Immune–tracheal intercellular signalling coordinates the muscle injury response in Drosophila

Immune–Vascular intercellular signalling coordinates tissue plasticity during adult muscle repair in Drosophila | bioRxiv /* */ /* */ <!-- <!-- /*! * yepnope1.5.4 * (c) WTFPL, GPLv2 */ (function(a,b,c){function d(a){return"[object Function]"==o.call(a)}function e(a){return"string"==typeof a}function f(){}function g(a){return!a||"loaded"==a||"complete"==a||"uninitialized"==a}function h(){var a=p.shift();q=1,a?a.t?m(function(){("c"==a.t?B.injectCss:B.injectJs)(a.s,0,a.a,a.x,a.e,1)},0):(a(),h()):q=0}function i(a,c,d,e,f,i,j){function k(b){if(!o&&g(l.readyState)&&(u.r=o=1,!q&&h(),l.onload=l.onreadystatechange=null,b)){"img"!=a&&m(function(){t.removeChild(l)},50);for(var d in y[c])y[c].hasOwnProperty(d)&&y[c][d].onload()}}var j=j||B.errorTimeout,l=b.createElement(a),o=0,r=0,u={t:d,s:c,e:f,a:i,x:j};1===y[c]&&(r=1,y[c]=[]),"object"==a?l.data=c:(l.src=c,l.type=a),l.width=l.height="0",l.onerror=l.onload=l.onreadystatechange=function(){k.call(this,r)},p.splice(e,0,u),"img"!=a&&(r||2===y[c]?(t.insertBefore(l,s?null:n),m(k,j)):y[c].push(l))}function j(a,b,c,d,f){return q=0,b=b||"j",e(a)?i("c"==b?v:u,a,b,this.i++,c,d,f):(p.splice(this.i++,0,a),1==p.length&&h()),this}function k(){var a=B;return a.loader={load:j,i:0},a}var l=b.documentElement,m=a.setTimeout,n=b.getElementsByTagName("script")[0],o={}.toString,p=[],q=0,r="MozAppearance"in l.style,s=r&&!!b.createRange().compareNode,t=s?l:n.parentNode,l=a.opera&&"[object Opera]"==o.call(a.opera),l=!!b.attachEvent&&!l,u=r?"object":l?"script":"img",v=l?"script":u,w=Array.isArray||function(a){return"[object Array]"==o.call(a)},x=[],y={},z={timeout:function(a,b){return b.length&&(a.timeout=b[0]),a}},A,B;B=function(a){function b(a){var a=a.split("!"),b=x.length,c=a.pop(),d=a.length,c={url:c,origUrl:c,prefixes:a},e,f,g;for(f=0;f<d;f++)g=a[f].split("="),(e=z[g.shift()])&&(c=e(c,g));for(f=0;f<b;f++)c=x[f](c);return c}function g(a,e,f,g,h){var i=b(a),j=i.autoCallback;i.url.split(".").pop().split("?").shift(),i.bypass||(e&&(e=d(e)?e:e[a]||e[g]||e[a.split("/").pop().split("?")[0]]),i.instead?i.instead(a,e,f,g,h):(y[i.url]?i.noexec=!0:y[i.url]=1,f.load(i.url,i.forceCSS||!i.forceJS&&"css"==i.url.split(".").pop().split("?").shift()?"c":c,i.noexec,i.attrs,i.timeout),(d(e)||d(j))&&f.load(function(){k(),e&&e(i.origUrl,h,g),j&&j(i.origUrl,h,g),y[i.url]=2})))}function h(a,b){function c(a,c){if(a){if(e(a))c||(j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}),g(a,j,b,0,h);else if(Object(a)===a)for(n in m=function(){var b=0,c;for(c in a)a.hasOwnProperty(c)&&b++;return b}(),a)a.hasOwnProperty(n)&&(!c&&!--m&&(d(j)?j=function(){var a=[].slice.call(arguments);k.apply(this,a),l()}:j[n]=function(a){return function(){var b=[].slice.call(arguments);a&&a.apply(this,b),l()}}(k[n])),g(a[n],j,b,n,h))}else!c&&l()}var h=!!a.test,i=a.load||a.both,j=a.callback||f,k=j,l=a.complete||f,m,n;c(h?a.yep:a.nope,!!i),i&&c(i)}var i,j,l=this.yepnope.loader;if(e(a))g(a,0,l,0);else if(w(a))for(i=0;i (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0];var j=d.createElement(s);var dl=l!='dataLayer'?'&l='+l:'';j.src='//www.googletagmanager.com/gtm.js?id='+i+dl;j.type='text/javascript';j.async=true;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-M677548'); Skip to main content Home About Submit ALERTS / RSS Search for this keyword Advanced Search New Results Immune–Vascular intercellular signalling coordinates tissue plasticity during adult muscle repair in Drosophila Nourhene Ammar , Olivier Josse , Aurélien Guillou , Jessica Perochon , Hadi Boukhatmi doi: https://doi.org/10.1101/2025.06.25.661466 Nourhene Ammar 1 Institut de Génétique et Développement de Rennes (IGDR), Université de Rennes , CNRS UMR6290, Rennes, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site Olivier Josse 1 Institut de Génétique et Développement de Rennes (IGDR), Université de Rennes , CNRS UMR6290, Rennes, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site Aurélien Guillou 1 Institut de Génétique et Développement de Rennes (IGDR), Université de Rennes , CNRS UMR6290, Rennes, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site Jessica Perochon 2 Institute of Cancer Sciences−University of Glasgow, Wolfson Wohl Cancer Research Centre , Glasgow, UK Find this author on Google Scholar Find this author on PubMed Search for this author on this site Hadi Boukhatmi 1 Institut de Génétique et Développement de Rennes (IGDR), Université de Rennes , CNRS UMR6290, Rennes, France Find this author on Google Scholar Find this author on PubMed Search for this author on this site For correspondence: hadi.boukhatmi{at}univ-rennes.fr Abstract Full Text Info/History Metrics Preview PDF Summary In skeletal muscle, immune and vascular responses are essential for regeneration, yet the cellular and molecular mechanisms that coordinate their activities to restore tissue function remain unclear. Here, using Drosophila , we uncover a multi-organ signaling program that integrates muscle fibers, macrophages, vascular-like tracheal cells, and the extracellular matrix (ECM) to drive adult skeletal muscle repair. Muscle injury induces rapid macrophage recruitment and secretion of the FGF-like ligand Branchless (Bnl), which activates FGF/FGFR signalling in tracheal cells and promotes their targeted expansion toward damaged muscle. Concomitantly, macrophages deposit ECM components on the damaged muscle, including Collagen IV, forming a localized microenvironment that restricts Bnl ligand diffusion and facilitates directed tracheal remodelling. Genetic disruption of macrophage-derived Bnl or macrophage-mediated ECM deposition abolishes tracheal remodeling and compromises muscle recovery. Together, these findings reveal a previously unrecognized immune muscle vascular program in which macrophages coordinate growth factor signalling and ECM organization to shape muscle microenvironment and preserve muscle function following acute damage. Introduction Effective tissue repair relies on precise communication and coordination across multiple cell types and tissue systems [ 1 ]. Skeletal muscle provides a prime example of this interdependent cellular network, as successful regeneration depends on tightly orchestrated interactions among diverse cell populations, including macrophages, muscle stem cells (MuSCs; also known as satellite cells), and the fibro/adipogenic progenitors (FAPs) [ 2 , 3 ]. Following injury, infiltrating macrophages promote muscle regeneration by signalling to MuSCs and regulating their activation, proliferation, and differentiation [ 4 , 5 ]. In parallel, extensive remodelling of both the vasculature and the extracellular matrix (ECM) is required to meet increased metabolic demands and provide structural support [ 6 , 7 ]. Immune, vascular, and ECM-mediated responses are therefore critical to muscle regeneration; however, how these systems are mechanistically integrated in vivo to drive effective repair remains poorly understood. Addressing this question has been limited by the lack of experimental systems that enable to track and manipulate multiple interacting cell types in vivo during muscle repair. Drosophila has emerged as a powerful in vivo model for dissecting multi-organ communication during regeneration, owing to its capacity to combine precise genetic manipulation with high-resolution imaging [ 8 ]. As in vertebrates, adult Drosophila indirect flight muscles (IFMs) harbor MuSCs that support muscle maintenance and have been implicated in regenerative responses following injury [ 9 – 11 ]. The IFMs muscles are organized into six stereotypically arranged bilateral pairs within the adult thorax [ 12 ]. Studies, including our own, have shown that macrophage-like immune cells, termed hemocytes, populate the IFMs under homeostatic conditions [ 9 , 13 ]. In addition to hemocytes, the IFMs are infiltrated by a sophisticated network of air-filled tracheal cells, which function analogously to the vertebrate vascular and respiratory systems by ensuring tissue oxygenation [ 14 , 15 ]. While tracheal branching has been extensively characterized at the morphological, cellular, and genetic levels during development [ 16 , 17 ], recent work has revealed that adult tracheal networks retain a remarkable capacity for dynamic