- Research article
- Open Access
- Open Peer Review
Monocyte depletion increases local proliferation of macrophage subsets after skeletal muscle injury
© Côté et al.; licensee BioMed Central Ltd. 2013
- Received: 2 July 2013
- Accepted: 8 December 2013
- Published: 19 December 2013
Sequential accumulation of M1 and M2 macrophages is critical for skeletal muscle recovery after an acute injury. While M1 accumulation is believed to rely on monocyte infiltration, the mechanisms of M2 accumulation remain controversial, but could involve an infiltrating precursor. Yet, strong depletion of monocytes only partially impairs skeletal muscle healing, supporting the existence of alternative mechanisms to palliate the loss of infiltrating macrophage progenitors. The aims of this study are thus to investigate if proliferation occurs in macrophage subsets within injured skeletal muscles; and to determine if monocyte depletion leads to increased proliferation of macrophages after injury.
Injury was induced by bupivacaine injection in the tibialis anterior muscle of rats. Blood monocytes were depleted by daily intravenous injections of liposome-encapsulated clodronate, starting 24 h prior to injury. In separate experiments, irradiation of hind limb was also performed to prevent resident cell proliferation. Upon euthanasia, blood and muscles were collected for flow cytometric analyses of macrophage/monocyte subsets.
Clodronate induced a 80%-90% depletion of monocyte but only led to 57% and 41% decrease of M1 and M2 macrophage accumulation, respectively, 2 d following injury. Conversely, the number of M1 macrophages in monocyte-depleted rats was 2.4-fold higher than in non-depleted rats 4 d after injury. This was associated with a 16-fold increase in the number of proliferative M1 macrophages, which was reduced by 46% in irradiated animals. Proliferation of M2 macrophages was increased tenfold by clodronate treatment 4 d post injury. The accumulation of M2 macrophages was partially impaired by irradiation, regardless of monocyte depletion.
M1 and M2 subsets proliferate after skeletal muscle injury and their proliferation is enhanced under condition of monocyte depletion. Our study supports the conclusion that both infiltrating and resident precursors could contribute to M1 or M2 macrophage accumulation in muscle injury.
- Flow cytometry
Macrophages are classically known for their pro-inflammatory roles in innate immunity and more recently for their active contribution to the resolution of inflammation and tissue repair [1–6]. This versatility is reflected by their ability to adopt distinct phenotypes depending on the microenvironment [7–9]. Macrophages can be divided in two main subsets according to their mode of activation and specific functions. In the context of skeletal muscle tissue injury, “classically activated”  M1 macrophages are found during the inflammatory phase and are associated with phagocytosis, while “alternatively activated”  M2 macrophages accumulate at the site of injury once necrotic tissue has been removed and participate to the repair and remodeling processes [7–9]. In rats, former nomenclature ED1+ and ED2+ refers to M1 (CD68+ CD163-) and M2 (CD68+ CD163+) macrophage subsets, respectively. For the sake of clarity we will thereafter refer to M1 and M2 macrophages. Macrophages insuring homeostasis of uninjured muscles are believed to be resident cells and would present a M2 phenotype [7, 9]. Macrophage involvement is clearly a prerequisite for skeletal muscle repair and their deregulation can determine the outcome of skeletal muscle healing. The current dogma is that macrophages must freely accumulate in the injured muscle to ensure an adequate healing [1–3, 11–13]. However, depletion of macrophages has only led to partial alteration of skeletal muscle recovery after injury , , supporting the existence of alternative mechanisms to ensure the functions of macrophages. Given the critical involvement of macrophage subsets in skeletal muscle healing, a better understanding of the mechanisms governing their accumulation may reveal new points of regulation for intervention.
The mechanisms of M1 and M2 macrophage accumulation after sterile injury remain elusive. In models like peritoneal infection, M1 or M2 macrophage accumulation results from the differentiation of distinct infiltrating M1 and M2 myeloid precursors . In contrast, recent evidences support in situ proliferation of M1 and M2 macrophage subsets in TH2-mediated inflammation [14, 15]. In the context of muscle injury, M1 macrophage accumulation is thought to result exclusively from the infiltration and differentiation of a precise monocyte subset into macrophages. On the other hand, the origin of the increased number of M2 macrophages is a matter of debate. A number of hypotheses are currently put forward including sequential mobilization of M1 and M2 macrophage circulating precursors , differentiation of monocyte-derived M1 macrophages into M2 macrophages following phagocytic activity after skeletal muscle injury  or M2 macrophage proliferation . It appears that the mechanisms of M1 and M2 accumulation in this specific situation are complex, sometimes overlapping, and most likely determined by the context of the immune response.
