Abstract
Macrophages are a highly heterogeneous population of cells, with this diversity stemming in part from the existence of tissue resident populations and an ability to adopt a variety of activation states in response to stimuli. Drosophila blood cells (hemocytes) are dominated by a lineage of cells considered to be the functional equivalents of mammalian macrophages (plasmatocytes). Until very recently plasmatocytes were thought to be a homogeneous population. Here, we identify enhancer elements that label subpopulations of plasmatocytes, which vary in abundance across the lifecourse of the fly. We demonstrate that these plasmatocyte subpopulations behave in a functionally-distinct manner when compared to the overall population, including more potent migratory responses to injury and decreased clearance of apoptotic cells within the developing embryo. Additionally, these subpopulations display differential localisation and dynamics in pupae and adults, hinting at the presence of tissue-resident macrophages in the fly. Our enhancer analysis also allows us to identify novel candidate genes involved in plasmatocyte behaviour in vivo. Misexpression of one such enhancer-linked gene (calnexin14D) in all plasmatocytes improves wound responses, causing the overall population to behave more like the subpopulation marked by the calnexin14D-associated enhancer. Finally, we show that, we are able to modulate the number of cells within some subpopulations via exposure to increased levels of apoptotic cell death, thereby decreasing the number of plasmatocytes within more wound-responsive subpopulations. Taken together our data demonstrates the existence of macrophage heterogeneity in Drosophila and identifies mechanisms involved in the specification and function of these plasmatocyte subpopulations. Furthermore, this work identifies key molecular tools with which Drosophila can be used as a highly genetically-tractable, in vivo system to study the biology of macrophage heterogeneity.
Introduction
Macrophages are key innate immune cells responsible for clearing infections, debris and apoptotic cells, the promotion of wound healing and are necessary for normal development [1]. However, their aberrant behaviour can also cause or exacerbate numerous human disease states, including cancer, atherosclerosis and neurodegeneration [1]. Macrophages are a highly heterogeneous population of cells, which enables them to carry out their wide variety of roles, and this heterogeneity arises from diverse processes. These processes include the dissemination and maintenance of tissue resident populations [2] and the ability to adopt a spectrum of different activation states (termed macrophage polarisation), which can range from pro-inflammatory (historically termed as M1-like) to anti-inflammatory, pro-healing (M2-like) macrophage activation states [3,4].
Macrophage heterogeneity appears to be conserved across jawed vertebrate lineages. Evidence suggests the existence of pro-inflammatory macrophage populations [5] and myeloid-derived microglia in zebrafish [6,7], with polarisation also a well-defined phenomenon in other fish species [8]. Vertebrate macrophages interact with and can become polarised in response to signals produced by Th1 and Th2 cells, leading to acquisition of M1-like and M2-like activation states, respectively. To date this form of heterogeneity has been considered to be restricted to organisms containing both an adaptive and an innate immune system. B and T cell-based adaptive immunity is thought to have evolved in teleost fish [9] and the diversity of macrophage populations in organisms possessing only an innate immune system appears more restricted. However, even comparing mammals as closely related as mice and humans, macrophage markers can be highly divergent [10], therefore other approaches and markers might be required to identify equivalent macrophage diversity in lower organisms.
Macrophage heterogeneity has been extensively studied in mammalian systems and, although this has provided a good understanding of how macrophages determine their polarisation state, this has also identified considerable complexity with many activation states possible [11]. Additional complexity arises with both M1-like and M2-like macrophages found at the same sites of pathology, for example within atherosclerotic plaques [12]. Furthermore, the cytokine profiles that can be induced in vitro depend on the exact activation methods used experimentally and these do not necessarily reflect polarisation states in vivo [13], while other macrophage subpopulations may be missed by in vitro approaches. Given these intricacies, it is clear that we still need to better understand the fundamental components and pathways responsible for the specification of different macrophage subtypes, particularly in vivo. Recently the “macrophage-first” hypothesis has been proposed, re-emphasising the idea that acute signals polarise macrophages ahead of the involvement of T cells [8]. Consequently, organisms without a fully-developed adaptive immune system represent intriguing models in which to examine this idea and better understand macrophage heterogeneity in vivo.
