Identification of polarized macrophage subsets in zebrafish

1Behind the generic name ‘macrophage’ hides various cell types with distinct phenotypes and functions. Currently, it is well established that macrophages are not just important immune effector cells but also cells with critical homeostatic roles, exerting a myriad of functions in development, homeostasis, and tissue repair and playing a pivotal role in disease progression. Therefore, there is a high interest in a better characterization of these cells to establish an early and accurate diagnosis. The wide variety of macrophage functions might be explained by the outstanding plasticity and versatility of macrophages that efficiently respond to environmental challenges and changes in tissue physiology by modifying their phenotype. Although there is a consensus that macrophages are a diversified set of cells, macrophage subtypes are still poorly characterized. Indeed, although these cell populations have been extensively investigated in mouse and human, these studies were mostly performed in vitro using monocyte-derived macrophages induced under specific stimuli. A comprehensive characterization of macrophage subsets that takes into account their specific behaviour, phenotypic diversity, functions, and modulation shall rely on a real-time tracking in the whole organism in response to environmental challenges.

2Mouse and human macrophages have been classified according to their polarization state. In this classification, M1 macrophages, also referred as classically activated macrophages, are pro-inflammatory cells associated with the first phases of inflammation, while M2 macrophages, also known as alternatively activated macrophages, are involved in the resolution of inflammation and tissue remodelling. Differential cytokine and chemokine production and receptor expression define the polarization state of macrophages. However, it is worthwhile to note that such binary naming does not fully reflect the diversity of macrophage phenotypes in complex in vivo environments in which several cytokines and growth factors are released and adjust the final differentiated state. Macrophages might adopt intermediate activation phenotypes classified by the relative levels of macrophage subset-specific markers. Therefore, macrophage plasticity results in a full spectrum of macrophage subsets with a myriad of functions. Although the possible phenotype conversion of macrophages from M1 to M2 has been suggested in in vitro studies, a recent study argues for the sequential homing of M1 and M2 macrophages to the site of injury. Such controversies highlight the lack of accurate real-time tracing of macrophage subtypes in vivo in the entire animal.

3Inflammation is a model of choice to study the wide range of macrophage subsets involved from its initiation to its resolution. Therefore, in the present study, we propose to decipher in vivo in real time the kinetic of macrophage subset recruitment, their behaviour and their phenotypic plasticity at the molecular level during a multiple-step inflammatory process. We used the zebrafish larvae model for its easy genetic manipulation, transparency, and availability of fluorescent reporter lines to track macrophages. While the existence of macrophage subtypes in zebrafish embryos has been suggested, they have not been fully characterized. Here, we report a new reporter transgene for TNFa, a central inflammatory cytokine and well-established marker of M1 macrophages, instrumental to discriminate macrophage subsets during intravital imaging.