The importance of sequential accumulation of M1 and M2 macrophages for optimal muscle healing is now well accepted. However, the cellular origin and respective contribution of proliferation vs. infiltration remain elusive following skeletal muscle injury. In addition, there is no information on how those mechanisms might be altered under anti-inflammatory conditions. The goal of this study was to determine if local proliferation could contribute to M1 and M2 macrophage accumulation following skeletal muscle injury, under normal or monocyte depletion conditions. Given that M1 macrophage accumulation in the context of sterile skeletal muscle injury is believed to rely exclusively on monocyte infiltration [2, 16], and that M2 macrophage accumulation could be derived from M1 , our working hypothesis is that following sterile muscle injury in rat, blood monocyte depletion will impair M1 and M2 tissue macrophage accumulation.
Female Wistar rats weighing between 125 – 150 g were purchased from Charles River (St-Constant, QC, Canada) and housed 3 per cage. They were maintained on a 12 h – 12 h light – dark cycle. Water and food were provided ad libitum. All care, handling and experimental procedures were approved by the Université Laval Research Center Animal Care (Permit Number: 2012–025) and Use Committee according to the guidelines of the Canadian Council on Animal Care.
At 0 d, the right tibialis anterior (TA) muscle was chemically injured with bupivacaine. Rats received 0.05 mg/kg buprenorphine (Temgesic®, Reckitt Benckiser Healthcare (UK) Ltd) intraperitoneally (i.p.) as an analgesic 15 min before surgery and were then anesthetized with 1.5 – 2% isoflurane (Abbott Laboratories, Montreal, QC, Canada) under a flow of 400 – 800 mL/min of oxygen. Fur of the anterior side of the right hind limb was shaved off and skin was disinfected with isopropyl alcohol. Then, 120 μL of bupivacaine hydrocloride 0.5% (Marcaine; Hospira, Lake Forest, IL, USA) was injected through the skin in 3 sites of 40 μL along the right TA using a syringe with a 29 G needle. A single dose of 0.05 mg/kg of buprenorphine was also administrated i.p. 1 d after the injury.
Liposome-encapsulated dichloromethylene diphosphonate (Cl2MDP; clodronate) was used to deplete blood monocytes/macrophages . Clodronate was a gift from Roche Diagnostics GmbH (Mannheim, Germany). Rats received 1 mL, at the concentration established by the provider, of the liposome-encapsulated clodronate suspension. Injections were made daily in the tail vein or in the internal jugular veins under anesthesia with isoflurane starting 24 h before injury until sacrifice at 1, 2, 3 or 4 d post-injury.
Single leg irradiation
Liposome-encapsulated clodronate does not deplete resident macrophages in muscle. To study the contribution of resident macrophages, irradiation was performed since it prevents proliferation by inducing DNA damage, especially in immune cells [18, 19]. 24 h before injury, rats were anesthetized by i.p. injection of a ketamine/xylazine cocktail (80 mg/kg ketamine and 10 mg/kg xylazine) and then installed under a 4 cm-thick lead shield plate with their right hind limb facing a hole allowing radiation to pass for local irradiation. The right leg received a single dose of 20 Gy of γ rays delivered at 1.1 Gy per minute using a Gammacell® 40 Exactor (Best Theratronics Ltd, Ottawa, ON, Canada). This dose prevents resident leukocyte replication without killing mature muscle cells [18, 20–23].
Isolation of peripheral blood mononuclear cells (PBMCs)
5 mL of blood was obtained by cardiac puncture under anesthesia with isoflurane and collected in BD Vacutainer® blood collection tubes containing EDTA (BD, Franklin Lakes, NJ, USA). PBMCs were isolated with a density-gradient centrifugation on Histopaque 1083 (Sigma-Aldrich, St. Louis, MO, USA). Blood was deposited on 3 mL of Histopaque 1083 in a 15 mL conical tube and centrifuged for 30 min at 400 × g at room temperature. The upper layer was discarded and the opaque interface containing mononuclear cells (buffy coat) was aspirated with a Pasteur pipette and transferred to a new 15 mL centrifuge tube. Cells were washed twice with PBS and contaminating erythrocytes were lysed with a 4 min incubation with 1 mL of sterile erythrocyte lysis buffer containing 155 mM NH4Cl, 10 mM KHCO3 and 0.342 mM EDTA. 10 mL of PBS was then added to stop the lysis. After washing twice with PBS, cells were resuspended in 0.5 mL PBS and counted with a hemocytometer using trypan blue exclusion.