Drosophila melanogaster has been extensively used to study innate immunity [14], but lacks an adaptive immune system. Fruit flies possess three types of blood cell (also referred to as hemocytes): plasmatocytes, crystal cells and lamellocytes. Of these, plasmatocytes are functionally equivalent to vertebrate macrophages [15,16], with the capacity to phagocytose apoptotic cells and pathogens, secrete extracellular matrix, disperse during development and migrate to sites of injury [17]. Although Drosophila blood lineages are considerably less complex than their vertebrate equivalents, they are specified via transcription factors related to those used during vertebrate myelopoiesis, including GATA and Runx-related proteins [15]. Furthermore, plasmatocytes utilise evolutionarily-conserved genes in common with vertebrate innate immune cells to migrate (e.g. SCAR/WAVE, integrins and Rho GTPases [18–22]) and phagocytose (e.g. the CED-1 family member Draper [23] and CD36-related receptor Croquemort [24]). Given these striking levels of functional and molecular conservation, Drosophila has been extensively used for research into macrophage behaviour in vivo with its genetic tractability and in vivo imaging capabilities facilitating elucidation of different macrophage behaviours conserved through evolution [16,17]. However, despite these evolutionarily-conserved commonalities, the plasmatocyte lineage has, until very recently, been considered a homogeneous cell population. Hints that Drosophila plasmatocytes may exhibit heterogeneity exist in the literature with variation in marker expression observed in larval hemocytes [25] and non-uniform expression of TGF-β homologues upon injury or infection in adults [26]. Recent single-cell RNA-sequencing (scRNA-seq) experiments performed on larval hemocytes demonstrated the presence of multiple clusters of cells, which were interpreted as representing either different stages of differentiation or functional groupings [27,28]. However, the in vivo identification of subtypes and insights into the roles and specification mechanisms of potential macrophage subtypes in Drosophila has not yet been described.
Here, we describe the first identification and characterisation of molecularly and functionally-distinct plasmatocyte subpopulations within Drosophila melanogaster. Drawing on a collection of reporter lines [29], we have identified regulatory elements that define novel plasmatocyte subpopulations in vivo. We show that these molecularly-distinct subpopulations exhibit functional differences compared to the overall plasmatocyte population and that the proportions of cells within these subpopulations can be modulated by external stimuli such as increased levels of apoptosis. Furthermore, we show that misexpression of a gene associated with a subpopulation-specific enhancer element is able to modulate plasmatocyte behaviour in vivo, thereby identifying novel effector genes of plasmatocyte subpopulation function. Together our findings reveal that macrophage heterogeneity is a fundamental and evolutionarily-conserved characteristic of innate immunity that pre-dates the development of the adaptive immune system. This significantly extends the utility of an already powerful genetic model system and provides further avenues to understand regulation of innate immunity and macrophage heterogeneity.
Results
Drosophila embryonic plasmatocytes do not behave as a uniform population of cells
The macrophage lineage of hemocytes (plasmatocytes) has historically been considered a homogeneous population of cells. However, careful analysis of plasmatocyte behaviour in vivo suggested to us that this lineage might not be functionally uniform. For instance, imaging the inflammatory responses of plasmatocytes to epithelial wounds, we find that some cells close to injury sites rapidly respond by migrating to the wound, while other neighbouring cells fail to respond, (Figure 1a; Supplementary Movie 1). We also find that plasmatocytes exhibit variation in their expression of well-characterised plasmatocyte markers (crq-GAL4 [19,24]; Figure 1b-b’) and display a broad diversity in their migration speeds within the embryo (random migration at stage 15, Figure 1c-d). These professional phagocytes also display differences in their capacities to phagocytose apoptotic cells with some cells engulfing many apoptotic particles, whereas others engulf very few, if any (Figure 1e). Furthermore, phagocytosis of microorganisms by larval hemocytes also varies significantly from cell-to-cell in vitro (Figure 1f). These differences within the plasmatocyte lineage led us to hypothesise that this cell population is more heterogeneous than previously appreciated.
