If you don't remember your password, you can reset it by entering your email address and clicking the Reset Password button. You will then receive an email that contains a secure link for resetting your password
If the address matches a valid account an email will be sent to __email__ with instructions for resetting your password
Macrophages are important for effective iron recycling and erythropoiesis, but they also play a crucial role in wound healing, orchestrating tissue repair. Recently, we demonstrated a significant accumulation of iron in healing wounds and a requirement of iron for effective repair. Herein, we sought to determine the influence of iron on macrophage function in the context of wound healing. Interestingly, wound macrophages extensively sequestered iron throughout healing, associated with a prohealing M2 phenotype. In delayed healing diabetic mouse wounds, both macrophage polarization and iron sequestration were impaired. In vitro studies revealed that iron promotes differentiation, while skewing macrophages toward a hypersecretory M2-like polarization state. These macrophages produced high levels of chemokine (C-C motif) ligands 17 and 22, promoting wound reepithelialization and extracellular matrix deposition in a human ex vivo wound healing model. Together, these findings reveal a novel, unappreciated role for iron in modulating macrophage behavior to promote subsequent wound repair. These findings support therapeutic evaluation of iron use to promote wound healing in the clinic.
Macrophages play a crucial role in orchestrating the wound healing response, fulfilling prodigiously versatile functions.
Most, but not all, wound macrophages originate from circulating bone marrow–derived monocytes. Once at the site of injury, these cells undergo marked, yet transient, morphologic and behavioral changes in response to the local wound milieu.
M1-stimulated macrophages are induced in early wound repair by proinflammatory mediators, such as lipopolysaccharides and interferon-γ, driving inflammation by secreting reactive oxygen species and proinflammatory cytokines (eg, IL-1, IL-6, and tumor necrosis factor-α
Thus, iron must be tightly regulated within the body to maintain normal physiology and avoid tissue pathology.
In the context of tissue repair, the cellular importance of iron is less clear. We have recently shown a substantial, previously unreported, accumulation of iron in the latter stages of wound healing that is attenuated in pathologic diabetic wounds.
The hypothesis of the current study is that iron plays an important role in modulating macrophage behavior to promote healing. Using a combination of in vitro and ex vivo studies, we explore the effect of iron on macrophage polarization, secretory profile, and ability to promote healing in a human ex vivo wound model. The data demonstrate an important new role for iron in mediating macrophage function and subsequent contribution to wound healing.
Materials and Methods
Animal Housing and Experimentation
Young (8- to 10-week–old) female wild-type (C57/Bl6), nondiabetic (NDb; Lepr−/+), and diabetic (Db; Lepr−/−) mice were purchased from Envigo Ltd. (Huntingdon, Cambridgeshire, UK) and housed at the University of Manchester Biological Services Facility. Mice received food and water ad libitum and were kept under a 12-hour light/dark cycle with constant temperature and humidity. Regulated procedures were performed according to UK Home Office regulations under project license 70/8136. Two equidistant 6-mm dorsal excisional wounds were made, as described previously.
Normal skin (day 0) and excisional wounds were collected and fixed in neutral-buffered formalin for histology.