Preparation of single cell suspension from TA muscle
Rats were euthanized by cervical dislocation under anesthesia with isoflurane. TA muscles were dissected and rinsed in PBS. Minced muscles were incubated for 3 h at 37°C with 5 mL of Roswell Park Memorial Institute media 1640 (RPMI 1640) (HyClone, Logan, UT, USA) containing 3 mg/mL collagenase D (Roche Diagnostics, Laval, QC, Canada). Collagenase-digested samples were then homogenized by trituration using a 1000 μL micropipette. 10 mL PBS-EDTA (26 mM EDTA in PBS) was added to this suspension and centrifuged at 500 × g for 5 min at 4°C. After red blood cell lysis, the pellet was resuspended in 30% isotonic Percoll (GE healthcare, Waukesha, WI, USA) and centrifuged at 500 × g for 15 min at 4°C. The supernatant was discarded and the pellet resuspended in 1 mL PBS-EDTA. Cells were counted with a hemocytometer using trypan blue exclusion.
Flow cytometric analyses
Sequential extracellular and intracellular stainings were performed as described earlier . Single cell suspensions were washed in staining buffer (PBS containing 2% FBS). Cells were then resuspended in staining buffer containing Fc Block (anti-CD32) (BD Biosciences, Mississauga, ON, Canada). A cocktail of antibodies containing Pacific Blue-coupled anti-CD11b (Serotec, Kidlington, UK), biotin-coupled anti-CD3 (Serotec, Kidlington, UK), biotin-coupled anti-CD45RA (Biolegend, San Diego, CA, USA), Alexa Fluor® 647-coupled anti-CD163 (Serotec, Kidlington, UK) or Alexa Fluor® 647 coupled-IgG1 isotype control (Serotec, Kidlington, UK) was added to the cell suspension and incubated for 20 min on ice. Cells were washed in staining buffer and incubated with Alexa Fluor® 700-coupled streptavidin (Invitrogen Life Technologies, Burlington, Canada) for 20 min on ice. Cells were washed in staining buffer and fixed in PBS containing 2% paraformaldehyde for 20 min on ice, and then permeabilized in staining buffer containing 0.2% saponin for staining with anti RPE-coupled anti-CD68 (Serotec, Kidlington, UK) and Pe-Cy7-coupled anti-Ki-67 (BD Biosciences, Mississauga, ON, Canada) or RPE-coupled IgG1 isotype control (Serotec, Kidlington, UK) and Pe-Cy7-coupled k IgG1 isotype control (BD Biosciences, Mississauga, ON, Canada). Intracellular staining was performed for 30 min on ice. Single color controls were performed using CompBead Plus (BD Biosciences, Mississauga, ON, Canada). Data was acquired with a FACS Diva-driven FACS Aria II (Becton Dickinson) and analyzed with FlowJo (Tree Star Inc. Ashland, OR, USA). Isotype controls or omission of primary antibodies (when appropriate) were used to set gates. To determine numbers of specific cell subsets (from blood or skeletal muscles), the percentages obtained from flow cytometric analyses were multiplied by the absolute cell numbers obtained from the hemocytometer count.
All values are expressed as means and standard error. The construct of these experiments allowed comparisons between groups to be performed by Student’s t-test or one-way ANOVA followed by Tukey-Kramer post-hoc test, when appropriate (InStat GraphPad Software Inc., La Jolla, CA, USA). Significance was defined as p < 0.05.
Blood monocyte depletion modifies macrophage accumulation in injured muscle
Monocyte depletion is associated with proliferation of macrophages in injured muscles
The M2 Marker CD163 is not expressed by circulating monocytes
Macrophages adopt various phenotypes based on their environment and participate to many physiological processes [4, 8, 28, 29]. Substantial efforts have been deployed to characterize these various phenotypes with the hope that a better understanding of macrophage subsets could help delineate detrimental from the beneficial effects. It is now known that both M1 and M2 macrophages positively influence myogenesis, but can also contribute to pathological conditions [30–33]. Although mechanisms of macrophage replenishment were described in other tissues [34–36], the intrinsic mechanisms supporting their accumulation in the specific context of muscle injury are still a matter of debate and how these mechanisms are modulated under various anti-inflammatory strategies remains unknown. The main findings of this paper are that following muscle injury 1) both M1 and M2 macrophages can proliferate locally and 2) the major source of M1 macrophages are circulating precursors while M2 accumulation relies about equally on local (radiation sensitive) and infiltrating precursors. 3) We show for the first time that proliferation of macrophages is increased under condition of monocyte depletion.