Discrete subpopulations of plasmatocytes are present in the developing Drosophila embryo
Given the diversity in plasmatocyte behaviour (Figure 1), we hypothesised that macrophage heterogeneity represents an evolutionarily-conserved feature of innate immunity, which therefore originally evolved in the absence of an adaptive immune system. To address this and look for molecular differences between plasmatocytes, we examined transgenic enhancer reporter lines (VT-GAL4 lines) produced as part of a recent large-scale tilling array screen [29] that had been annotated as labelling hemocytes (http://enhancers.starklab.org/). Based on examination of the published VT-GAL4 expression patterns, we identified VT-GAL4 lines that appeared to label reduced numbers of plasmatocytes in the embryo, reasoning that plasmatocyte subpopulations could be molecularly identified on the basis of differences in reporter expression. While a number of the enhancers appeared to label all plasmatocytes (e.g. VT41692-GAL4), we identified several that labelled discrete numbers of plasmatocytes (Figure 2a). We next confirmed that the cells labelled by these VT-GAL4 lines were plasmatocytes by using these constructs to drive expression of UAS-tdTomato in the background of a GAL4-independent, pan-hemocyte marker (srp-GMA (GFP-tagged actin-binding domain of moesin); Figure 2b-d; [30]). As initially predicted based on their morphology and position during embryogenesis, each of the VT-GAL4 lines marking potential subpopulations did indeed express in the hemocyte lineage Figure 2e). These subpopulation cells were identified as plasmatocytes based upon their morphology, the absence of lamellocytes in embryos and the non-migratory nature of crystal cells (Figure 2e; [16]) and could be observed to follow both the dorsal and ventral migration routes [17] used by these cells during their developmental dispersal (Figure 2e). In order to quantify the proportion of cells labelled by each VT-GAL4 line, we counted the number of cells labelled on the ventral midline of the developing stage 15 embryo, using VT-GAL4 lines to drive GFP expression. This verified reproducible and consistent labelling of discrete subsets of plasmatocytes (Figure 2f-h), suggesting that these cells represent stable subpopulations within this macrophage lineage.
Subpopulations of Drosophila plasmatocytes vary across development: subpopulation dynamics in larvae and white pre-pupae
Having identified subpopulations of plasmatocytes in the embryo, we then tested other stages of the life cycle to see how expression might be maintained or modulated throughout development. In order to exclude potential expression in non-hemocyte cells (e.g. non-plasmatocyte cells apparent in Figure 2e), we labelled subpopulation cells specifically using a split GAL4 approach [31], via which only cells expressing both serpent (a well-characterised hemocyte marker; [32]) and the named VT enhancer would be labelled via transcriptional activation of UAS transgenes (Supplementary Figure 1).
Imaging of L3 larvae containing split GAL4 constructs (srp-AD;VT-DBD – henceforth abbreviated to VTn) driving UAS-stinger revealed that very few subpopulation cells were present at this stage (Figure 3a-f). From larval development onwards, we cannot use cell morphology to discriminate between plasmatocytes and other hemocyte lineages (crystal cells and lamellocytes) and therefore refer to subpopulation cells as hemocytes for these subsequent stages. Using this approach, VT32897 and VT17559 labelled the most cells (Figure 3c-d), with only the occasional cell present in VT57089 larvae (Figure 3e) and cells essentially absent from VT62766 larvae (Figure 3f). Labelled cells were also present in the head region, along the dorsal vessel (the fly heart) and between the salivary glands (which exhibit non-specific labelling) in VT32897 larvae. The VT32897 head region cells are potentially specifically localised hemocytes, whereas cells at the remaining two sites are likely to correspond to serpent-positive nephrocytes and garland cells [33,34], respectively (Figure 3d). VT57089 shows additional staining in the head region (potentially the Bolwig organ; Figure 3e) and, as per the dorsal vessel-associated cells in VT32897 (Figure 3d), this can be observed in positive controls (data not shown). These patterns closely resemble patterns observed using the initial VT-GAL4 reporters, albeit with more restricted labelling due to our split GAL4 approach (data not shown).
Imaging of white pre-pupae (WPP), the stage that marks the beginning of pupal development and metamorphosis, showed very similar patterns across the split GAL4 VT enhancer lines (Figure 3g-l), but with a further reduction in the numbers of cells labelled. It was possible to observe the occasional cell moving in circulation within WPP, strongly suggesting these cells are hemocytes (Supplementary Movies 2 and 3). Live imaging of VT32897 WPP also confirmed association of cells with the pumping dorsal vessel (Supplementary Movie 4). Significantly, this data indicates that the presence of subpopulations within embryos is not simply a consequence of slow accumulation of fluorescent proteins by weak drivers, since these enhancer-based reporters do not label an ever-increasing number of cells as development proceeds. Overall, the numbers of hemocytes within subpopulations decreases over larval and early pupal stages demonstrating that plasmatocyte subpopulations are developmentally regulated. Such changes could reflect specific and changing requirements for specialised plasmatocyte subpopulations across the life cycle, for example an association with processes required for organogenesis [35–37]. This specific localisation of subpopulation cells also indicates the potential for tissue-resident macrophages in Drosophila.