remodelling in response to tissue damage, including during regeneration of the adult gut [ 18 – 20 ]. Finally, IFMs are ensheathed by a specialized basement membrane enriched in conserved ECM components, which provide structural integrity and mechanical support to the contractile apparatus [ 15 , 21 ]. These features provide an in vivo context in which interactions between muscle fibres, immune cells, vascular-like trachea, and the ECM can be explored during muscle repair. We show that muscle damage triggers rapid and sustained hemocyte recruitment, which is required for targeted tracheal remodelling through FGF/FGFR signalling, enabling selective recognition and support of damaged fibers. In parallel, hemocytes deposit ECM components around injured fibres, which spatially confine FGF-like ligand and promote tracheal recruitment. Disrupting tracheal remodelling or hemocyte-mediated ECM delivery compromises muscle performance after injury, revealing an immune–muscle–vascular signalling module that couples metabolic support to ECM organization to preserve muscle function. Results Hemocytes are recruited to the injured adult flight muscle Accurate analysis of muscle injury responses requires a robust molecular readout that distinguishes damaged muscle fibers from adjacent intact muscles. Although Drosophila is one model for studying muscle repair, it lacks molecular markers that specifically identify injured muscle fibers. In vertebrates, re-expression of embryonic Myosin Heavy Chain (MyHC-emb) in damaged fibers is a common marker of muscle injury and regeneration [ 22 ]. To assess whether Myosin Heavy Chain (Mhc) expression could similarly serve as an indicator of muscle damage in Drosophila , we analysed its expression following injury. Our results show that Mhc expression is drastically upregulated in injured muscles ( Figure 1 A-F ) and inversely correlated with the downregulation of Mef2, providing a robust molecular readout of muscle damage ( Figure S1 A-B’’ ). This upregulation persists for at least 20 days post-injury (DPI) ( Figure S2 C-L ). In Drosophila , several Mhc isoforms are generated via alternative splicing of a single Mhc gene [ 23 ]. To determine which isoform respond to injury, we analysed the expression of various GFP-tagged Mhc isoforms [ 23 , 24 ]. Only the shorter adult Mhc isoforms-K, L, and M-were upregulated in response to muscle damage ( Figure S1 C-E’’ ). To rule out the possibility that Mhc upregulation is an artifact of the injury method used, we examined its expression in Naa30A mutants, which exhibit developmental defects in flight muscles and spontaneous muscle detachment in eight-day-old adults, along with loss of nuclear Mef2 expression [ 25 ]. In these mutants, Mhc upregulation coincided with Mef2 downregulation ( Figure S1 F-G’’ ), further supporting Mhc as a reliable marker of muscle damage in Drosophila . Download figure Open in new tab Figure S1. Mhc upregulation marks the damaged muscle fiber. (A-B’’). IFMs stained for Mhc (magenta) and Mef2 (cyan) show upregulation of Mhc and downregulation of Mef2 in the injured fiber (asterisks, B’-B’’) at 5DPI, compared to the uninjured IFMs (A’-A’’). (A’, B’). 2X magnification of the boxed regions in A and B. Scale bars: 200 μm. (C-C’’). Expression of the embryonic Mhc isoform ( weeP26 , green) in uninjured (C) and injured (C’-C’’) IFMs. Injury does not alter weeP26 expression (C’ - C’’). (D-D’’). The shorter Mhc isoforms ( Mhc-Iso-K, L, M (fTRG500) ; green) colocalize with anti-Mhc staining (magenta) and are upregulated in injured muscle (D’-D’’). (E-E’’). The longer Mhc-GFP isoforms ( Mhc-Iso-A, F, G (fTRG519) ; green) are not expressed in adult IFMs and are not upregulated in response to muscle injury (E’-E’’). Scale bars: 200 μm. (F-G’’). In Naa30A mutant IFMs, defective muscle attachment correlates with increased Mhc expression (magenta, G), upregulation of short Mhc-GFP isoforms (green, G’), and a loss of Mef2 (cyan, G’’). (H-I’’). Co-expression of hemocytes markers HmlΔ-Gal4, UAS-mCD8GFP (green) and Pxn-RFP (magenta) reveals two subsets of hemocytes associated with the IFMs: Pxn+Hml+ (arrows) and Pxn+Hml- (arrowheads). Nuclei are visualized with TO-PRO (blue). Scale bars: 200 μm. (H’’-I’’). 3X magnification of the boxed regions in H and I. Scale bars: 100 μm. (J-M’). Co-expression analysis of Enh3-Gal4; UAS-mcD8GFP (green) and Pxn-RFP (magenta) shows that all Enh3+ cells express Pxn (J’-K’). No Mef2 expression (cyan) is observed in these cells, in either undamaged (L’) or damaged (M’, dashed lines) fibers. Scale bars: 200 μm. Yellow asterisks in panels F’, G’’ and L’ mark non-specific signal detected in motoneurons. Download figure Open in new tab Figure S2. Early recruitment of hemocytes initiates tracheal remodeling after muscle damage. (A-B’). In Naa30A mutant flies, defective muscle attachment is associated with tracheal enrichment (gray, B’) and increased hemocytes accumulation (green, B’) compared to controls (A-A’). (C-K’). Time-course analysis of tracheal remodeling and hemocytes recruitment in Enh3-Gal4; UAS-mCD8GFP adult IFMs following localized stab injury. Tracheal branches are visualized by autofluorescence (gray), and hemocytes are labeled with GFP (green). Muscles are labeled with Mhc (magenta). Time points range from 1-hour post-injury (1HPI) to 20 days post-injury (20DPI). (L). Quantification of normalized Mhc intensity. (M). Quantification of the hemocytes number in the injured fiber shows a significant increase starting at 1HPI which remains elevated until 20DPI. (N). Tracheal quantification shows a marked increase in its density from 3DPI and remains sustained until 20 DPI. (****p < 0.0001, ***p < 0.001; Student’s t-test; n ≥ 10 heminota per condition). Scale bars: 200 μm. Download figure Open in new tab Figure 1. Hemocytes are specifically recruited to the damaged muscle fiber. (A-F). Adult indirect flight muscles (IFMs) imaged under uninjured conditions and at 2-and 5-days post-injury (2DPI and 5DPI). Mhc (magenta) marks muscles. Hemocytes (green) are visualized using the following reporters: Pxn-RFP in (A-C’’) and Enh3-Gal4; UAS-mCD8GFP in (D-F’’). Dashed yellow outlines indicate injured muscle fibers showing elevated Mhc expression. Boxed regions are magnified 5X. (G). Time-course analysis of hemocytes recruitment at 1, 2, 3, 4 and 5 DPI reveals heterogeneous kinetics of response to muscle damage. Enh3-positive, Pxn-positive, and Hml-positive hemocytes are represented by light gray, black, and dark gray bars, respectively (****p < 0.0001, Student’s t-test; n ≥ 10 heminota per condition). (G’). Schematic representation of the proportion of recruited hemocytes expressing different reporters. (H-I’’’). Distribution of hemocytes visualized with Enh3-Gal4; UAS-mCD8GFP (green) in control muscles (H-H’’’) and Naa30A mutants (I-I’’’). Muscles and muscle nuclei are stained with Mhc (magenta) and Mef2 (cyan) respectively. Scale bars: 200μm. Asterisks in panels H’’’ and I’’’ mark non-specific signal detected in motoneurons. The contribution of the hemocytes in IFMs repair remains largely unexplored. To address this, we started by tracking these cells at various time points after injury using the hemocytes reporters Peroxidasin ( Pxn ) ( Figure 1 A-C’’ ) and Hemolectin ( Hml ) ( Figure S1 H-I’’ ). Co-staining for these markers and Mhc revealed extensive recruitment of hemocytes to the injured muscle fibers ( Figure 1 A-C’ ; Figure S1 H-I’’ ). Notably, while Pxn+ hemocytes persist at the injured muscle for at least 5 DPI, Hml+ hemocytes enrichment declines over time ( Figure 1 G ), highlighting the heterogeneity of the adult hemocyte population, consistent with previous findings [ 26 ]. To further characterize the hemocytes recruitment dynamics, we examined the zinc finger homeodomain transcription factor Zfh1 reporter ( Enh3-Gal4 > UAS-mCD8GFP ), another hemocytes marker [ 9 , 27 ] ( Figure 1 D-F’’ ). Enh3+ hemocytes exhibited recruitment kinetics similar to Pxn+ cells ( Figure 1 G ). Importantly, at 5 DPI all Pxn+ cells were also Enh3+, whereas only a subset of Pxn+ cells were Hml+ ( Figure 1 G’ and Figure S1 H-K’’’ ). None of the recruited Enh3+ hemocytes expressed the myogenic marker Mef2 ( Figure S1 L’-M’ ). To confirm that hemocytes recruitment is a response to muscle damage, we