We observed a low but quantifiable proliferation of M1 and M2 macrophage subsets following muscle injury at 2 d post-injury in non-depleted animals. This is in accordance with the reported proliferative ability of a number of macrophage subtypes including macrophages located in brain [35, 37] or peritoneum , bone marrow-derived macrophages in culture , and the macrophage cell line RAW-264.7 . Moreover, Hashimoto et al. have recently demonstrated that some tissue-resident macrophages repopulate locally via cellular proliferation with minimal contribution from monocytes, but these findings were observed under steady-state situation in alveolar macrophages, which are known to possess unique features such as a half-life 3 to 4 times longer than other macrophage subsets . Ajami et al. also found no evidence of microglia progenitor recruitment from the circulation, but these results were observed in denervation and CNS neurodegenerative disease models . Macrophage proliferation was also observed during chronic inflammation  and Khmelewski et al. (2004) revealed that both subsets of macrophages accumulated around collateral vessels in spite of monocyte depletion during femoral artery occlusion-induced arteriogenesis . However, in these studies the authors failed to discriminate the proliferative capacity of M1 and M2 macrophages [41, 42]. Jenkins et al. (2011) clearly showed that tissue macrophages with M2-like phenotype and M1 macrophages can undergo proliferation, but this was done in a model of TH-2-driven inflammation where the presence of IL-4 was essential . Thus, macrophage proliferation has been demonstrated in various tissues and experimental models quite distinct from the one presented here. This is the first evidence that macrophage proliferation occurs specifically in skeletal muscle tissue and that proliferation contributes to macrophage accumulation following muscle injury. Our results are therefore unique and represent, to the best of our knowledge, the first demonstration that proliferation of M1 and M2 macrophages normally occurs following sterile muscle injury.
M1 macrophage accumulation
Up to now, most studies pointed toward the infiltration of “inflammatory monocytes” to explain the accumulation of tissue macrophages after non-infectious tissue injury like skeletal muscle trauma. However, recent evidence shows that macrophages can undergo local proliferation in TH2-mediated inflammation . This prompted us to verify the effect of sustained monocyte depletion, in an attempt to delineate if monocyte infiltration is the sole contributor to macrophage accumulation in muscle injury. In accordance with other studies [3, 43, 44], monocyte depletion induced a significant decrease in the absolute number of M1 macrophages present at 2 d post-injury. We surprisingly observed a higher number of M1 macrophages observed in the monocyte-depleted animals at 4 d post-injury, when compared to non-depleted injured animals. It was very unlikely that this phenomenon could be explained by an increased recruitment at 4 d, since current literature suggests that signals for monocyte recruitment are not produced at this time point . Importantly, numerous reports showed delayed peaks of macrophage accumulation following acute muscle injury when using diverse anti-inflammatory strategies; our results are the first to suggest that the mechanisms leading to this delayed accumulation differ from that of normal accumulation and relies on local proliferation. One could argue that the preconditioning with clodronate has enriched the residual monocyte population for the highly proliferative monocytes. To verify it, we have assessed the effect of clodronate treatment on the percentage of proliferative monocytes into PBMCs population. We observed that clodronate treatment tended to enrich CD11b+CD68+Ki-67+ proliferative blood monocyte population by about 2.5-fold at all time points tested in comparison to untreated animals (data not shown). This increase represents a small absolute percentage of proliferative monocytes into PBMCs population (0.01% in non-treated vs. 0.025% in treated animals), and given the short timeframe of this study, those could have contributed partially the large increase of CD11b+CD68+Ki-67+ proliferative macrophages observed into muscle.
Our data strongly suggest that a precursor located within skeletal muscles prior to injury can contribute to M1 macrophage accumulation when monocyte infiltration is blocked. Irradiation, which prevents replication of resident macrophages, did not have any effect on the absolute number of M1 macrophages in non-depleted animals following injury, suggesting that infiltration of blood-derived monocytes was the main mechanism in those conditions. Inversely, irradiation short-circuited the increase of M1 macrophage number that was induced at 4 d with continuous depletion of monocytes. We conclude that when monocyte infiltration is artificially decreased or delayed, an alternative mechanism based on proliferation of a local M1 precursor is triggered to ensure the accumulation of that specific subset.