Subpopulation cells return in large numbers during pupal development
Since subpopulation cells appear associated with stages of development when organogenesis and tissue remodelling occurs, we hypothesised that hemocyte subpopulations would return during metamorphosis. Imaging pupae at various times after puparium formation (APF) revealed that subpopulation cells re-emerged in large numbers during this stage, but with distinct dynamics (Figure 4a-f): VT17559 cells have already returned in very substantial numbers by 18h APF (Figure 4c), whereas VT32897 reporter expression reappeared between 24 and 48h APF (Figure 4d). VT57089 and VT62766-labelled cells increased in numbers more gradually over the course of pupal development (Figure 4e-f).
Subpopulations display distinct dynamics and localisation in adults
Immediately after adults hatch, large numbers of split GAL4-labelled cells can be observed across all lines and are present in selective regions that overlap with the overall adult hemocyte population (Figure 5a-e). The overall hemocyte population remains detectible as adults age (0-6 weeks; Figure 5a), however not all subpopulations exhibit an identical localisation or dynamics during this time (Figure 5b-e). VT57089 and VT62766 cells largely disappear by 1 week (Figure 5d-e) and the majority of VT17559-labelled cells are absent by 2 weeks (Figure 5b). By contrast, VT32897 cells persist for at least 6 weeks of adult life and are particularly prominent in the thorax at 4 weeks (Figure 5c). Other differences in localisation are also apparent with cells particularly obvious in the legs for the VT17559 line (Figure 5b, day 1-2 weeks), whereas VT57089 and VT62766-labelled cells are more closely associated with the thorax and dorsal abdomen (Figure 5d-e, day 1). Labelled cells are also present in the proboscis for several lines (Figure 5c-e). The distinct dynamics of subpopulation cells strongly suggests these subpopulations are at least partially distinct from each other and highlights their plasticity during development, with their presence, return and disappearance correlating with changes in the biology of blood cells over the entire lifecourse.
Subpopulation cells behave in a functionally-distinct manner compared to the overall plasmatocyte population
Given that the VT lines identified above are specifically and dynamically expressed in subpopulations of hemocytes during Drosophila development, we next set out to investigate whether the labelled subpopulations are also functionally distinct using a range of immune-relevant assays. The ability of vertebrate macrophages to respond to pro-inflammatory stimuli, such as injuries, can vary according to their activation status [38,39]. To investigate this in our system, a well-established assay of inflammatory migration [19] was employed (Figure 1a; Supplementary Movie 1). Strikingly, following laser-induced wounding, cells labelled by three VT-GAL4 lines (VT17559-GAL4, VT32897-GAL4 and VT62766-GAL4) showed a significantly more potent migratory response to injury. In each case a greater proportion of labelled subpopulation cells migrated to wounds, compared to the overall hemocyte population as labelled by a pan-plasmatocyte driver (Figure 6a-c). Consistent with our results above, plasmatocytes labelled by the VT lines represent a subset of the total number of hemocytes present ventrally in stage 15 embryos (Figure 6d).
We next investigated in vivo migration speeds of the embryonic plasmatocyte subpopulations (as per Figure 1c-d). Stage 15 embryos were imaged for 1 hour and individual plasmatocyte movements were tracked (Figure 6e-f). Only the VT17559-GAL4 labelled plasmatocyte subpopulation displayed statistically significantly faster rates of migration compared to the overall plasmatocyte population (labelled using srp-Gal4; Figure 6g). There were no differences in directionality (cell displacement divided by total path length) for any of the subpopulations, suggesting that the mode of migration was similar across these lines and with that of the overall population (Figure 6h).
Apoptotic cell clearance (efferocytosis) represents another evolutionarily-conserved function performed by embryonic plasmatocytes (Figure 1e; [40]). Therefore, we investigated this function in subpopulations, using numbers of vacuoles per cell as a proxy for this process [18]. Cells labelled via VT17559-GAL4, VT57089-GAL4 and VT62766-GAL4 (but not VT32897-GAL4) contained fewer vacuoles than the overall plasmatocyte population (Figure 6i-k), suggesting that these discrete populations of cells are less effective at removing apoptotic cells inside the developing embryo.