4Fin wounding-induced inflammation and Escherichia coli inoculation in zebrafish larvae of 3 dpf are two well-established models triggering macrophage recruitment. Using in situ hybridization, we observed that the expression of the tumour necrosis factor alpha (tnfa), a consensus marker of M1 macrophages, was induced in cells accumulated in the caudal fin and the muscle following amputation (nlarvae = 29/33) and E. coli inoculation (nlarvae = 12/12), respectively (Figure 1A–E). To study the cells that express the tnfa transcripts, we established the Tg(tnfa:eGFP-F) transgenic zebrafish line expressing a farnesylated (membrane-bound) eGFP (eGFP-F) under the control of the tnfa promoter. While eGFP-F was undetectable in intact fins of Tg(tnfa:eGFP-F) larvae (nlarvae = 10/10), it was expressed in cells recruited to the wound at 6 hr post-amputation (hpA) (nlarvae = 16/16, Figure 1F,G). Similarly, eGFP-F expression was upregulated in cells accumulated in the muscle of Tg(tnfa:eGFP-F) larvae at 16 hr post-inoculation (hpi) of DsRed-expressing E. coli (nlarvae = 8/8), compared to Phosphate-buffered saline (PBS) injection (nlarvae = 3/3, Figure 1H,I). Confocal analysis confirmed the presence of a membrane-bound eGFP in cells displaying a typical myeloid morphology (Figure 1J). To demonstrate that the Tg(tnfa:eGFP-F) line recapitulates transcriptional activation of tnfa, we performed a simultaneous detection of tnfa mRNA by in situ hybridization and GFP-F protein by immunofluorescence in amputated larvae 6 hpA. We observed a consistent overlap between tnfa and GFP-F signal in the fin (nlarvae = 11/11), showing the direct correlation of eGFP-F and tnfa transcriptional activation in the fin of the reporter line (Figure 1K). In addition, we FACS-sorted GFP+ cells from wounded Tg(TNFa:eGFP-F) larvae 6 hpA and performed RT-qPCR to analyze tnfa expression. We observed a significant increase of tnfa mRNA level in eGFP+ cells as compared to eGFP− cells (Figure 1L,M). All together these results indicate that the Tg(TNFa:eGFP-F) reporter line recapitulates transcriptional activation of tnfa. Then, with the ability to specifically track tnfa-expressing cells, we used Tg(tnfa:eGFP-F) fish to study macrophage activation by mating them with Tg(mpeg1:mCherryF) fish in which macrophages express farnesylated mCherry (mCherryF) under the control of the macrophage-specific mpeg1 promoter. In intact Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae, no eGFP-F was observed in macrophages (Figure 2A). We imaged double transgenic larvae Tg(tnfa:eGFP-F/mpeg1:mCherryF) using 4D confocal microscopy from 45 min post-amputation and found that macrophages were recruited to the wound from 1 hpA, some starting to express eGFP from 3 hpA (Figure 2B,C and Video 1). From 5 hpA already activated macrophages, that is, expressing tnfa arrived at the wound (Video 1 and Figure 2C). Similarly, infection with a crimson-expressing E. coli in the muscle induced the expression of tnfa in phagocytes few hours following the infection (Figure 2—figure supplement 1, Video 2). Imaging of the double transgenic larvae Tg(tnfa:eGFP-F/mpeg1:mCherryF) showed that tnfa-expressing phagocytes were mainly macrophages (Figure 2—figure supplement 1, Video 2). These results show the dynamic macrophage activation in real-time in vivo including recruitment and rapid phenotypic change. During the revision of this paper, a similar result has been published.

5To test whether tnfa+ and tnfa− macrophages harboured different cellular characteristics, we first analyzed their morphology in fin-wounded Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae. tnfa+mpeg1+ cells displayed flattened and lobulated morphology (Figure 2D), while tnfa−mpeg1+ were elongated and dendritic (Figure 2E). As we observed that tnfa+mpeg1+ cells were predominant at the wound at 18 hpA and tnfa−mpeg1+ cells at 24 hpA (data not shown), we imaged the behaviour of these macrophage populations in wounded fins from Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae at these time points. tnfa+mpeg1+ cells presented a lower velocity (0.32 μm/min) than tnfa−mpeg1+ macrophages (1.09 μm/min, Figure 2F) but a higher cell–cell contact frequency (0.036 VS 0.016 contacts/min) with other macrophages (Figure 2G,I,J and Videos 3, 4). Measurements of the duration of macrophage–macrophages contacts showed that these contacts lasted longer (48.6 min/contact) than that of tnfa−mpeg1+ macrophages (15.9 min/contact, Figure 2H–J and Videos 3, 4). All together these data highlight different morphology and behaviour of macrophage phenotypes in live zebrafish suggesting the existence of macrophage subsets exhibiting different functions.