M2 macrophage accumulation
Based on the present results, we conclude that M2 macrophage accumulation following injury relies on mixed mechanisms involving infiltration of blood monocytes as well as proliferation of local and radiation-sensitive precursors. Different mechanisms have been proposed to explain M2 macrophage accumulation; these include a specific circulating M2 precursor (CX3CR1hi/Ly6Clo monocytes) , a switch of phenotype from M1 toward M2 , and lastly, the capacity of M2 macrophages to undergo local proliferation . Data obtained in models of muscle injury led us to originally hypothesize that depletion of blood monocyte would eliminate M1 macrophages and thus indirectly decrease the number of M2 macrophages . As predicted, CD163+ macrophage accumulation was significantly reduced at 2 d post-injury in monocyte-depleted animals, but this difference was lost at later time points. In an attempt to explain this phenomenon, we showed that irradiation had a significant impact on the absolute number of M2 macrophages in injured rats treated or not with clodronate. These results thus suggest that M2 macrophage accumulation is ensured, at least partially, by a local progenitor such as resident cells expressing M2 phenotype or resulting from the conversion of previously infiltrated M1 macrophages into M2 [7, 16]. Conversely, M2 macrophage accumulation was still significant when muscle was irradiated, which argues against the hypothesis that local radiation-sensitive precursors are the only source for M2 macrophages. Overall, the data obtained by combining monocyte depletion and irradiation suggest that different pools of infiltrating and local M2 precursors could differently contribute to their accumulation. In order to identify a circulating progenitor for M2 macrophages, we assessed if a circulating monocyte subset expressed the M2 marker CD163 but were unable to detect such cells. Thus, the possibility of a CD68+CD163- precursor or of an unknown circulating precursor for M2 macrophages remains open. Moreover, non-circulating progenitor cannot be excluded since it has been shown in brain that microglia are maintained throughout life independently of any blood input .
In summary, the present study shows for the first time that macrophages have the capacity to proliferate following sterile muscle injury and that the number of proliferating macrophages is increased within muscles when monocyte infiltration is blocked. We conclude that under normal physiological conditions, the main source of M1 macrophages is circulating monocytes while M2 macrophage accumulation relies on both local and infiltrating precursors. As a whole, our study highlights overlapping mechanisms involved in macrophage accumulation after sterile skeletal muscle injury and suggests that these mechanisms are modulated when monocyte infiltration is impaired like is the case of anti-inflammatory conditions.
We want to thank M. Marc Veillette for his intellectual contribution and technical support and Dr. Jérôme Frenette for his innovative ideas.
- Segawa M, Fukada S, Yamamoto Y, Yahagi H, Kanematsu M, Sato M, Ito T, Uezumi A, Hayashi S, Miyagoe-Suzuki Y: Suppression of macrophage functions impairs skeletal muscle regeneration with severe fibrosis. Exp Cell Res. 2008, 314 (17): 3232-3244. 10.1016/j.yexcr.2008.08.008.View ArticlePubMedGoogle Scholar
- Arnold L, Henry A, Poron F, Baba-Amer Y, van Rooijen N, Plonquet A, Gherardi RK, Chazaud B: Inflammatory monocytes recruited after skeletal muscle injury switch into antiinflammatory macrophages to support myogenesis. J Exp Med. 2007, 204 (5): 1057-1069. 10.1084/jem.20070075.View ArticlePubMedPubMed CentralGoogle Scholar