Finally, we examined cell size and shape of labelled plasmatocyte subpopulations. Vertebrate macrophages are highly heterogeneous, with distinct morphologies dependent upon their tissue of residence or polarisation status [41–43]. We found no obvious size or shape differences between VT-GAL4 labelled cells and the overall plasmatocyte population (Supplementary Figure 2a-e). This was also the case when VT-GAL4 positive cells were compared to internal controls (VT-GAL4 negative cells within the same embryos) for a range of shape descriptors (Supplementary Figure 2f-i). Similarly, we were unable to detect differences in ROS levels or the proportion of VT-GAL4 labelled plasmatocytes that phagocytosed pHrodo-labelled E. coli compared to controls (Supplementary Figures 3 and 4), two processes associated with pro-inflammatory activation of macrophages [44].
Taken together these data show that the subpopulations of plasmatocytes identified via the VT-GAL4 reporters exhibit functional differences compared to the overall plasmatocyte population (Table 1). Therefore, as well as displaying molecular differences in the form of differential enhancer activity, and hence reporter expression, these discrete populations of cells behave differently. This strongly suggests that these cells represent functionally-distinct subpopulations and that the plasmatocyte lineage is not homogeneous. Furthermore, not all subpopulations displayed identical functional characteristics, suggesting that there are multiple distinct subtypes present in vivo, although some overlap between subpopulations seems likely. For example, VT17559-GAL4 labelled cells were more effective at responding to wounds and migrated more rapidly, but carried out less phagocytosis of apoptotic cells. By contrast, VT32987-GAL4 labelled cells only displayed improved wound responses (Figure 6).
VT enhancers identify functionally active genes within plasmatocytes
In the original study that generated the VT-GAL4 collection, the majority of active enhancer fragments tested were found to control transcription of neighbouring genes [29]. Thus, genes proximal to enhancers that label plasmatocyte subpopulations represent candidate regulators of immune cell function (Table 2; Figure 7a). VT62766-GAL4 labels a subpopulation of plasmatocytes with enhanced migratory responses to injury (Figure 6) and this enhancer region is found within the genomic interval containing paralytic (para), which encodes a subunit of a voltage-gated sodium channel [45], and upstream of the 3’ end of calnexin14D (cnx14D) (Figure 7a). cnx14D encodes a calcium-binding chaperone protein resident in the endoplasmic reticulum [46]. Alterations in calcium dynamics are associated with clearance of apoptotic cells [47,48] and modulating calcium signalling within plasmatocytes alters their ability to respond to wounds [49]. Therefore, given the association of cnx14D with the VT62766 enhancer and the potential for plasmatocyte behaviours to be modulated by altered calcium dynamics, we examined whether misexpressing cnx14D in all plasmatocytes was sufficient to cause these cells to behave more similarly to the VT62766 subpopulation. Critically, pan-hemocyte expression of cnx14D stimulated wound responses with elevated numbers of plasmatocytes responding to injury compared to controls (Figure 7b-c), consistent with the enhanced wound responses of the endogenous VT62766-GAL4 positive plasmatocyte subpopulation (Figure 6c). This reveals that genes proximal to subpopulation-defining enhancers represent candidate genes in dictating the biology of cells in those subpopulations. More importantly, misexpression of a subpopulation-linked gene promotes a similar behaviour to that subpopulation in the wider plasmatocyte population.
Plasmatocyte subpopulations can be modulated via exposure to enhanced levels of apoptosis
Having defined functional differences in embryonic plasmatocyte subpopulations and characterised how these populations shift during development and ageing, we sought to identify the processes via which these subpopulations were specified. In vertebrates, a range of stimuli drive macrophage heterogeneity and polarisation [3,4], with apoptotic cells able to polarise macrophages towards anti-inflammatory phenotypes [50,51]. In the developing fly embryo, high apoptotic cell burdens impair wound responses [52,53], consistent with reprogramming of plasmatocytes towards less wound-responsive states. In order to test whether apoptotic cells might regulate plasmatocyte subpopulations, we exposed plasmatocytes to increased levels of apoptosis in vivo. In the developing fly embryo, both glial cells and plasmatocytes contribute to the clearance of apoptotic cells. We, and others, have previously shown that loss of repo, a transcription factor required for glial specification [54–56], leads to decreased apoptotic cell clearance by glia [57], and a subsequent challenge of plasmatocytes with increased levels of developmental apoptosis (Figure 8a-b; [53]). Therefore, a repo mutant background represents an established model with which to stimulate plasmatocytes with enhanced levels of apoptosis.