Paraffin sections were cut (5 μm thick), dewaxed in xylene, and brought to water down an ethanol gradient. Antigen retrieval was performed with citrate buffer (pH 6) before primary antibody incubation at 4°C overnight. For immunofluorescence, anti-rabbit ferritin (clone EPR3004Y), anti-mouse α-smooth muscle actin (clone 1A4; Abcam, Cambridge, UK), and anti-rat Mac-3 (clone M3/84; BD Biosciences, Reading, UK) were used at 5 μg/mL. Anti-rabbit keratin 14 (clone Poly19053; Biolegend, San Diego, CA), anti-mouse COL1A1 (clone 3G3), and anti-mouse COL3A1 (clone C-6; Santa Cruz Biotechnology, Heidelberg, Germany) were used at 1 μg/mL. Alexa Fluor 488–conjugated (Mac-3, α-smooth muscle actin, and COL1A1) and Alexa Fluor 594–conjugated (ferritin, keratin 14, COL1A1, and COL3A1) secondary antibodies were used at 10 μg/mL for 1 hour at room temperature (all from Thermo Fisher Scientific, Paisley, UK). Slides were mounted with Mowiol 488 (Sigma-Aldrich, Dorset, UK) containing DAPI (Thermo Fisher Scientific). Images were taken on an LSM710 laser scanning confocal microscope (Carl Zeiss Ltd, Cambridge, UK) using a 20× objective and the 405-nm diode, 488-nm argon, and 561-nm Dulbecco's phosphate buffered saline lasers. Mac-3–, ferritin-, and α-smooth muscle actin–positive cells were counted in ImageJ software version 1.8.0 (NIH, Bethesda, MD; http://imagej.nih.gov/ij). For immunoperoxidase staining, anti-rat Mac-3 (above), anti-rabbit nitric oxide synthase 2, and anti-goat arginase-1 antibodies (both from Santa Cruz Biotechnology) were used at 1 μg/mL and detected using appropriate VECTASTAIN ELITE ABC kits and NovaRED peroxidase substrate (Vector Laboratories, Peterborough, UK). Bright-field images were obtained on a Nikon E400 microscope (Nikon Instruments Inc., Melville, NY) and SPOT camera (SPOT Imaging, Sterling Heights, MI). Cell counts were performed in ImagePro-Plus version 6.3.0 (Media Cybernetics, Cambridge, UK).
THP-1 Cell Culture
THP-1 cells (ATCC TIB-202; ATCC, Middlesex, UK) were used as a human monocyte model.
THP-1 cells were grown in RPMI 1640 medium (Thermo Fisher Scientific) with 10% heat-inactivated fetal bovine serum (Thermo Fisher Scientific) and 1% penicillin-streptomycin solution (Thermo Fisher Scientific).
THP-1 Cell Differentiation Experiments
Phorbol 12-myristate 13-acetate (PMA) was used at 200 ng/mL to induce differentiation in THP-1 cells. THP-1 cells were treated with or without PMA and iron in the form of ferric ammonium citrate (FAC) at 10 and 100 μmol/L concentrations. After 3 days of incubation, THP-1 cells were collected for RNA (below) to assess the expression of the differentiation markers, CD11b, CD14, CD36, and CD54, via quantitative real-time PCR.
After PMA stimulation for 3 days to induce differentiation, THP-1–derived macrophages (TDMs) were stimulated to M1 (interferon-γ and lipopolysaccharide) and M2 (IL-4 and IL-13) states using reagents at 20 ng/mL concentration (all from Thermo Fisher Scientific) or left unstimulated (M0 treatment group). For these polarization experiments, TDMs were treated with FAC at the same time as adding cytokines. TDMs were collected after 24 hours for quantitative real-time PCR, immunocytochemistry (using ferritin antibody, as above), and Western blot analysis.
Quantitative Real-Time PCR
Macrophages were collected in Ambion TRIzol reagent (Thermo Fisher Scientific), and RNA was isolated using phenol/chloroform.
RNA was purified using the Ambion PureLink Mini Kit (Thermo Fisher Scientific), as per manufacturer's instructions, and reverse transcribed with Bioscript (Bioline, London, UK) and Random Primers (Promega, Southampton, UK). Quantitative real-time PCR was performed with 2× Takyon SYBR Green mastermix (Eurogentec, Hampshire, UK) and a CFX Connect thermocycler (Bio-Rad Laboratories, Hertfordshire, UK). Primer sequences are provided in Table 1.
Table 1Primer Sequences for Quantitative Real-time PCR
THP-1 cell adhesion was assessed after PMA and FAC stimulation (as per differentiation experiments). Herein, wells were rinsed with Dulbecco's phosphate-buffered saline (Thermo Fisher Scientific) to remove nonadherent cells; and CellTiter 96 AQueous One Solution (Promega) was added to the wells, as per manufacturer's guidelines. Zymography was also performed on conditioned medium (CM) collected from these THP-1 cells, as described previously.