analyzed Naa30A mutants. Consistently, we observed a selective recruitment of Enh3+ hemocytes to injured muscle fibers, identified by high Mhc expression and the loss of Mef2 nuclear expression ( Figure 1 H-I’’’ ). Altogether, these data show that adult hemocytes are rapidly and specifically recruited to injured muscle fibers, with distinct subpopulations displaying differential persistence over time. Hemocytes recruitment coordinates with tracheal remodelling following muscle injury We observed that hemocytes are closely associated with muscle tracheal cells under homeostatic conditions ( Figure 2A–A’ ). Previous studies have shown that adult tracheal cells can undergo dynamic remodeling and extend new branches toward damaged regions to support gut regeneration [ 18 , 19 ]. We therefore asked whether tracheal remodeling also occurs in response to IFMs injury and whether this process involves intercellular communication between tracheal cells and hemocytes. Our analysis revealed that muscle damage, whether caused by physical injury or in Naa30A mutant animals, leads to a significant increase in tracheal coverage of the injured fiber ( Figure 2 B-D and Figure S2 A-B’ ). The invasion is directed toward the injury site, with the highest branching density close to the damaged area and progressively decreasing with distance from the injury ( Figure 2 E ). We further observed that hemocytes remain spatially close to the tracheal branches, even under injured conditions, suggesting sustained interactions between these two cell types during the muscle damage response ( Figure 2 C’’ ). Download figure Open in new tab Figure 2. Muscle damage induces directed tracheal remodelling. (A-A’). Hemocytes (green/magenta; Enh3-Gal4, UAS-mCD8GFP; UAS-H2B-RFP ) are associated to the muscle tracheal cells (gray, autofluorescence). Scale bars: 25 μm. (B-C’’). Distribution of tracheal branches (gray, autofluorescence) and hemocytes (green, Enh3-Gal4; UAS-mCD8GFP ) in uninjured IFMs (B-B’’) and at 5DPI (C-C’’). Muscles are labeled with Mhc (magenta). (B’’, C’’). Boxed regions from (B and C), magnified 2X. Scale bars: 200 μm. (D). Quantification of tracheal coverage in (B) and (C) (****p < 0.0001, Student’s t-test; n = 19 heminota per condition; light, dark, and intermediate shading indicate data points from three independent replicates). (E). Quantification of tracheal coverage in (C) at the injury site (orange box), adjacent to the injury site (green box, approximately 135 μm away from the injury site), and at distant regions (blue box, within 300 μm of the lesion border) (*p= 0.0142, Student’s t-test; n=19 heminota). (F-F’). dSRF-Gal4 marks muscle-associated terminal tracheal cells (TTCs). TTC membranes (green) and nuclei (magenta) are indicated by arrowheads and arrows, respectively ( dSRF-Gal4, UAS-mCD8GFP; UAS-H2B-RFP ). dSRF-Gal4 is inactive in primary tracheal branches (asterisks in F’). Muscles are labeled with Mhc (blue). Scale bars: 25 μm. (G-H’’’). TTCs distribution in IFMs (green, dSRF-Gal4, UAS-mCD8GFP ) in uninjured muscles (G-G’’’) and at 5DPI (H-H’’’). Muscles are labeled with Mhc (magenta). (G’’, H’’). 4.5X magnifications of boxed regions in (G’) and (H’). Scale bar: 200 μm. (G’’’, H’’’). Orthogonal views of (G’’) and (H’’). Arrow in (H’’’) indicates TTC enrichment at the injured muscle. (I). Quantification of TTCs coverage from (G) and (H) (****p < 0.0001, Student’s t-test; n = 14 heminota per condition; light, dark, and intermediate shading points represent data points from three independent replicates). (J-J’). dSRF (green) marks TTCs nuclei associated with IFMs. Scale bars: 25 μm. (K-L’’’). Distribution of TTCs nuclei (green) in IFMs under uninjured conditions (K-K’’’) and at 5DPI (L-L’’’). Muscles are labeled with phalloidin (blue); tracheae are visualized by autofluorescence (gray). Anti-PH3 (magenta) is used to assess cell division. (K’’, L’’). 2X magnification of boxed regions in (K) and (L). Scale bars: 200 μm. Arrowheads and arrows indicate dSRF expression in the muscle and TTCs nuclei, respectively. (M). Proportion of TTCs nuclei located within a selected muscle relative to the total number of TTCs nuclei in the imaged sample (*p = 0.036, Student’s t-test; n = 17 heminota per condition; light, dark, and intermediate shading indicate data points from three independent replicates). To further investigate the tracheal response to injury, we focused on terminal tracheal cells (TTCs), specialized branches of the tracheal system known for their structural plasticity and ability to invade tissues, including muscle fibers, to deliver oxygen [ 14 , 15 , 28 ]. TTCs can be reliably identified by their distinctive morphology and the expression of the transcription factor Serum Response Factor (dSRF) ( Figure 2 F-F’ and J-J’ ). We visualized TTCs by expressing membrane-bound GFP under the control of dSRF-Gal4 ( dSRF-Gal4 > UAS-mCD8GFP ) ( Figure 2 G-H’’’ ). We observed a high density of TTCs within injured muscle fibers, with the highest concentration near the injury site ( Figure 2 H-H’’’ and I ). Orthogonal views showed that the recruited trachea are found not only on the muscle surface but also extend branches into the injured fibers ( Figure 2 G’’’-H’’’ ), consistent with the atypical muscle-tracheal organization optimized to deliver oxygen deep within the muscle, directly to sites of high metabolic demand [ 14 , 15 ]. Labelling TTCs nuclei with anti-dSRF confirmed the significant enrichment of dSRF+ nuclei along the injured fibers comparing to the uninjured muscles ( Figure 2 K-L’’ and M ). The tracheal enrichment at the injured muscle results from TTCs branching and recruitment rather than proliferation, as no PH3+ cells were detected in the muscle trachea cells ( Figure 2 K’’’-L’’’ ). Altogether, these data indicate that tracheal cells respond to muscle injury and target specifically the damaged fiber. We found that tracheal enrichment around injured muscles persists throughout the repair process and is still present 20 days post-injury ( Figure S2 C-K’ and N ), a stage at which muscle healing is expected to be complete [ 10 , 11 ]. A comparison of hemocyte recruitment kinetics with tracheal coverage density revealed that hemocyte recruitment precedes tracheal coverage of the injured fibers ( Figure S2 M and N ), raising the possibility of an interplay between the two-cell population in supporting the injured muscle fiber. Tracheal invasion of injured muscle fibers is triggered by local hypoxia The tracheal recruitment to injured muscle fibers ( Figure 2 ) suggests a critical oxygen demand of the damaged muscle. If this is the case, local hypoxia within injured fibers could serve as a key cue for tracheal attraction and directional growth toward the site of injury, as seen in other physiological contexts [ 29 ]. To assess oxygen variations within the injured muscle, we first used the Synthetic HRE (Hypoxia response element) nuclear sensor (SyHREns), a hypoxia reporter that serves as a proxy for evaluating intracellular oxygen fluctuations [ 30 ]. In normal condition, the SyHREns was found to be expressed in some TTCs, identifiable by both their triangular shape and expression of dSRF ( Figure 3 A-A’’ ). Importantly, we observed a pronounced enrichment of SyHREns-positive cells surrounding injured muscle fibers, in close proximity to the recruited tracheal branches ( Figure 3 B-D’’ ). The high number of SyHREns+ cells around the injured fibers remained stable for at least 20 DPI ( Figure 3 E ), indicating that local hypoxia persists in parallel with the sustained tracheal remodelling observed over the same period. Download figure Open in new tab Figure 3. Local hypoxia in response to muscle damage drives tracheal branching. (A-A’’). SyHREns reporter activity (green) is detected in a subset of TTCs marked by dSRF (magenta, arrows). Scale bars: 25 μm. (B-D’’). Expression of the SyHREns reporter (green) in unijured IFMs (B-B’’), at 5DPI (C-C’’) and 20DPI (D-D’’). Muscles are labeled with Mhc (magenta), and tracheal branches are visualized via autofluorescence (gray). (B’’, C’’ and D’’). 