Using srp-H2A-mCherry to mark all plasmatocytes within the embryo (Figure 8c), we quantified the proportion of plasmatocytes labelled via VT-GAL4 transgenes in repo mutants compared to controls (Figure 8d-h). Increased exposure to apoptotic death shifted plasmatocytes out of each subpopulation (Figure 8d-h). Subpopulations exhibited differing sensitivities to contact to apoptotic cells, with VT62766-GAL4 labelled cells undergoing the largest decrease in labelled cells in a repo mutant background (Figure 8h). These results therefore reveal a mechanism via which the molecularly and functionally-distinct subpopulations of plasmatocytes we have identified can be manipulated using an evolutionarily-conserved, physiological stimulus (apoptotic cells) relevant to immune cell programming.
Discussion
We have identified molecularly and functionally-distinct subpopulations of Drosophila macrophages (plasmatocytes). These subpopulations showed functional differences compared to the overall plasmatocyte population, exhibiting enhanced responses to injury, faster migration rates and reduced rates of apoptotic cell clearance within the developing embryo. These subpopulations are highly plastic with their numbers varying across development, in line with the changing behaviours of Drosophila blood cells across the lifecourse. That these discrete populations of plasmatocytes represent bona fide subpopulations is evidenced by the finding that numbers of cells within subpopulations can be manipulated via exposure to enhanced levels of apoptotic cell death in vivo. Furthermore, pan-hemocyte expression of a gene (cnx14D) linked to one of the enhancers used to visualise these subpopulations (VT62766-GAL4) shifts the behaviour of these cells towards a more wound-responsive state, resembling the behaviour of VT62766-GAL4 labelled cells. Taken together this data strongly suggests that Drosophila blood cell lineages are more complex than previously known.
Vertebrate macrophage lineages show considerable heterogeneity due to the presence of circulating monocytes, a wide variety of tissue resident macrophages and a spectrum of activation states that can be achieved. Whether more simple organisms such as Drosophila exhibit heterogeneity within their macrophage-like lineages has been a topic of much discussion and hints in the literature suggest this as a possibility. The ease of extracting larval hemocytes has meant these cells have received more attention than their embryonic counterparts. Braun and colleagues identified heterogeneity in reporter expression within plasmatocytes in an enhancer trap screen, but without associating these with functional differences [58]. Non-uniform expression has also been reported for plasmatocyte genes such as hemolectin [59], hemese, nimrod [60,61], croquemort [26], TGF-β family members [26] and the iron transporter malvolio [62], though some of these differences are likely due to incomplete differentiation from a pro-hemocyte state [25]. Recent transcriptional profiling approaches via scRNA-seq have suggested the existence of distinct larval blood cell populations in Drosophila [27,28]. One study interpreted this data as reflecting different progenitor/differentiation states [27]; another identified a number of potentially different functional groups, including more activated cell populations displaying expression signatures reflective of active Toll and JNK signalling [28]. Our identification of developmentally-regulated subpopulations, coupled with this recent evidence from larvae, points to heterogeneity within the plasmatocyte lineage.
The subpopulations we have identified are almost entirely absent from L3 larvae (the stage used in the aforementioned scRNA-seq studies) and presumably represent additional heterogeneity specific to other developmental stages. It is clear that the biology of Drosophila blood cells varies significantly across the lifecourse: for instance plasmatocytes play strikingly different functional roles in embryos and larvae [35,36], shifting from developmental roles to host defence. Additionally, modes of migration to sites of injury are similar in embryos and pupae (directional migration [19,63]), but larval cells are captured from circulation via adhesion [64]. These functional differences are reflected in molecular differences between embryonic and larval blood cells revealed via bulk RNA-seq [28], with reprogramming within larvae potentially explaining why our VT enhancer-labelled subpopulations are absent at that stage. Transcriptional changes are also associated with steroid hormone-mediated signalling in pupae [37], which may drive re-emergence of subpopulations in time for metamorphosis.
