibit enhanced inflammatory response to LPS17. This prompted us to look at their function at these different timepoints. Even though the number of M1 macrophages significantly decreased, we saw a significant increase in the oxidative burst capability of these macrophages. This could be demonstrative of a separate subset of macrophages, uncaptured by our phenotyping, with increasing in inflammatory capacity after injury. In our understanding of macrophage physiology, it is becoming increasingly evident that macrophage polarization is malleable. There is a multitude of different macrophage subtypes that could be responsible for this discrepancy between phenotype and function. These fall on a spectrum of polarization with clearly defined M1 and M2 being the two extreme poles, and further study using multiplexing assays such as CyTOF or RNA sequencing may better delineate these subtypes. The unexpected discrepancy between function and phenotype also prompts us to think about the possibility of using functional changes in macrophages rather than phenotypic changes to characterize macrophages. Targeting macrophages based on their function could also open up new therapeutic potentials.
Limitations
This study has several limitations. While we were able to identify the clearly polarized M1 and M2 macrophages in the different tissues, we were unable to capture the evolving phenotypes that have not reached a mature polarization state. These include different subtypes of macrophages that are on the spectrum of polarization. This study also did not look at macrophages through a single cell perspective which might have offered a more detailed phenotypic background. In addition to that, being that this study is the first to evaluate the dynamic changes in macrophage population over time after polytrauma, a sample size of 6 was chosen as a pilot study. These results will be utilized to power future analyses. In addition, the study was conducted with male C57B6 mice and therefore may not be generalizable. We also acknowledge that we chose to concentrate on CD68+ cells and did not focus solely on tissue resident (F4/80+) macrophages, in order to include evaluation of more immature circulating macrophages. We also chose to represent the M1 and M2 phenotypes by their most classical markers, as others (CCR2 and CD206) were much more variable in nature, possibly due to the fluctuating phenotype of these cells.
Conclusion
Macrophages play an important immunomodulatory role and their ability to respond to external stimulus allowed them to become a target of therapeutic potential. We evaluated macrophages after trauma in different organ tissues and focused on the two main phenotypes: the M1 proinflammatory phenotype and the M2 anti-inflammatory phenotype. We found a discrepancy in macrophage phenotype and function in the lung which may signify a change in subset polarization after injury. An important next step will be to confirm these findings in the setting of a secondary bacterial infection in the trauma recovery phase, to better understand the response to infection after trauma.
MDN has the following to disclose:
Scientific Advisory Board, Haima Therapeutics
Research funding: DoD, NIH, Janssen, Haemonetics, Instrumentation Laboratory
Honoraria: Meredian, Haemonetics, CSL Behring
Appendix 1:
Per 1 ready solution: Per 1 stock solution, diH2O, 2.23 uM ethylene glycol-bis(2-aminoethylether)-N,N,N’,N’-tetraacetic acid (EGTA).
Per 1 stock solution: diH2O, 1.42 M Sodium Chloride (NaCl), 67 mM Potassium Chloride (KCl), 100 mM HEPES.
Per 2 ready solution: Per 2 stock solution, 60 mg of Collagenase H (Roche cat#11087789001).
Per 2 stock solution: 19 mM calcium chloride, diH2O, 151.52 mM Albumin, 67 mM Sodium chloride, 6.7 mM potassium chloride, 100 mM HEPES.
Appendix 2: Flow cytometry markers
| Marker | Color | Product detail |
|---|---|---|
| Live/dead fixable stain – 405 nm excitation | Aqua | Thermofisher L34957 |
| F4/80 | BUV395 | BD bioscience 565614 |
| CD68 | PE-Cy7 | Biolegend 137016 |
| iNOS | FITC | Thermofisher 53-5920-82 |
| iNOS isotype control: Rat IgG2a kappa Isotype Control | Alexa Fluor 488 | Thermofisher 53-4321-80 |
| Arg1 | APC | R&D IC5868A |
| Arg1 isotype control | Sheep IgG | R&D 1C016A |
Footnotes
Conflict of Interest Declaration:
LH, PL, SH, DR, NA, EA, JB, AS, KW, JA, RS, JK, JAL, AS have no affiliation with or involvement in any organization or entity with any financial interest in the subject matter or materials discussed in this manuscript.
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