2X magnification of the boxed regions in B’, C’ and D’. Scale bars: 200 μm. (E). Quantification of SyHREns positive cells (green) per injured muscle fiber (****p 20 heminota per condition; light, dark and intermediate shading indicate data points from three independent replicates). (F-H’’). Sima (HIF-1α) downregulation in TTCs ( dSRF-Gal4; UAS-Sima RNAi ) prevents tracheal recruitment at the injured fiber. (F’’, G’’ and H’’). 2X magnification of the boxed regions in E’, F’ and G’. Scale bars: 200 μm. (I). Quantification of trachea coverage (****p < 0.0001, Student’s t-test; n = 13 heminota per condition). (J-K’’). Detection of Bnl (green, Bnl::GFP endo ) in control IFMs (J-J’’) and at 5DPI (K-K’’). Muscles are labeled with Mhc (magenta), and tracheal cells are visualized via autofluorescence (gray). (J’’, K’’). 4.5X magnification of the boxed regions in J and K. Scale bars: 200 μm. (J’’’, K’’’). Orthogonal views of (J) and (K). Asterisk in K’’’ indicates Bnl::GFP endo enrichment at the injured muscle. (L). Quantification of normalized Bnl::GFP endo intensity level in uninjured and injured muscle fiber at 5DPI (***p<0.0001, Student t-test, n = 15 heminota per condition from three independent replicates). Hypoxia-induced tracheogenesis is controlled by the transcription factor Similar (Sima)-the Drosophila homolog of hypoxia-inducible factor-1α (HIF-1α)- a central regulator of hypoxic and angiogenic responses [ 29 ]. Accordingly, silencing sima in TTCs severely decreased tracheal expansion towards the injured fibers ( Figure 3 F-I ). Bnl is a well-established downstream effector of hypoxic signalling, deposited locally on hypoxic tissues to promote tracheal attraction [ 29 ]. We therefore hypothesised that if the injured muscle undergoes hypoxia, Bnl should accumulate locally at the damaged fiber to facilitate tracheal remodelling. Analysing the expression of an endogenously tagged Bnl::GFP endo allele [ 31 ] revealed marked enrichment of Bnl at injured muscle fibers compared to adjacent intact ones ( Figure 3 J-K’’’ and L ). Altogether, these data suggest that muscle damage induces local muscle oxygen starvation, which in turn drives tracheal cell recruitment to the injured tissue. Hemocytes deposit FGF-like ligand Bnl at injured muscles to mediate tracheal recruitment During both embryonic and post-embryonic stages, tracheal guidance is regulated in a dose-dependent manner by local Bnl concentrations [ 29 ]. In adult flight muscle development, muscle-tracheal connections are established through the action of Bnl, which is secreted by muscle fibers and acts in a paracrine manner by binding to its receptor, FGFR/Breathless (Btl), expressed on tracheal cells [ 14 ]. The observed enrichment of Bnl in injured muscles ( Figure 3 J-L ) raises two key questions: Is Bnl/Btl signaling functionally involved in directing tracheal recruitment to damaged fibers? If so, what is the source of Bnl enrichment on the injured fibers? To begin addressing these questions, we conditionally overexpressed Bnl in a restricted subset of adult muscle fibers using the flip-out system combined with a regionally expressed driver ( Figure S3 A-C and Materials and Methods section). This targeted overexpression was sufficient to recruit tracheal cells to the ectopic Bnl-expressing regions ( Figure 4 A-C’ ), and the extent of this recruitment depended on the duration of Bnl expression: prolonged expression (5 days) triggered more extensive tracheal remodelling than short-term expression (24 hours). These findings showed that the adult tracheal system remains responsive to elevated Bnl levels in the IFMs, supporting the hypothesis that Bnl/Btl signalling directs tracheal pathfinding in response to muscle injury. Download figure Open in new tab Figure S3. Tracheal branching requires precise control of Bnl provided by the hemocytes. (A-B’). Expression pattern of the Shavenbaby (Svb) muscle regulatory enhancer (SME) (green, SME-Gal4; UAS-mCD8GFP ). SME-Gal4 drives GFP expression (green) in a subset of larval muscle progenitors located in the posterior region of the wing disc (A-A’) and in a subset of adult muscle nuclei (green and blue (DAPI), B-B’). Scale bars: 200 μm. (C). Schematic representation of the FLP-OUT system used to drive UAS-Bnl::GFP expression in a subset of adult muscle nuclei, using the Hs-FLP; FRT-SME-Stop-FRT-Gal4/CyO; tub-Gal80ts combination. (D-D’). The endogenous Bnl (green, Bnl::GFP endo ) is detected in indirect flight muscles (IFMs, labelled with Mhc, blue; arrowheads), tracheal branches (autofluorescence, gray; asterisks), and hemocytes ( Pxn-RFP , magenta; arrows). Scale bars: 25 μm. (E-E’). Expression of the Bnl-LacZ reporter (green) is observed in terminal tracheal cells (TTCs) marked by dSRF (magenta; asterisks), and in IFMs nuclei (arrowheads). Scale bars: 25 μm. (F-F’). Bnl-LacZ (green) is detected in IFMs associated hemocytes labelled with Pxn-RFP (magenta; arrows). Scale bars: 25 μm. (G-I’’). Targeted downregulation of bnl in a subset of hemocytes using HmlΔ-Gal4 driver significantly reduces tracheal enrichment at the injured fiber, as quantified in (J). (*p < 0.05, Student’s t-test; n ≥ 10 heminota per condition). Scale bars: 200 μm. Download figure Open in new tab Figure 4. Hemocytes deliver Bnl to injured muscle to promote tracheal recruitment. (A-C’). Local overexpression of UAS-Bnl::GFP (green) in adult IFMs using SME-Gal4 driver induces enrichment of terminal tracheal cells (TTCs; dSRF, magenta) and increases tracheal branching (gray), compared with controls (A). Scale bars: 200 μm. (D-E’’). Bnl-LacZ reporter expression (green) is downregulated specifically in of the injured muscle fibers overexpressing Mhc (magenta), the nuclei are marked with DAPI (blue). (D’, E’’). 3X magnification of the boxed regions in D and E. Scale bars: 200 μm. (F-G’’). Endogenous Bnl expression (green, Bnl::GFP endo ) in uninjured IFMs (F-F’’) and at 5DPI (G-G’’). Muscles are labeled with Mhc (blue), tracheal branches are visualized via autofluorescence (gray), and hemocytes detected with the Pxn-RFP reporter (magenta). Arrows in G’’ indicate Bnl localization around hemocytes recruited to the injured muscle. Scale bars: 200 μm. (H-J’’). Downregulating bnl in the hemocytes (green, Enh3-Gal4; UAS-mCD8GFP, UAS-Bnl RNAi ) do not alter hemocytes recruitment but decreases tracheal invasion to the damaged fiber as shown in (J’-J’’) and quantified in (K) (**** p< 0.0001, Student t-test; n = 16 heminota from two independent replicates). Scale bars: 200 μm. (L). Schematic representation of the flight-assay setup used to assess flight performance. Flies are released at the top of a graduated cylinder, and landing position across three vertical zones is recorded. (L’). Quantification of the proportion of flies distributed across three sections under three conditions: Control Uninjured , Control Injured (5DPI) and Enh3-Gal4; UAS-Bnl RNAi Injured (5DPI) . Bars indicate the percentage of flies landing in each of the three flight-assay zones (***p<0.001, * p < 0.05 Chi-2 test). (M). Schematic model of hemocyte-driven tracheal recruitment required to maintain flight capacity following muscle injury. We next sought to identify the cellular source responsible for Bnl accumulation in injured muscle fibers. Under wild-type conditions, Bnl::GFP endo signal was detected in hemocytes, tracheal cells, and IFMs ( Figure S3 D-D’ ). However, since Bnl is a diffusible protein, these observations alone cannot distinguish between cells that produce Bnl and those that merely take it up. To address this, we used the Bnl-lacZ transcriptional reporter, which revealed active bnl expression in hemocytes, adult IFMs, and TTCs, confirming that these cell types are bona fide sources of Bnl under normal conditions ( Figure S3 E-F’ ). Given that canonical Bnl/Btl signalling operates in a paracrine manner, injured muscles and hemocytes emerge as the most plausible sources of injury-induced Bnl accumulation. To distinguish between these two possibilities, we examined Bnl-lacZ expression following injury. Surprisingly, we found that injured fibers nuclei lacked Bnl-lacZ activity, whereas neighboring uninjured fibers retained its expression ( Figure 4 D-E’’ ). These findings indicate that the injured muscle fibers themselves do not contribute to the localized Bnl accumulation after injury and suggest that the source of Bnl is extrinsic to the damaged tissue. By contrast, we observed a marked enrichment of Bnl::GFP endo signal