6To quantify the respective frequency of tnfa+ macrophages (mCherry+eGFP+ referred as dbl+) and tnfa− macrophages (mCherry+eGFP− referred as mCh+), we performed flow cytometry analysis on cells isolated from Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae at different time points following caudal fin amputation or E. coli. inoculation (Figure 3A,B). While only 5.6% ± 0.9 (s.e.m.) dbl+ cells were detected in the mpeg1+ population of the intact larvae, a steady increase of the dbl+ population from 6 to 20 hpA (up to 27.33 ± 0.2%) was observed. This percentage decreased dramatically at 26 hpA to 8.75 ± 1%. In E. coli inoculation experiments, the frequency of dbl+ cells increased as soon as 3 hpi (55.60 ± 0.6%) and remained stable until 26 hpi. These results demonstrate that wound-induced macrophage activation is transient compared to infection-induced macrophage activation.

7To characterize at the molecular level tnfa− and tnfa+ macrophage populations during early and late phases of inflammation, we FACS-sorted dbl+ and mCh+ cells from Tg(tnfa:eGFP-F/mpeg1:mCherryF) tail-amputated larvae (Figure 3C) and analyzed them by qRT-PCR. In mammals, M1 and M2 macrophages are reported to be involved, respectively, in the initial phase of inflammation and in the resolution phase. Cell sorting was thus performed at 6 hpA and 26 hpA following caudal fin amputation since the kinetic analysis of macrophage subset activation (Figure 3B) suggested that these two time points correspond to initiation and resolution of inflammation, respectively. As expected, high levels of mpeg1 expression was observed in the mCh+ and dbl+ sorted cells at 6 and 26 hpA (Figure 3D), and high levels of tnfa expression was detected in double-positive populations at 6 hpA (Figure 3E). These observations demonstrated that fluorescence of these transgenes can be efficiently used to track and separate macrophage sub-populations. At 6 hpA, dbl+ macrophages expressed high levels of tnfb, il1b, and il6 compared to mCh+ macrophages (Figure 3F,G) that are well-known markers of M1 macrophages in mammals. By contrast, mCh+ macrophages expressed low levels of these pro-inflammatory cytokines at both 6 and 26 hpA (Figure 3F,G), but expressed high levels of tgfb1, ccr2, and cxcr4b (Figure 3H), that are specifically expressed in mammalian M2 macrophages. Of note, neither Arginase 1 (Arg1), which is largely used as a M2 marker in mouse but not in human, nor il10 (data not shown), a known M2 marker in mammals, was detected in zebrafish macrophages. Importantly, based on the stability of the eGFP in Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae allowing us to specifically track the behaviour and fate of pro-inflammatory macrophages, we found that the dbl+ pro-inflammatory macrophages changed their phenotype at 26 hpA. Indeed, dbl+ macrophages negative for M2 markers at 6 hpA, displayed at 26 hpA, in parallel to a significant decrease of tnfa, il1b, and il6 expression level, a significant increased expression level of ccr2 and cxcr4b (Figure 3E,H). Of note, a tendency toward differential expression level was observed for tnfb and tgfb1 between 6 and 26 hpA. To go further and demonstrate that the same macrophages are present at the wound site during inflammation and its resolution, we generated the Tg(mpeg1:GAL4/UAS:Kaede) larvae to track macrophages exploiting the conversion of the native green fluorescence of Kaede into red fluorescence under UV light. Recruited macrophages were photoconverted 6 hpA and imaged at 26 hpA revealing that early recruited macrophages were still present at the wound area 20 hr later (Figure 4—figure supplement 1). Then, GFP+ macrophages were specifically tracked using time-lapse imaging of wounded Tg(mpeg1:mCherryF/TNFa:GFP-F) fins from 6 to 26 hpA. We show that initially recruited eGFP+ macrophages remain at the injury site and still express the GFP (Figure 4A–C and Video 5). The analysis of macrophage behaviour over time shows that among eGFP+ macrophages displaying an amyboid phenotype at the wound edge 6 hpA, 50% change toward a fibroblastic phenotype from 11 hpA when they moved distally (Video 5). All together these data show that pro-inflammatory macrophages underwent a phenotypic conversion toward an intermediate phenotype in which both M1 and M2 markers are expressed. In addition, this molecular characterization of macrophages in zebrafish reveals the conservation of macrophage subtypes between zebrafish and human.