Human skin was collected from theaters at Castle Hill Hospital (Cottingham, Hull, UK) under local research ethics committee (17/SC/0220) approval and full, informed patient consent. Human dermal fibroblasts (HDFs) were isolated, as previously described.
HDFs were cultured in minimum essential medium (Gibco, Thermo Fisher Scientific) in 2% fetal bovine serum with FAC for 11 days. Cells were removed (denuded) from the HDF-derived ECM with 20 mmol/L ammonium hydroxide (Sigma-Aldrich) in 0.5% Triton X-100 in Dulbecco's phosphate-buffered saline. Denuded ECM was treated with 10 μg/mL DNAse (Sigma-Aldrich) and extensively washed. THP-1 cells were seeded on this HDF-derived ECM with or without 200 ng/mL PMA, left for 3 days, and collected for differentiation assessment via quantitative real-time PCR. Immunocytochemistry and Perls Prussian Blue staining were performed on the HDF-derived ECM.
A ferrozine assay allowed for the detection of ferrous and ferric iron.
Denuded HDF-derived ECM was scraped on ice in 50 mmol/L NaOH containing 1% HALT protease inhibitor cocktail (Thermo Fisher Scientific). HCl (10 mmol/L) was added to each lysate with 200 μL iron-releasing reagent (8.6% v/v HCl and 280 μmol/L KMnO4 in distilled water) for 2 hours at 60°C. Standards of known concentrations of FAC (100 mmol/L, 10 mmol/L, 1 mmol/L, 100 μmol/L, 10 μmol/L, 1 μmol/L, and 0 μmol/L) were prepared and incubated, as described. Samples were cooled at room temperature, and 50 μL iron detection reagent (650 μmol/L ferrozine, 650 μmol/L neocuproine, 2.8 mmol/L ammonium acetate, and 1 mmol/L ascorbic acid in distilled water) was added at room temperature for 30 minutes. Collected supernatant was read at 570 nm, and cellular iron concentration (in μmol/L) was calculated from the FAC standard curve.
Perls Prussian Blue Staining
Perls Prussian Blue staining was used to visualize iron deposits. HDFs were left on their deposited ECM or removed from their ECM before staining. Plates were fixed (4% formaldehyde solution) and incubated in a 50:50 solution of 5% (w/v) potassium ferricyanide (in distilled water; Sigma-Aldrich) and 5% (v/v) hydrochloric acid for 10 minutes at room temperature, then rinsed in distilled water
Bone marrow was isolated from NDb and Db mice with Dulbecco's modified Eagle's medium (Gibco, Thermo Fisher Scientific). Mononuclear phagocytes were differentiated to macrophages with L929 (CCL-1; ATCC) CM, containing macrophage colony-stimulating factor. Macrophages were differentiated for 7 days, serum starved, and then stimulated. Macrophages were left unstimulated (M0) or polarized to M1 or M2 states
in the presence of FAC and left for 24 hours before collecting for quantitative real-time PCR.
Western Blot Analysis
Western blot analysis was performed on FAC-treated polarized TDMs. Blots were probed with anti-mouse ferritin (clone B-12; Santa Cruz Biotechnology) at 1 μg/mL concentration for 1 hour at room temperature and detected with SuperSignal West Pico PLUS Chemiluminescent Substrate (Thermo Fisher Scientific).
THP-1 cells were differentiated and polarized, as described above, but in Transwell inserts (0.4 μm; Corning, Flintshire, UK). After polarization, inserts were washed in Dulbecco's phosphate-buffered saline, to remove exogenous cytokines and FAC, and transferred to plates seeded with primary HDFs. THP-1 cell–containing inserts were replaced with fresh inserts after 3 days of culture. On day 6, Transwell inserts were removed and HDFs were fixed and stained using anti-mouse collagen type I primary antibody at 1 μg/mL concentration (clone 3G3; Santa Cruz Biotechnology). Images were taken via confocal microscopy (above) and analysis of collagen production performed in ImageJ version 1.8.0. Quantitative real-time PCR was also performed on HDFs after Transwell co-culture.