surrounding Pxn+ hemocytes recruited to the damaged fibers, suggesting that hemocytes are responsible for depositing Bnl along injured muscles ( Figure 4 F-G’’ ). To assess the functional contribution of hemocyte-derived Bnl to tracheal invasion, we selectively silenced bnl in hemocytes using either the Enh3-Gal4 or HmlΔ-Gal4 drivers. While hemocyte recruitment to injured muscles was unaffected, bnl knockdown significantly reduced tracheal invasion into damaged fibers compared with controls ( Figure 4 H-J’’ and K ; Figure S3 G-I’’ and J ). This signalling logic between the two cell types aligns with their close anatomical association under homeostatic conditions ( Figure 2 A-A’ and [ 13 ]. Given the critical role of the tracheal system in supporting tissue function [ 16 ], we next asked whether hemocyte-mediated tracheal recruitment via Bnl is required to maintain muscle flight performance after injury. To test this, we performed flight assays on uninjured controls, injured flies, and injured flies in which bnl was specifically depleted from hemocytes. Injured flies exhibited reduced flight capability compared with uninjured controls ( Figure 4 L-L’ ). Importantly, flies lacking hemocyte-derived Bnl showed an even greater impairment in flight, underscoring the functional importance of this signalling in promoting tracheal remodelling and preserving muscle performance following injury ( Figure 4 M ). The receptor FGFR/Btl is essential for injury-induced tracheal guided outgrowth The finding that Bnl orchestrates tracheal morphogenesis in response to muscle damage suggested in turn that it activates its receptor Btl, on tracheal cells, thereby driving guided outgrowth and branching. To investigate this, we examined Btl expression in adult flight muscles using an endogenously tagged Btl::cherry endo allele, which faithfully reports Btl expression [ 31 ]. Btl::cherry endo was detected along tracheal branches under homeostatic conditions and within tracheal structures invading the injured muscle ( Figure 5 A-B’’ ). To complement this approach, we expressed a membrane-tethered GFP (mCD8-GFP) under the control of a Btl-Gal4 driver ( Figure 5 C-E’ ). While Btl::cherry endo reflects consistent endogenous expression, Btl-Gal4 drives a more heterogeneous pattern across tracheal cells ( Figure 5 C-C’ ). Nevertheless, we observed a striking accumulation of tracheal branches labelled by Btl-Gal4 at injured muscles, reinforcing that Btl-expressing tracheal cells are actively recruited to damaged fibers ( Figure 5 D-E’ ). We then tested whether Btl function is required for proper tracheal pathfinding. To this end, btl was specifically silenced in tracheal cells using either dSRF-Gal4 or Btl-Gal4 drivers ( Figure 5 F-I’’ ). In both cases, tracheal invasion into injured fibers was significantly impaired compared to the control ( Figure 5 J ). Notably, the impairment was more pronounced with dSRF-Gal4 , consistent with its more uniform expression in terminal tracheal cells ( Figure 2 F-F’ ) compared with Btl-Gal4 ( Figure 5 C-C’ ). Together, these findings show that the Bnl/Btl signalling axis plays a pivotal role in guiding tracheal remodelling after muscle damage. While Btl expression is restricted to tracheal cells, localized Bnl secretion from hemocytes at the damaged fiber refines tracheal pathfinding, ensuring precise and targeted tracheal invasion into injured muscle ( Figure 5 K ). Download figure Open in new tab Figure 5. The FGF receptor Breathless (Btl) is expressed along tracheal branches and is required for tracheal recruitment to the damaged muscle. (A-B’’). Btl::Cherry endo (magenta) is detected along tracheal branches (gray) associated with indirect flight muscles (IFMs) labelled with Mhc (blue), in either uninjured (A) or injured muscles (B). Scale bars: 50 μm. Dashed yellow outlines indicate injured muscle fibers showing elevated Mhc expression. (C-C’). The Btl-Gal4 driver (green and magenta; Btl-Gal4, UAS-mCD8GFP; UASH2B-RFP ) is active in a subset of tracheal branches (arrows, C-C’). Arrowheads in C indicate the tracheal branches that do not express the Btl-Gal4 driver. Scale bars: 25 μm. (D-E’). Activity of the Btl-Gal4 driver (green, Btl-Gal4, UAS-mCD8GFP ) in uninjured condition (D-D’) and at 5DPI (E-E’). Muscles are labelled with Mhc (magenta). Scale bars: 200 μm. (F-I’’). Downregulating btl in the tracheal cells with either Btl-Gal4 driver ( Btl-Gal4; UAS-Btl RNAi , H-H’’) or dSRF-Gal4 driver ( dSRF-Gal4, UAS-Btl RNAi , I-I’’) prevents tracheal invasion to the damaged fiber. (J). Quantification of trachea coverage (****p <0.0001, **p=0.0053, Student’s t-test; n = 7 heminota per condition). Scale bars: 200 μm. (K). Schematic model of hemocyte-driven Bnl activates tracheal cells recruitment via Bnl/Btl signalling pathway. Hemocyte-derived ECM retains Bnl ligand at injured muscles Although Bnl is a diffusible molecule, we observed that it is specifically retained at the injured fiber ( Figure 3 K’-K’’ ), raising the question of what restricts its spatial distribution. Previous study has shown that Collagen IV, a principal structural component of the muscle ECM, is critical for retaining diffusible signals within defined tissue regions [ 32 ]. We therefore hypothesized that ECM components may similarly act to confine Bnl to the damaged fiber. Hemocytes are known to be major producers of ECM components during both Drosophila development and post-developmental stages [ 33 , 34 ], suggesting they could supply Collagen IV in response to adult muscle injury. To test this, we analysed Collagen IV distribution in adult flight muscles using the Vkg::GFP line ( Figure 6 A-B’’ ). Under homeostatic conditions, we observed a basal level of Vkg::GFP surrounding muscle fibers and along tracheal branches ( Figure 6 A’ ), and we found that hemocytes express Collagen IV ( Figure 6 A’’ ). Five days after injury, Vkg::GFP was markedly enriched in damaged fibers compared with intact muscles and uninjured controls ( Figure 6 A-B and A’’-B’’ ). To further assess the relationship between hemocytes and ECM remodeling, we examined Collagen IV distribution across multiple time points post-injury (1-20 DPI) and compared this to hemocyte recruitment dynamics ( Figure 6 C-D’’ and Figure S4 A-F ). Collagen IV accumulation closely correlated with the presence and distribution of hemocytes. Notably, regions where hemocytes were evenly distributed along the injured fiber displayed stronger and more uniform Vkg::GFP signal. Together, these data suggest that hemocytes are, at least in part, a source of Collagen IV within injured muscle tissue. Download figure Open in new tab Figure S4. Analysis of hemocyte recruitment, Collagen IV deposition, and Bnl localization following muscle injury. (A-F). Hemocyte recruitment (magenta, Pxn-RFP ) and Collagen IV distribution (green, Vkg::GFP ) in uninjured muscles (A) and at successive time points after injury (B-F), from 1 to 20 DPI. Scale bar: 200 μm. (G-G’’). The endogenous Bnl ligand (green, Bnl::GFP endo ) is detected around the hemocytes labeled with Zfh1 (magenta, arrows). (H-I’’). Depletion of Collagen IV in the hemocytes using Enh3-Gal4 driver (Enh3-Gal4, UAS-Vkg RNAi ) does not affect Bnl production by hemocytes (green, Bnl::GFP endo ; arrows) nor their recruitment to injured fibers (Zfh1, magenta) as quantified in (J) (ns, Student’s t-test; n ≥ 10 heminota per condition from two independent replicates). Scale bars: 200 µm. (H’’’-H’’’’ and I’’’-I’’’’). Hemocytes-specific knockdown of Collagen IV (Enh3-Gal4, UAS-Vkg RNAi ) does not affect the trachea enrichment in the damaged fiber (Quantifications in (K) ns, Student’s t-test; n ≥ 10 heminota per condition from two independent replicates). The ratio of tracheal enrichment in damaged fiber relative to adjacent non-injured fiber is significantly reduced upon vkg knockdown (K’) . Scale bars: 200 µm. (**p<0.01, Student’s t-test; n ≥ 10 heminota per condition from two independent replicates). (L-M’). Muscle-associated hemocytes (arrows; round-shaped nuclei), but not muscle nuclei (arrowheads; flattened nuclei), express both the SPARC-Gal4, UAS-myr-RFP reporter (magenta) and BM40-SPARC-GFP (green) in uninjured and injured muscles. Nuclei are labelled with DAPI (blue). Scale bars: 25 µm. Download figure Open in new tab