- Summan M, Warren GL, Mercer RR, Chapman R, Hulderman T, Van Rooijen N, Simeonova PP: Macrophages and skeletal muscle regeneration: a clodronate-containing liposome depletion study. Am J Physiol Regul Integr Comp Physiol. 2006, 290 (6): R1488-R1495. 10.1152/ajpregu.00465.2005.View ArticlePubMedGoogle Scholar
- Chazaud B, Brigitte M, Yacoub-Youssef H, Arnold L, Gherardi R, Sonnet C, Lafuste P, Chretien F: Dual and beneficial roles of macrophages during skeletal muscle regeneration. Exerc Sport Sci Rev. 2009, 37 (1): 18-22. 10.1097/JES.0b013e318190ebdb.View ArticlePubMedGoogle Scholar
- Tidball JG, Villalta SA: Regulatory interactions between muscle and the immune system during muscle regeneration. Am J Physiol Regul Integr Comp Physiol. 2010, 298 (5): R1173-R1187. 10.1152/ajpregu.00735.2009.View ArticlePubMedPubMed CentralGoogle Scholar
- Villalta SA, Rinaldi C, Deng B, Liu G, Fedor B, Tidball JG: Interleukin-10 reduces the pathology of mdx muscular dystrophy by deactivating M1 macrophages and modulating macrophage phenotype. Human molecular genetics. 2011, 20 (4): 790-805. 10.1093/hmg/ddq523.View ArticlePubMedGoogle Scholar
- Murray PJ, Wynn TA: Protective and pathogenic functions of macrophage subsets. Nat Rev Immunol. 2011, 11 (11): 723-737. 10.1038/nri3073.View ArticlePubMedPubMed CentralGoogle Scholar
- Lawrence T, Natoli G: Transcriptional regulation of macrophage polarization: enabling diversity with identity. Nat Rev Immunol. 2011, 11 (11): 750-761. 10.1038/nri3088.View ArticlePubMedGoogle Scholar
- Gordon S, Taylor PR: Monocyte and macrophage heterogeneity. Nat Rev Immunol. 2005, 5 (12): 953-964. 10.1038/nri1733.View ArticlePubMedGoogle Scholar
- Mosser DM, Edwards JP: Exploring the full spectrum of macrophage activation. Nat Rev Immunol. 2008, 8 (12): 958-969. 10.1038/nri2448.View ArticlePubMedPubMed CentralGoogle Scholar
- Martinez CO, McHale MJ, Wells JT, Ochoa O, Michalek JE, McManus LM, Shireman PK: Regulation of skeletal muscle regeneration by CCR2-activating chemokines is directly related to macrophage recruitment. Am J Physiol Regul Integr Comp Physiol. 2010, 299 (3): R832-R842. 10.1152/ajpregu.00797.2009.View ArticlePubMedPubMed CentralGoogle Scholar
- Sun D, Martinez CO, Ochoa O, Ruiz-Willhite L, Bonilla JR, Centonze VE, Waite LL, Michalek JE, McManus LM, Shireman PK: Bone marrow-derived cell regulation of skeletal muscle regeneration. FASEB J. 2009, 23 (2): 382-395.View ArticlePubMedPubMed CentralGoogle Scholar
- Tidball JG, Wehling-Henricks M: Macrophages promote muscle membrane repair and muscle fibre growth and regeneration during modified muscle loading in mice in vivo. The Journal of physiology. 2007, 578 (Pt 1): 327-336.View ArticlePubMedGoogle Scholar
- Geissmann F, Auffray C, Palframan R, Wirrig C, Ciocca A, Campisi L, Narni-Mancinelli E, Lauvau G: Blood monocytes: distinct subsets, how they relate to dendritic cells, and their possible roles in the regulation of T-cell responses. Immunology and cell biology. 2008, 86 (5): 398-408. 10.1038/icb.2008.19.View ArticlePubMedGoogle Scholar
- Jenkins SJ, Ruckerl D, Cook PC, Jones LH, Finkelman FD, van Rooijen N, MacDonald AS, Allen JE: Local macrophage proliferation, rather than recruitment from the blood, is a signature of TH2 inflammation. Science. 2011, 332 (6035): 1284-1288. 10.1126/science.1204351.View ArticlePubMedPubMed CentralGoogle Scholar
- Nahrendorf M, Swirski FK, Aikawa E, Stangenberg L, Wurdinger T, Figueiredo JL, Libby P, Weissleder R, Pittet MJ: The healing myocardium sequentially mobilizes two monocyte subsets with divergent and complementary functions. J Exp Med. 2007, 204 (12): 3037-3047. 10.1084/jem.20070885.View ArticlePubMedPubMed CentralGoogle Scholar
- van Rooijen N, van Kesteren-Hendrikx E: In vivo" depletion of macrophages by liposome-mediated "suicide. Methods in enzymology. 2003, 373: 3-16.View ArticlePubMedGoogle Scholar