Proteome Profiler Chemokine Arrays
TDMs were polarized in the presence of FAC for 24 hours, washed extensively, and then given fresh RPMI 1640 media with no cytokines for 2 days. This medium was collected and sterile filtered before performing Proteome Profiler chemokine arrays (Biotechne, Oxford, UK), as per manufacturer's instructions.
Human ex Vivo Wounding
Human skin was collected, as described above, and defatted before washing in 2% (v/v) antibiotic-antimycotic solution (Thermo Fisher Scientific). Partial-thickness wounds were generated (2-mm diameter) in the center of 6-mm–diameter skin explants using sterile biopsy punches (Stiefel Laboratories, Research Triangle Park, NC). Wounds were cultured in Petri dishes on a stack of two absorbent pads and a nitrocellulose 0.22-μm membrane using Dulbecco's modified Eagle's medium with 10% fetal bovine serum and 1% antibiotic-antimycotic solution. Wounds were treated with a vehicle (1% bovine serum albumin plus 0.002% dimethyl sulfoxide), 100 nmol/L CCR4 antagonist (C 021 dihydrochloride), 50 ng/mL CCL17, and/or 50 ng/mL CCL22 (R&D Systems, Oxford, UK), applied topically every 2 days. Wounds were collected in 4% neutral-buffered formalin 3 days after wounding for histologic analysis.
Means ± SEM were used to present all data. One- and two-way analyses of variance were performed where applicable, with appropriate post-hoc analyses. Significance was deduced at P < 0.05. Statistical analysis was performed in GraphPad Prism version 7.0 (GraphPad Software, San Diego, CA).
Macrophages Accumulate Iron across a Murine Healing Time Course
We previously demonstrated the temporal accumulation of iron in wound repair.
Herein, the cellular nature of this accumulated iron (measured as bioavailable ferritin) was assessed across a wound healing time course. Total ferritin was significantly increased on day 3 (P < 0.001) and day 7 (P < 0.001) after injury (Figure 1, A–C ), with a large proportion of iron sequestration occurring in macrophages (P < 0.001) (Figure 1D) and, to a lesser extent, α-smooth muscle actin–positive fibroblasts (P < 0.05) (Figure 1E). Of note, wound macrophage phenotype changes significantly over this time frame; nitric oxide synthase 2–positive (proinflammatory) macrophages peaked at day 3 after injury, whereas arginase-1–positive (anti-inflammatory) macrophages dominated at day 7 after injury (Supplemental Figure S1). Hence, wound iron preferentially accumulated in macrophages, predominately correlating with tissue infiltration of prohealing (M2) macrophages.
Impaired Macrophage Iron Accumulation in Diabetic Wounds Correlates with Altered Macrophage Polarization
In both Db and NDb mouse wounds, ferritin increased at day 3 (P < 0.01 and P < 0.05) and day 7 (P < 0.001 and P < 0.01) after injury, respectively (Figure 2, A and B ). However, the absolute ferritin level was reduced in Db mice at all time points examined, significant at day 7 after injury (P < 0.05). Interestingly, although the proportion of wound tissue ferritin-positive macrophages was approximately equal between Db and NDb mice day 3 wounds, there was a clear reduction in Db macrophage iron sequestration at day 7 after injury (Figure 2C). This reduced proportion of ferritin-positive macrophages in day 7 Db wounds was even more meaningful given that total macrophages (Figure 2E) and nitric oxide synthase 2–positive cells (M1 macrophage marker) (Figure 2F) were both increased in Db mouse wounds. By contrast, wound arginase-1–positive cells (M2 macrophage marker) were reduced in day 7 Db versus NDb wounds (P < 0.05) (Figure 2, D and G), mirroring ferritin-positive macrophages (Figure 2C). Collectively, these data reveal altered macrophage iron sequestration and polarization in Db wounds.
Endogenous Iron or Iron-Rich Extracellular Matrix Promotes Macrophage Differentiation
The tissue microenvironment drives monocyte to macrophage differentiation.