Figure 6. Hemocytes deposit ECM components at injury muscles to constrain Bnl to damaged muscle fibers and maintain muscle function. (A-A’’). Collagen IV (green, Vkg::GFP ) is detected at basal levels around muscle fibers (arrow, A’), tracheal branches (arrowheads, A’), and hemocytes (magenta, Pxn-RFP ; arrow, A’’) in uninjured conditions. (B-B’’). At 5DPI, Collagen IV is strongly enriched on the injured muscle fibers, labelled with Mhc (blue). (C-D’’). Hemocytes recruitment (magenta, Pxn-RFP ) coincides with Collagen IV (green, Vkg::GFP ) distribution on the injured muscle, at 1DPI and 4DPI. Muscles are labeled with phalloidin (blue). Scale bars: 200 μm. (E-F’’). The endogenous Bnl (green, Bnl::GFP endo ) is enriched along damaged muscle fibers (magenta, Mhc) and is markedly reduced upon hemocyte-specific depletion of Collagen IV ( Enh3-Gal4, UAS-Vkg RNAi ). (G). Quantification of normalized Bnl::GFP endo intensity in injured fibers (****p27 heminota per condition from three independent replicates). (H). Schematic model of hemocyte-mediated ECM delivery required to confine Bnl to injured muscle fibers. (I-I’). SPARC (magenta) is detected in the IFMs associated hemocytes visualized by Enh3-Gal4 driver (green, Enh3-Gal4, UAS-mCD8GFP ). (J-J’). SPARC expression is increased in injured fibers compared to uninjured condition (I-I’). (K-K’). Adult hemocytes (magenta; SPARC-Gal4, UAS-myr-RFP ) express BM40-SPARC-GFP (green). (L-L’). BM40-SPARC-GFP produced by adult hemocytes is broadly distributed along the injured muscle fibers. (M-N’). Downregulation of SPARC in adult hemocytes ( Enh3-Gal4 > UAS-SPARC RNAi ) reduces Collagen IV enrichment (green, Vkg::GFP ) and impairs tracheal recruitment in injured fiber (gray, autofluorescence), as quantified in panels (O) and (P) respectively (O. ****p<0.0001, Student t-test, n = 20 heminota per condition from two independent replicates, P. ***p<0.001, Student t-test, n = 18 heminota per condition from two independent replicates). (Q). Quantification of the proportion of flies distributed across three sections (Top, Middle and Bottom) under three conditions: Control Uninjured, Control Injured (5DPI) and Enh3-Gal4; UAS-SPARC RNAi Injured (5DPI) (***p<0.001, ** p < 0.01 Chi-2 test). To test whether hemocyte-supplied Collagen IV is necessary for Bnl retention at injured fibers, we knocked down Collagen IV specifically in hemocytes and assessed Bnl::GFP endo localization ( Figure 6 E-F’’ and Figure S4 G-J ). In control flies, Bnl signal was strongly enriched along the injured fiber; however, this enrichment was significantly reduced following hemocyte-specific Collagen IV depletion ( Figure 6 E-F’’ and G ). Importantly, hemocytes recruitment to damaged fibers was not affected by Collagen IV depletion ( Figure S4 G-J ), indicating that Collagen IV is not required for immune cell recruitment to injured fibers. These data show that hemocyte-derived Collagen IV is required to spatially confine Bnl to the damaged fibers ( Figure 6 H ). We next asked whether loss of Bnl confinement following hemocyte-specific Collagen IV depletion affects tracheal remodelling after muscle injury. Absolute tracheal enrichment at injured fibers was comparable between control and Collagen IV–depleted conditions ( Figure S4 K ). However, the ratio of tracheal coverage at injured muscles relative to neighbouring intact muscles was significantly reduced when Collagen IV was depleted in hemocytes ( Figure S4 H’’’-H’’’’, I’’’-I’’’’ and K’ ). These data indicate that loss of ECM induces Bnl signal to spread beyond the injury site, thereby provoking non-specific tracheal remodelling in adjacent, intact muscles. Hemocyte-derived SPARC supports Collagen IV deposition, tracheal remodleling and muscle recovery Incorporation of Collagen IV into the extracellular matrix (ECM) requires accessory molecules for proper deposition and stabilization. SPARC is a collagen-binding glycoprotein essential for Collagen IV diffusion, stabilization, and incorporation into basement membranes [ 35 , 36 ]. We therefore investigated whether hemocytes also supply SPARC during muscle repair. We first examined SPARC localization in hemocytes associated with adult IFMs. Hemocytes, identified using Enh3-Gal4 driver, were positive for SPARC ( Figure 6 I-I’ ). Importantly, following muscle injury, SPARC levels were elevated along the injured fiber compared with controls, mirroring the distribution of Collagen IV ( Figure 6 I-J’ ). These observations suggest that SPARC is locally produced by hemocytes in adult muscle. To further confirm the cellular origin of SPARC enrichment in injured muscles, we employed the SPARC-Gal4 driver, a Gal4 trap line inserted into the endogenous SPARC locus, to drive membrane-targeted RFP (myr-RFP) expression in flies also carrying a BM40-SPARC-GFP fosmid reporter ( Figure 6 K-L’ and Figure S4 L-M’ ). This dual-reporter approach allowed simultaneous visualization of SPARC (via BM40-SPARC-GFP) and identification of SPARC-producing cells (via SPARC-Gal4 > myr-RFP). In uninjured muscle, BM40-SPARC-GFP and SPARC-Gal4-driven myr-RFP were co-expressed in hemocytes. Following injury, BM40-SPARC-GFP was broadly enriched along the damaged fiber, resembling endogenous SPARC protein, whereas SPARC-Gal4 activity remained restricted to hemocytes and was not detected in muscle nuclei ( Figure 6 L-L’ and Figure S4 M-M’ ). Together, these data demonstrate that recruited hemocytes are the primary source of SPARC at injured muscle fibers. We then tested whether hemocyte-derived SPARC is required for Collagen IV enrichment and tracheal remodelling. Depletion of SPARC specifically in hemocytes was sufficient to prevent both Collagen IV accumulation and tracheal remodelling at the injured fiber ( Figure 6 M-N’’ and O-P ). Finally, to evaluate the functional importance of SPARC in the muscle repair process, we performed flight assays and found that injured flies lacking hemocyte-derived SPARC exhibited significantly reduced flight ability compared with controls ( Figure 6 Q ). Altogether, these results show that hemocyte-derived SPARC is essential for ECM assembly, tracheal remodelling, and functional recovery following muscle injury. They also highlight that hemocytes act as multifunctional signalling hubs, simultaneously delivering trophic, structural, and guidance cues required to coordinate efficient muscle repair. Discussion Tissue repair requires the coordinated integration of immune, vascular, and structural responses, yet how these processes are mechanistically linked following muscle damage remains poorly understood. Here, we identify a signalling program in which hemocytes link muscle damage to vascular remodelling and extracellular matrix organization. We show that hemocytes respond to injury by locally delivering the FGF ligand Branchless to guide tracheal outgrowth toward damaged muscle, while simultaneously supplying extracellular matrix components, both of which are required for muscle maintenance after injury. Together, these findings indicate that hemocytes play a central role in organizing the regenerative microenvironment in adult Drosophila muscle ( Figure 7 ). Download figure Open in new tab Figure 7. Hemocyte-muscle-tracheal intercellular signaling orchestrates tracheal remodelling and muscle repair after injury. (A). Schematic representation of the hemocyte-muscle-tracheal signaling module activated in response to muscle injury. (B). Upon muscle damage, hemocytes are rapidly recruited to the injured fiber, where they secrete the FGF-like ligand Branchless (Bnl), which activates its receptor Breathless (Btl) on tracheal cells. Bnl-Btl signaling drives tracheal remodeling and directional outgrowth toward the injured muscle. In parallel, hemocytes deposit extracellular matrix (ECM) components that retain Bnl and stabilize trachea-muscle attachments. Together, these coordinated cellular mechanisms promote muscle repair and preserve flight performance. Immune-derived FGF acts as an intercellular signalling cue to coordinate vascular remodelling during muscle repair FGF signalling contributes to multiple aspects of vertebrate myogenesis and muscle regeneration [ 37 – 40 ]. For example, FGF2 stimulates myoblasts proliferation while suppressing differentiation in vitro , whereas FGF6 is