- Hodgetts SI, Grounds MD: Irradiation of dystrophic host tissue prior to myoblast transfer therapy enhances initial (but not long-term) survival of donor myoblasts. J Cell Sci. 2003, 116 (Pt 20): 4131-4146.View ArticlePubMedGoogle Scholar
- Denekamp J, Rojas A: Cell kinetics and radiation pathology. Experientia. 1989, 45 (1): 33-41. 10.1007/BF01990450.View ArticlePubMedGoogle Scholar
- Robertson TA, Grounds MD, Papadimitriou JM: Elucidation of aspects of murine skeletal muscle regeneration using local and whole body irradiation. J Anat. 1992, 181 (Pt 2): 265-276.PubMedPubMed CentralGoogle Scholar
- Gross JG, Morgan JE: Muscle precursor cells injected into irradiated mdx mouse muscle persist after serial injury. Muscle Nerve. 1999, 22 (2): 174-185. 10.1002/(SICI)1097-4598(199902)22:2<174::AID-MUS5>3.0.CO;2-S.View ArticlePubMedGoogle Scholar
- Gulati AK: The effect of X-irradiation on skeletal muscle regeneration in the adult rat. J Neurol Sci. 1987, 78 (1): 111-120. 10.1016/0022-510X(87)90083-9.View ArticlePubMedGoogle Scholar
- Gross JG, Bou-Gharios G, Morgan JE: Potentiation of myoblast transplantation by host muscle irradiation is dependent on the rate of radiation delivery. Cell Tissue Res. 1999, 298 (2): 371-375. 10.1007/s004419900062.View ArticlePubMedGoogle Scholar
- Marsolais D, Hahm B, Walsh KB, Edelmann KH, McGavern D, Hatta Y, Kawaoka Y, Rosen H, Oldstone MB: A critical role for the sphingosine analog AAL-R in dampening the cytokine response during influenza virus infection. Proc Natl Acad Sci USA. 2009, 106 (5): 1560-1565. 10.1073/pnas.0812689106.View ArticlePubMedPubMed CentralGoogle Scholar
- Kim WK, Alvarez X, Fisher J, Bronfin B, Westmoreland S, McLaurin J, Williams K: CD163 identifies perivascular macrophages in normal and viral encephalitic brains and potential precursors to perivascular macrophages in blood. Am J Pathol. 2006, 168 (3): 822-834. 10.2353/ajpath.2006.050215.View ArticlePubMedPubMed CentralGoogle Scholar
- Tippett E, Cheng WJ, Westhorpe C, Cameron PU, Brew BJ, Lewin SR, Jaworowski A, Crowe SM: Differential expression of CD163 on monocyte subsets in healthy and HIV-1 infected individuals. PloS one. 2011, 6 (5): e19968-10.1371/journal.pone.0019968.View ArticlePubMedPubMed CentralGoogle Scholar
- Polfliet MM, Fabriek BO, Daniels WP, Dijkstra CD, van den Berg TK: The rat macrophage scavenger receptor CD163: expression, regulation and role in inflammatory mediator production. Immunobiology. 2006, 211 (6–8): 419-425.View ArticlePubMedGoogle Scholar
- Mills CD, Kincaid K, Alt JM, Heilman MJ, Hill AM: M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol. 2000, 164 (12): 6166-6173.View ArticlePubMedGoogle Scholar
- Mantovani A, Sica A, Sozzani S, Allavena P, Vecchi A, Locati M: The chemokine system in diverse forms of macrophage activation and polarization. Trends Immunol. 2004, 25 (12): 677-686. 10.1016/j.it.2004.09.015.View ArticlePubMedGoogle Scholar
- Moyer AL, Wagner KR: Regeneration versus fibrosis in skeletal muscle. Current opinion in rheumatology. 2011, 23 (6): 568-573. 10.1097/BOR.0b013e32834bac92.View ArticlePubMedGoogle Scholar
- Bot A, Smith KA, von Herrath M: Molecular and cellular control of T1/T2 immunity at the interface between antimicrobial defense and immune pathology. DNA and cell biology. 2004, 23 (6): 341-350. 10.1089/104454904323145227.View ArticlePubMedGoogle Scholar
- Khallou-Laschet J, Varthaman A, Fornasa G, Compain C, Gaston AT, Clement M, Dussiot M, Levillain O, Graff-Dubois S, Nicoletti A: Macrophage plasticity in experimental atherosclerosis. PloS one. 2010, 5 (1): e8852-10.1371/journal.pone.0008852.View ArticlePubMedPubMed CentralGoogle Scholar