We, thus, investigated the influence of exogenous iron on macrophage differentiation in THP-1 cells in vitro. Herein, iron (100 μmol/L FAC) administration significantly potentiated THP-1 monocyte differentiation in response to PMA treatment, marked by enhanced expression of the differentiation markers CD11b (P < 0.001), CD14 (P < 0.001), CD36, and CD54 (P < 0.001) (Figure 3, A–D ). FAC also increased cellular adhesion, a phenotypic marker of differentiation (P < 0.001) (Figure 3E) and significantly induced cellular matrix metalloproteinase 9 activity (P < 0.001) (Figure 3, F and G).
In the wound environment, macrophages will be exposed to iron deposited in the ECM; it was, therefore, tested how ECM derived from FAC-treated HDFs influenced macrophage differentiation. HDFs treated with FAC deposited an extensive ECM (Figure 3H) rich in iron, as demonstrated by both Perls Prussian Blue staining (Figure 3I) and ferrozine assay (Figure 3J). THP-1 monocytes were seeded onto this denuded ECM and stimulated with PMA to induce differentiation. Exposure to the iron-rich ECM significantly increased expression of the differentiation markers CD11b (P < 0.001), CD14 (P < 0.05), CD36 (P < 0.001), and CD54 (Figure 3, K–N). Clearly, both exogenous iron treatment and exposure to iron-rich ECM stimulate macrophage differentiation.
Iron Drives Cytokine-Stimulated Macrophages toward a Prohealing Phenotype
FAC treatment (100 μmol/L) led to a significant decrease in the M1 markers IL-1β (P < 0.01) (Figure 4A) and tumor necrosis factor-α (P < 0.001) (Figure 4B) in M1-stimulated TDMs, whereas the expression of the M2 markers CCL17 (P < 0.001) (Figure 4C) and CCL22 (P < 0.001) (Figure 4D) was elevated in M2-stimulated TDMs after 100 μmol/L FAC. Similar effects were observed in NDb murine bone marrow–derived macrophages (BMDMs), where Il-1β expression was reduced at both FAC concentrations (P < 0.001) (Figure 4E). Db BMDMs were less responsive to iron, with Il-1β expression only reduced by 100 μmol/L FAC treatment (P < 0.05) (Figure 4I) whereas FAC treatment modestly reduced Tnf in NDb and Db BMDMs (Figure 4, J and L). Intriguingly, 100 μmol/L FAC treatment elevated Ccl17 and Ccl22 in M0, M1, and M2 polarized BMDMs from NDb and Db mice (Figure 4, G, H, K, and L). Together, these data reveal that iron skews cytokine profiles in both TDMs and BMDMs.
Iron-Loaded M2 Macrophages Secrete Factors that Stimulate Extracellular Matrix Production
TDMs treated with 100 μmol/L FAC showed significantly greater ferritin storage in M0 (P < 0.001) and M2 (P < 0.001) states (Figure 5, A and B ) by immunohistochemistry, and in all three activation states via Western blot analysis (P < 0.01 to P < 0.001) (Figure 5C). For both techniques, the highest levels of ferritin were observed in 100 μmol/L FAC–treated M2-polarized TDMs. As macrophages play a key role in ECM remodeling,
a Transwell co-culture approach was used to test the paracrine effects of FAC-stimulated TDMs on HDF ECM production. Transwell co-culture of M2-stimulated FAC-treated TDMs resulted in up-regulation of COL1A1 (P < 0.05) (Figure 5D), COL1A2 (P < 0.001) (Figure 5E), and COL3A1 (P < 0.05) (Figure 5F) expression in HDFs. Collagen I protein levels were also highest in HDFs co-cultured with M2-stimulated, FAC-treated TDMs (P < 0.001) (Figure 5, G and H). Finally, CM from iron-treated M2-polarized TDMs was profiled using chemokine arrays to characterize secreted factors of interest (Figure 5I). Herein, CM from M2-stimulated TDMs treated with 100 μmol/L FAC displayed higher levels of multiple chemokines, including CCL17 and CCL22.