required to maintain full regenerative capacity and MuSC pool size [ 41 , 42 ]. However, whether FGF signalling also mediates vascular remodelling during vertebrate muscle repair, as it does in other regenerative contexts, remains unclear. One explanation for this gap may be the extensive functional redundancy of the vertebrate FGF family, which comprises multiple ligands with overlapping expression patterns and signalling activities. This redundancy complicates precise genetic dissection of individual FGF functions during regeneration. Moreover, the cellular sources of FGF ligands during muscle regeneration remain poorly defined [ 42 ], further limiting targeted genetic ablation strategies. Here, we show that following muscle damage, hemocytes are rapidly recruited to and spread over the injured muscle surface, where they secrete the FGF ligand Branchless (Bnl) to promote localized tracheal outgrowth and precise branch targeting ( Figure 7 ). This mode of immune-driven FGF delivery contrasts with the prevailing model in which the tissue requiring vascular or tracheal infiltration is itself thought to produce the FGF signal [ 29 , 43 ]. The identification of a non–cell-autonomous source of FGF during muscle repair is therefore unexpected and raises an important question: why does the injured muscle fiber not itself express Bnl to attract tracheal branches? One possible explanation is the need to tightly restrict tracheal growth. Bnl is a potent chemoattractant, and its widespread expression by muscle fibers could lead to excessive or ectopic tracheogenesis, resulting in inappropriate branch recruitment and disruption of normal tracheal network architecture. In contrast, hemocytes are comparatively sparse and become selectively localized to damaged regions, enabling the spatially restricted secretion of Bnl. Such targeted delivery would ensure that tracheal remodelling occurs specifically at sites of injury, where oxygen delivery and tissue repair are most acutely required. Local ECM delivery optimizes directed vascular outgrowth during muscle repair The ECM is essential for muscle integrity and function [ 44 ]. However, its role in muscle regeneration—particularly how its remodelling is coordinated with other cellular processes during muscle repair—remains incompletely understood. In vertebrates, fibro-adipogenic progenitors (FAPs) are the principal source of ECM within muscle tissue [ 45 ]. Drosophila lack a dedicated FAP population. During development, hemocytes are a major source of ECM components in flies [ 33 , 46 ]. Here, we show that adult hemocytes also produce key ECM components, including Collagen IV and SPARC, that are required for muscle repair. These findings suggest that Drosophila hemocytes share functional similarities with mammalian FAPs and may compensate for their absence, pointing to an evolutionarily flexible strategy for muscle regeneration in organisms with fewer specialized cell types. Moreover, we find that hemocyte-derived ECM forms a specialized local microenvironment in which FGF distribution is spatially constrained, thereby directing efficient vascular growth toward damaged muscle. Whether the ECM physically interacts with FGF or instead acts primarily as a diffusion barrier remains to be determined. Overall, these findings provide a mechanistic example of how extracellular matrix remodelling is coordinated with vascular remodelling to support muscle repair. Do Drosophila repair or regenerate their flight muscles? The IFMs generate the high-frequency wing beats that enable flight, a behavior essential for predator avoidance, foraging, and mating. Given their vital function, IFMs must be maintained in optimal condition to ensure sustained performance throughout the animal’s lifespan. This need for resilience is also underscored by the observation that over 30% of wild-caught flies exhibit physical injuries, frequently located in the thorax and abdomen [ 47 ]. Although a rare population of satellite-like cells has been identified in adult flies, the regenerative capacity of adult Drosophila muscles remains debated [ 48 ]. A major limitation has been the lack of appropriate molecular markers to monitor both muscle regeneration and muscle stem cell (MuSC) activity. In our study, we identified a specific isoform of the Myosin Heavy Chain (Mhc) that is consistently upregulated in damaged fibers. However, unlike in vertebrates, where MyHC-emb levels return to baseline following regeneration, Mhc expression in Drosophila remains elevated even 20 days post-injury ( Figure S2 ). This makes Mhc a sensitive marker for identifying injury, but less useful for assessing complete muscle recovery. Adult MuSCs have been identified via the transcription factor Zfh1/ZEB and its reporter Enh3 [ 9 , 10 ], the only presently known marker for these cells. Zfh1 is required for MuSCs maintenance, an equivalent role of ZEB was subsequently shown in mice [ 49 ]. Interestingly, both Zfh1 and ZEB are expressed in the muscle associated immune cells [ 9 , 49 ]. In our study, we found that following muscle injury, 100% of the cells recruited to damaged fibers are both Enh3⁺ and Pxn⁺ (a plasmatocytes marker), while being negative for the muscle-specific marker Mef2 ( Figure S1 L-M’ ). This strongly suggests that hemocytes are the predominant, if not exclusive cell population migrating to injured muscle. These findings suggest that adult Drosophila flight muscles possess limited intrinsic regenerative capacity and may instead rely on compensatory mechanisms such as increased oxygenation through tracheal remodelling and structural reinforcement via extracellular matrix (ECM) deposition-to cope with muscle damage. This is consistent with the broader idea that short-lived species prioritize immediate survival over long-term regenerative investment [ 50 ]. In such organisms, evolution favors fast and efficient repair mechanisms that support rapid recovery and fitness over complex, energy-intensive regeneration programs. Materials and Methods Drosophila melanogater strains and genetics All Drosophila melanogaster stocks were grown on standard medium at 25°C. The following stains were used: w 118 as wild type (wt), Pxn-RFP [ 51 ], HmlΔ-Gal4 (BL#30140) , Enh3-Gal4 [ 9 ] , Mhc-Iso-K, L, M (fTRG500) and Mhc-Iso-A, F, G (fTRG519) [ 24 ], weeP26 [ 23 ], Bnl::GFP endo and Btl::cherry endo [ 31 ], Bnl-LacZ (BL#11704),, dSRF-GAL4 (BL#25753), Btl-Gal4 (BL#78328), UAS-Btl RNAi (BL#435544), UAS-Bnl RNAi (BL#34572), UAS-Bnl::GFP [ 52 ], Vkg::GFP (BL#98343), BM40-SPARC-GFP (Sarov et al., 2016), SME-Gal4 (VT 0576871), UAS-H2B-RFP [ 53 ], UAS-mCD8GFP (BL#5137), UAS-myr-mRFP (BL#7118), SyHREns [ 30 ], UAS-Sima RNAi (BL#26207), SPARC-Gal4 (BL#77473), UAS-SPARC RNAi (BL#40885), UAS-Vkg RNAi (BL#50895). Naa30A deletion males were obtained from crosses between Naa30A Δ74 /FM0 females and either UAS-mCD8GFP; Enh3-Gal4 ( Figure 1 ) or Mhc-Iso-K, L, M ( Figure S1 ) males. RNAi experiments were carried out at 29°C. To achieve temporal control of gene expression in adult tissues, the tubGal80 ts was employed, allowing restriction of Gal4 activity to a specific time window. Fly crosses were initially maintained at 18°C, and after adult eclosion, the progenies were transferred to 29°C, where they remained until dissection. An exception was made for the Bnl knockdown experiments shown in Figure 4 J-J’’ , which were performed without the tubGal80 ts transgene. Generation of SME-FRT-stop-FRT-Gal4 fly line and flip-out induction procedure SME (Shavenbaby Muscle Enhancer) is one of several enhancers of the shavenbaby gene (VT057071-Gal4). It is active in larval muscle progenitors (AMPs) ( Figure S3 A-A’ ) and remains active in a subpopulation of localized muscle nuclei in the adult indirect flight muscles (IFMs) ( Figure S3 B-B’ ). To generate the SME-FRT-stop-FRT-Gal4 construct, the cis-regulatory element from the original shavenbaby enhancer construct was used to replace the pAct promoter in the pAct-FRT-stop-FRT3-FRT-FRT3-Gal4 vector (Addgene plasmid #52889). The resulting construct was integrated into the attP site at cytological location 51D on chromosome II via embryo injection into vas-int; attP-51D flies. To induce localized overexpression of Bnl::GFP in adult IFM nuclei, hsflp; FRT-SME-stop-FRT-Gal4/CyO; tubGal80 