- Mikita J, Dubourdieu-Cassagno N, Deloire MS, Vekris A, Biran M, Raffard G, Brochet B, Canron MH, Franconi JM, Boiziau C: Altered M1/M2 activation patterns of monocytes in severe relapsing experimental rat model of multiple sclerosis. Amelioration of clinical status by M2 activated monocyte administration. Mult Scler. 2011, 17 (1): 2-15. 10.1177/1352458510379243.View ArticlePubMedGoogle Scholar
- Hashimoto D, Chow A, Noizat C, Teo P, Beasley MB, Leboeuf M, Becker CD, See P, Price J, Lucas D: Tissue-resident macrophages self-maintain locally throughout adult life with minimal contribution from circulating monocytes. Immunity. 2013, 38 (4): 792-804. 10.1016/j.immuni.2013.04.004.View ArticlePubMedGoogle Scholar
- Ajami B, Bennett JL, Krieger C, Tetzlaff W, Rossi FM: Local self-renewal can sustain CNS microglia maintenance and function throughout adult life. Nature neuroscience. 2007, 10 (12): 1538-1543. 10.1038/nn2014.View ArticlePubMedGoogle Scholar
- Schulz C, Gomez Perdiguero E, Chorro L, Szabo-Rogers H, Cagnard N, Kierdorf K, Prinz M, Wu B, Jacobsen SE, Pollard JW: A lineage of myeloid cells independent of Myb and hematopoietic stem cells. Science. 2012, 336 (6077): 86-90. 10.1126/science.1219179.View ArticlePubMedGoogle Scholar
- Dobbertin A, Schmid P, Gelman M, Glowinski J, Mallat M: Neurons promote macrophage proliferation by producing transforming growth factor-beta2. The Journal of neuroscience: the official journal of the Society for Neuroscience. 1997, 17 (14): 5305-5315.Google Scholar
- Senokuchi T, Matsumura T, Sakai M, Yano M, Taguchi T, Matsuo T, Sonoda K, Kukidome D, Imoto K, Nishikawa T: Statins suppress oxidized low density lipoprotein-induced macrophage proliferation by inactivation of the small G protein-p38 MAPK pathway. J Biol Chem. 2005, 280 (8): 6627-6633. 10.1074/jbc.M412531200.View ArticlePubMedGoogle Scholar
- Celada A, Borras FE, Soler C, Lloberas J, Klemsz M, van Beveren C, McKercher S, Maki RA: The transcription factor PU.1 is involved in macrophage proliferation. J Exp Med. 1996, 184 (1): 61-69. 10.1084/jem.184.1.61.View ArticlePubMedGoogle Scholar
- Moeslinger T, Spieckermann PG: Urea-induced inducible nitric oxide synthase inhibition and macrophage proliferation. Kidney international Supplement. 2001, 78: S2-S8.View ArticlePubMedGoogle Scholar
- Spector WG, Wynne KM: Proliferation of macrophages in inflammation. Agents and actions. 1976, 6 (1–3): 123-126.View ArticlePubMedGoogle Scholar
- Khmelewski E, Becker A, Meinertz T, Ito WD: Tissue resident cells play a dominant role in arteriogenesis and concomitant macrophage accumulation. Circulation research. 2004, 95 (6): E56-E64. 10.1161/01.RES.0000143013.04985.E7.View ArticlePubMedGoogle Scholar
- Bryer SC, Fantuzzi G, Van Rooijen N, Koh TJ: Urokinase-type plasminogen activator plays essential roles in macrophage chemotaxis and skeletal muscle regeneration. J Immunol. 2008, 180 (2): 1179-1188.View ArticlePubMedGoogle Scholar
- DiPasquale DM, Cheng M, Billich W, Huang SA, van Rooijen N, Hornberger TA, Koh TJ: Urokinase-type plasminogen activator and macrophages are required for skeletal muscle hypertrophy in mice. Am J Physiol Cell Physiol. 2007, 293 (4): C1278-C1285. 10.1152/ajpcell.00201.2007.View ArticlePubMedGoogle Scholar
- Ginhoux F, Greter M, Leboeuf M, Nandi S, See P, Gokhan S, Mehler MF, Conway SJ, Ng LG, Stanley ER: Fate mapping analysis reveals that adult microglia derive from primitive macrophages. Science. 2010, 330 (6005): 841-845. 10.1126/science.1194637.View ArticlePubMedPubMed CentralGoogle Scholar
- The pre-publication history for this paper can be accessed here:http://www.biomedcentral.com/1471-2474/14/359/prepub
This article is published under license to BioMed Central Ltd. This is an open access article distributed under the terms of the Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.