CCL17 and CCL22 Promote Healing and ECM Deposition in Human ex Vivo Wound Repair
To test the biological significance of TDM-derived chemokines, an ex vivo wound model was used. Given that the CCL17-CCR4 receptor axis has previously been suggested to be important for healing,
were applied topically to human ex vivo wounds. Wound closure (re-epithelialisation) was significantly increased after treatment with CCL17 (P < 0.01) or CCL22 (P < 0.05) in isolation and by a combination of both ligands (P < 0.001) (Figure 6, A and B ). Although the CCR4 antagonist (C 021 dihydrochloride) had no direct effect on wound closure, it significantly blocked the healing promoting effect of CCL17 (P < 0.05) and a combination of both ligands (P < 0.01). Finally, CCL17 and CCL22 effects on ECM production were evaluated. Collagen I and collagen III contents were assessed from three regions close to the wound surface (Figure 6C). Collagen I was significantly increased after treatment with CCL17 (P < 0.05) or a combination of both ligands (P < 0.05), an effect that was again reversed by cotreatment with the antagonist, C 021 dihydrochloride (Figure 6, D and E). Collagen III was significantly elevated after treatment with both ligands alone and in combination (P < 0.05), which was again blocked by the receptor antagonist, C 021 dihydrochloride (Figure 6, D and F). Together, these data reveal a novel, previously unappreciated role for the macrophage-derived chemokines, CCL17 and CCL22, in promoting human wound reepithelialization and matrix deposition.
Macrophages are closely associated with the biological availability of iron. In addition to modulating heme storage and erythrocyte recycling,
yet how macrophage behavior is influenced by the complex wound microenvironment remains far from clear. Herein, we present new experimental findings linking iron modulation of macrophage behavior and the remodeling phase of wound repair.
We previously revealed temporal accumulation of iron in late-stage wound repair.
In concurrence, the present study revealed that exposure to high iron increased macrophage differentiation, but also significantly extended this observation to demonstrate a role for iron in modulating macrophage polarization and chemokine secretion.
Macrophages are able to perform versatile functions with high plasticity to allow effective repair. Functionally, CD11b-specific
ablation leads to delayed healing and excessive inflammation. This illustrates not only the importance of macrophage presence, but also effective temporal modulation of macrophage phenotype in wound healing. Although not extensively characterized, the late-stage M2 switch may be driven by degradation of M1 receptors
In the current study, it is shown, for the first time, that iron treatment of macrophages significantly potentiated the release of a range of chemokines, which promoted collagen production in HDFs. M2 macrophage CM is known to stimulate collagen 1 and collagen 3 production in fibroblasts in vitro,
These new data suggest that iron indirectly stimulates HDF ECM deposition via modulation of macrophage behavior, with potential therapeutic application in pathologic healing.
The iron-treated M2 macrophage screen identified the chemokines CCL17 and CCL22. Both CCL17 and CCL22 were shown to directly promote reepithelialization and collagen deposition in a human ex vivo wound repair model. This is in line with previous work in HaCaTs,
demonstrated improved healing in a CCR4−/− type 1 diabetes mellitus model and in wild-type type 1 diabetes mellitus mice treated with anti-CCL17 and anti-CCL22 antibodies. Thus, future studies investigating macrophage iron processing should be extended to address tissue- and pathology-specific differences.
In summary, these new data demonstrate a key role for iron in modulating macrophage responses to the wound environment. Iron skewed macrophages toward an M2-like state that produced a secretome highly enriched with ECM-stimulating factors, including the chemokines, CCL17 and CCL22. It is tempting to speculate that local iron administration, perhaps targeted to the latter stages of wound healing, could have clinical benefit. Further work is now required to explore how wound iron manipulation can be used as a conduit to improve skin healing pathology.
We thank Dr. Barbara Guinn for helpful discussions.
H.N.W. and M.J.H. designed the study; H.N.W., E.R.R., and A.R.S. acquired data; H.N.W., E.R.R., and K.L.B. analyzed the data; H.N.W. prepared the manuscript; H.N.W., E.R.R., K.A.M., and M.J.H. revised the manuscript; K.A.M. and M.J.H. supervised the study; all authors read and approved the final manuscript; M.J.H. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.