ts females were crossed to UAS-Bnl::GFP males. Progeny were raised at 18°C and subjected to a 2-hour heat shock at 37°C in young adulthood ( Figure 4 A-C’ ). Immunohistochemistry and confocal microscopy Preparation and immunofluorescence of wing discs and adult muscles were performed as previously described in [ 54 ]. The following primary antibody were used: Chicken anti-βGal (1:100, Abcam, ab9361), Chicken anti-GFP (1:300, Abcam, ab13970), Mouse anti-Mhc (1:200, DSHB, 3E8-3D3-s), Mouse anti-dSRF (1:20, a gift from Mark A. Krasnow, Stanford university California, USA), Rabbit anti-Mef2 (1:500, DSHB, Mef2-p), Rabbit anti-PH3 (1:100, Cell Signaling Technology, #9713), Rabbit anti-SPARC (1:500, a gift from Maurice Ringuette, University of Toronto, Canada), Rabbit anti-Zfh1 (1:5000, a gift from Ruth Lehmann, New York, USA), Rat anti-RFP (1:800, Chromotek, 5F8-20), Alexa-conjugated Phalloidin (1:200, Thermo fisher scientific life technologies). Induction of flight muscle injury Flight muscle injury was induced following previously described protocol [ 55 ]. Adult flies (2-3 days post-eclosion) were anesthetized and positioned laterally under a stereomicroscope. A single, localized stab wound was inflicted manually using a fine stainless-steel needle (Minutien Pins, 0.1 mm diameter; #26002–10) to minimize tissue damage and avoid extensive injury. Age-matched, uninjured flies served as controls. Flight assay Flight ability was assessed using a 1-m-long cylindrical flight column internally lined with paper coated with a thin layer of glue. The adhesive sheet was divided evenly into three longitudinal sections corresponding to the Top, Middle, and Bottom regions of the tube ( Figure 4L ). For each experimental condition, an equal number of flies (approximately 20–30) was released into the tube. After all flies had landed, the paper was removed, flattened, and imaged for analysis. Image quantification was performed semi-automatically using a custom R script, Fly-Spy (available at https://github.com/Olivier-Josse/Fly-spy ). Images were first converted to grayscale and digitally divided into the three predefined regions. High-intensity spots corresponding to individual flies were identified by applying a manually adjusted threshold to ensure accurate segmentation. The number of flies detected in each region was quantified and expressed as a percentage of the total recovered flies. The distribution of flies across the three regions—reflecting high, intermediate, or low/absent flight ability—was compared between experimental conditions using a chi-square (χ²) test. Results are presented as stacked bar histograms Microscopy and data analysis Samples were imaged using a Leica SP8 microscope (BIOSIT, University of Rennes) at 20×, 40×, or 63× magnification and a resolution of 1024×1024 pixels. Images were processed with ImageJ and assembled using Adobe Illustrator. To quantify fluorescence intensity, all samples were processed and analyzed in parallel under identical conditions. Confocal images were acquired using the same laser settings across all samples. Sum slice projections were generated from each confocal stack. Regions of interest (ROIs) were manually defined based on protein expression, and mean fluorescence intensities were measured within each ROI. These values were then normalized to background levels measured in comparable areas of the same sample. Tracheal branches were visualized either by their intrinsic autofluorescence using a 405 nm UV laser [ 15 ] or by the expression of membrane-bound GFP driven by a tracheal-specific Gal4 driver (UAS-mCD8GFP). Tracheal coverage was quantified as pixel area per muscle fiber from maximum intensity Z-projections acquired at 20× magnification. Quantification was performed using the “Skeletonize” plugin in ImageJ, based on the method described by Perochon et al. (2021) [ 18 ], with minor modifications. Briefly, maximum intensity projections were generated from each confocal stack, tracheal branches were segmented by thresholding, and the resulting images were skeletonized. Skeleton pixel intensity was then measured specifically within injured muscle fibers and their corresponding contralateral control fibers. For Figure 2E , square regions of interest (ROIs) were placed at the injury site, as well as in proximal and distal regions, to assess local tracheal enrichment. Hemocyte and dSRF-positive cell numbers were quantified manually using ImageJ. Graphs and statistical analyses were performed using GraphPad Prism. Pair comparisons were statistically tested with a two-sided Student’s t-test. NS, not significant (p>0.05); *p < 0.05, **p < 0.01, ***p < 0.001 ****p< 0.0001. Competing interest The authors declare that they have no competing interests. Acknowledgments We would like to thank Jennifer Zanet, Cedric Polesello, Sougata Roy, Ruth Lehmann, Maurice Ringuette, Jean-Paul Vincent, and Mark A. Krasnow, as well as the Bloomington Drosophila Stock Center, the Vienna Drosophila Resource Center (VDRC), and the Developmental Studies Hybridoma Bank for providing Drosophila strains and antibodies. We are grateful to Juila Cordero, Lucas Waltzer, Laetitia Bataille, Alain Vincent, and Michèle Crozatier for their critical reading of the manuscript, and to other members of H.B’s laboratory for valuable discussions. We also thank Emmanuel Gallaud for helpful input during the early phases of the project. We acknowledge Xavier Pinson from the Microscopy Rennes Imaging Center (MRic, BIOSIT, Biogenouest) for technical assistance, and Cyrille Surbled for fly stock maintenance and technical help. Work in the Boukhatmi lab was supported by an AFM-Téléthon Trampoline Grant (#23108), ATIP-Avenir (CNRS), and ANR GenepiMuSC (ANR-23-CE13-0006). N.A. was supported by an AFM-Téléthon PhD fellowship (#23846) and an FRM fellowship (FDT202404018608). Work in the Cordero lab was supported by a Sir Henry Dale Fellowship (Wellcome Trust and Royal Society, 104103/Z/14/Z), a Wellcome ISSF Returner Scheme (204820/Z/16/Z), and a Tenovus Fellowship (Tenovus/S22-28) to J.P. Footnotes In this revised manuscript, we have refined the text and added new data (Figure S4) providing further support for our conclusions. Reference 1. ↵ Sun F , Poss KD . Inter-organ communication during tissue regeneration . Development . 2023 ; 150 ( 23 ). Epub 20231127. doi: 10.1242/dev.202166 . PubMed PMID: 38010139 ; PubMed Central PMCID: PMCPMC10730022 . OpenUrl CrossRef PubMed 2. ↵ Rodriguez C , Timoteo-Ferreira F , Minchiotti G , Brunelli S , Guardiola O . Cellular interactions and microenvironment dynamics in skeletal muscle regeneration and disease . Front Cell Dev Biol . 2024 ; 12 : 1385399 . Epub 20240522. doi: 10.3389/fcell.2024.1385399 . PubMed PMID: 38840849 ; PubMed Central PMCID: PMCPMC11150574 . 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Share Immune–Vascular intercellular signalling coordinates tissue plasticity during adult muscle repair in Drosophila Nourhene Ammar , Olivier Josse , Aurélien Guillou , Jessica Perochon , Hadi Boukhatmi bioRxiv 2025.06.25.661466; doi: https://doi.org/10.1101/2025.06.25.661466 Share This Article: Copy Citation Tools Immune–Vascular intercellular signalling coordinates tissue plasticity during adult muscle repair in Drosophila Nourhene Ammar , Olivier Josse , Aurélien Guillou , Jessica Perochon , Hadi Boukhatmi bioRxiv 2025.06.25.661466; doi: https://doi.org/10.1101/2025.06.25.661466 Citation Manager Formats BibTeX Bookends EasyBib EndNote (tagged) EndNote 8 (xml) Medlars Mendeley Papers RefWorks Tagged Ref Manager RIS Zotero Tweet Widget Facebook Like Google Plus One Subject Area Developmental Biology Subject Areas All Articles Animal Behavior and Cognition (7640) Biochemistry (17706) Bioengineering (13902) Bioinformatics (41978) Biophysics (21465) Cancer Biology (18611) Cell Biology (25528) Clinical Trials (138) Developmental Biology (13387) Ecology (19920) Epidemiology (2067) Evolutionary Biology (24332) Genetics (15615) Genomics (22519) Immunology (17747) Microbiology (40424) Molecular Biology (17194) Neuroscience (88662) Paleontology (667) Pathology (2839) Pharmacology and Toxicology (4827) Physiology (7650) Plant Biology (15160) Scientific Communication and Education (2046) Synthetic Biology (4302) Systems Biology (9826) Zoology (2271)