8In conclusion, we identified macrophage subsets in zebrafish and described their behaviour and fate during a process of inflammation (Figure 4D). Live imaging of transparent transgenic zebrafish larvae allowed the first real-time visualization of macrophage activation and polarization. In parallel, a molecular analysis of macrophage sub-populations highlights the evolutionary conservation of macrophages from fish to mammals. We propose that in response to wounding zebrafish, unpolarized macrophages are recruited to the inflammation site and adopt a M1-like phenotype. Subsequently, they progressively convert their functional phenotype from M1-like to M2-like in response to progressive inflammatory microenvironment changes within the tissue (Figure 4D). Live imaging of the new transgenic line we generated opens new avenues to study in real time in live vertebrates the full spectrum of macrophage activation, polarization, and functions.

9All animal experiments described in the present study were conducted at the University of Montpellier according to European Union guidelines for handling of laboratory animals (http://ec.europa.eu/environment/chemicals/lab_animals/home_en.htm) and were approved by the Direction Sanitaire et Vétérinaire de l'Hérault and Comité d'Ethique pour l'Expérimentation Animale under reference CEEA-LR-13007.

10Fish and embryo maintenance, staging, and husbandry were as previously described. Experiments were performed using the AB zebrafish strain (ZIRC) and the transgenic line Tg(mpeg1:mCherryF) to visualize macrophages. For the photoconversion experiments, a cross of Tg(mpeg1:Gal4)gl25 and Tg(UAS:kaede)rk8 lines was used, using breeders selected for progeny with negligible silencing of the UAS transgene.

11The TNFa promoter (Gene ID: 405785) was amplified from zebrafish genomic DNA using primers zTNFaP4 (CCCGCATGCTCCACGTCTCC) and zTNFaE11N (TTATAGCGGCCGCCCGACTCTCAAGCTTCA). The resulting fragment was phosphorylated using T4PNK, digested by NotI and cloned in a farnesylated eGFP (eGFP-F) derivative of pBSKI2. The resulting plasmid (pI2promTNFa:eGFP-F) harbours a 3.8-kb fragment of the zebrafish tnfa promoter, including part of the first coding exon. It uses the endogenous ATG codon of tnfa to drive the translation of eGFP-F. The expressed eGFP-F harbours the first 7 amino acids of zebrafish TNFa at its N-terminus (MKLESRA). The expression cassette is flanked by two I-SceI sites. pI2promTNFa:eGFP-F was co-injected in fertilized eggs with the enzyme I-SceI (New England Biolabs, France). Developing embryos were injected with non-pathogenic E. coli at 3 dpf (days post-fertilization), and those that developed a specific green fluorescence were raised as putative founders. The offspring of putative founders was tested the same way in order to establish the stable transgenic line. Low expression of eGFP in the pharynx at 3 dpf was used to check larvae for the presence of the transgene.

12Caudal fin amputation was performed on 3 dpf larvae as described in. The caudal fin was transected with a sterile scalpel, posterior to muscle and notochord under anaesthesia with 0.016% Tricaine (ethyl 3-aminobenzoate, Sigma Aldrich, France) in zebrafish water. Larvae were inoculated at 3 dpf by 2.103 CFU E. coli K12 bacteria harbouring either DsRed or Crimson (Clontech, France) expression plasmid or no plasmid. Imaging was performed as previously described using a confocal TCS SP5 inverted microscope with a HCXPL APO 40×/1.25–0.75 oil objective (Leica, France). Image stacks for time-lapse videos were acquired every 3–5 min, typically spanning 30–60 μm at 2-μm intervals, 1024 × 512 or 512 × 512 pixel resolution. The 4D files generated from time-lapse acquisitions were processed using Image J. They were compressed into maximum intensity projections and cropped. Brightness, contrast, and colour levels were adjusted for maximal visibility. Velocity of macrophages was measured using Manual Tracking Image J plugin. Frequency of macrophage–macrophage interaction and duration of interactions were measured manually on stack images. For tracking of macrophages, eGFP-F+ mCherryF+ cells from Tg(mpeg1:mCherryF/tnfa:eGFP-F) wounded fins were tracked using time-lapse image series from 6 hpA (hours post-amputation) to 26 hpA.

13300 Tg(tnfa:eGFP-F/mpeg1:mCherryF) larvae were either amputated or infected as described above, then crushed on a 70-µm cell strainer (Falcon, France). Isolated cells were washed in PBS/2 mM ethylenediaminetetraacetic acid (EDTA)/2% Foetal Calf Serum (FCS), filtered through a 40-µm cell strainer, and counted. Counting of mCherry+eGFP− and mCherry+eGFP+ cells was performed on LSRFortessa (BD Bioscience, France), and data analyzed using the Flowjo software (Tree start, Ashland, Or, USA). Sorting was done using FACS ARIA (BD Bioscience, France) and collected in 50% FCS/50% Leibovitz L-15 medium (21083-027, Gibco, France) on ice. To isolate total RNA, cells were lysed in QIAzol Lysis Reagent (Qiagen, France) and RNA extracted using miRNeasy mini kit (Qiagen-21704, France). 20 ng of total RNA was reverse transcribed using High Capacity RNA Reverse Transcription kit (Applied Biosystems, France). QPCR were performed on a LightCycler 480 system (Roche, France), following manufacturer's instructions (SYBR Green format, Roche Applied Science, Meylan, France) and using primers in Supplementary file 1: denaturation 15 s at 95°C, annealing 10 s at 64°C, and elongation 20 s at 72°C. Expression levels were determined with the LightCycler analysis software (version-3.5) from 5 independent experiments. The relative amount of a given mRNA was calculated by using the formulae 2−∆Ct with ef1a as reference.

14Significance testing for Figures 1M, 3D–H was done using Mann–Whitney unpaired t-test, one-tail and Figure 2F–H using Mann–Whitney unpaired t-test, two-tails using GraphPad Prism 6 Software. *p < 0.05, **p < 0.01, ****p < 0.0001.

15A tnfa probe was amplified from total cDNA by PCR using tnfa.55 and tnfa.58 primers (Supplementary file 1) and cloned in plasmid pCRII-TOPO. Digoxigenin (DIG)-labelled (Roche, France) sense and anti-sense RNA probes were in vitro transcribed (Biolabs, France). In situ hybridizations on whole-mount embryos were as previously described. For simultaneous detection of eGFP-F proteins and tnfa mRNA by immuno-detection and in situ hybridization, fixed and rehydrated Tg(tnfa:eGFP-F) larvae were permeabilised in ice in 100% ethanol for 5 min, then in a mixture of 50% Xylene-50% ethanol for 1 hr and in 80% acetone for 10 min at −20°C as described in. After washes in PBS-0.1% Tween, larvae were post-fixed in 4% paraformaldehyde (PFA) for 20 min. Subsequent steps of hybridization, washes, and staining with NBT-BCIP (Roche, France) were as previously described in. Next, unspecific-binding sites were saturated in PBS-1% bovin serum albumin (BSA)-1% lamb serum-10% Goat serum and larvae incubated 3 days with an anti-GFP antibody (MBL, 1/500). After extensive washes, larvae were incubated with a goat anti-rabbit antibody. Stained embryos were imaged using a MVX10 Olympus microscope with MVPLAPO 1× objective and XC50 camera and using a Zeiss Axioimager with a Zeiss 40× Plan-Apo 1.3 oil objective.

16Tg(mpeg1:GAL4/UAS:Kaede) embryos were raised to 3 dpf in the dark, and caudal fin was transected as described above. At 6 hpA, larvae were mounted in 1% low-melting point agarose. A 405-nm Laser Cube 405-50C on a confocal TCS SP5 inverted microscope with a HCXPL APO 40×/1.25–0.75 oil objective (Leica) was used to photoconvert the Kaede-labelled cells using 6% laser power scanning for 60 s (optimized before the experiments; data not shown). Fins were imaged before and after the photoconversion (at 6 and 26 hpA) in the green and red channels.