In higher vertebrates, erythro-myeloid precursor/progenitor cells seed the developing embryo to give rise to tissue resident macrophage populations [65–67]. Intriguingly, the localisation of subpopulations in adult flies shows some biases between subpopulation lines and the overall population, hinting at the potential for some degree of tissue residency in Drosophila. Hemocytes localise to and/or play specialised roles at a range of tissues including the respiratory epithelia [68], dorsal vessel [69], ovaries [70], wings [71], gut [72] and proventriculus [73]. It is therefore tempting to speculate that particular subpopulations could be recruited or differentiate in situ in order to carry out specific functions.
Macrophage diversity enables these important innate immune cells to operate in a variety of niches and carry out a wide variety of functions in vertebrates. Our data demonstrate that not all macrophages are equivalent within the developing Drosophila embryo, although the enhancers we have used to identify plasmatocyte subpopulations do not correspond to markers used in defining macrophage polarisation or tissue resident populations in an obvious way. Therefore how the subpopulations we have uncovered map onto existing vertebrate paradigms remains an open question. Nonetheless, the subpopulations we have identified could be viewed as a displaying a pro-inflammatory skewing of immune cell behaviours, given their enhanced wound responses, faster rates of migration and decreased efferocytic capacity. Pro-inflammatory macrophages (M1-like) in vertebrates are associated with clearance of pathogens, release of pro-inflammatory cytokines and, most pertinently, initial responses to injury [44]. In contrast, anti-inflammatory macrophages (M2-like) are more allied with tissue development and repair [74] and can display enhanced rates of efferocytosis [75–77].
Apoptotic cell clearance can promote anti-inflammatory states in vertebrates [78]. Consequently, it is both consistent and compelling that exposure of Drosophila plasmatocytes to excessive levels of apoptotic cells dampens their inflammatory responses to injury and rates of migration in the developing embryo [18,52,53] and also shifts cells out of the more wound-responsive and potentially pro-inflammatory subpopulations we have discovered. Other precedents may be apparent in flies with shifts towards aerobic glycolysis occurring during infection [79], similar to those observed in vertebrate polarisation to pro-inflammatory states [80]. Furthermore, TGF-β signalling is associated with promotion of anti-inflammatory characteristics in vertebrates during resolution of inflammation [78] and these molecules can be found in discrete sets of hemocytes on injury and infection in adult flies [26]. Thus, despite significant evolutionary distance between flies and vertebrates, comparable processes and mechanisms may control the behaviours of their innate immune cells.
We have concentrated on using the VT enhancers as reporters to follow subpopulation behaviour in vivo, however these elements also potentially identify genes required for specific functions associated with each subpopulation. For instance, the VT17559 enhancer overlaps Lisencephaly-1, which has been shown to be expressed in hemocytes [81]. Furthermore, misexpression of cnx14D, located proximally to the VT62766 enhancer, was sufficient to improve overall wound responses, paralleling the behaviour of the VT62766-GAL4 labelled subpopulation. Cnx14D can bind calcium and therefore potentially modulates calcium signalling within plasmatocytes. Calcium signalling is known to influence wound responses in flies [49] and plays a central role during phagocytosis of apoptotic cells [47,48]. Therefore a molecule such as Cnx14D, which also has a known role in phagocytosis in Dictyostelium [82], could help fine-tune the behaviour of specific macrophage subpopulations. When considered in combination with the ability to manipulate the numbers of cells within subpopulations with physiologically relevant stimuli, the functional linkage of candidate genes with subpopulation behaviours strongly suggests that we have identified bona fide functionally and molecularly-distinct macrophage subpopulations in the fly.
In conclusion, we have demonstrated that Drosophila macrophages are a heterogeneous population of cells with distinct functional capabilities. We have characterised novel tools in which to visualise these subpopulations and have used these tools to reveal functional differences between these subpopulations and the general complement of hemocytes. Furthermore, we have shown that these subpopulations can be manipulated by exposure to apoptotic cells and can be linked to specific functional players. Therefore, we have further established Drosophila as a model for studying macrophage heterogeneity and immune programming and demonstrate that macrophage heterogeneity is a key feature of the innate immune system even in the absence of adaptive immunity and is conserved more widely across evolution than previously anticipated.
Methods
Fly genetics and reagents
Standard cornmeal/agar/molasses media was used to culture Drosophila at 25°C (see Supplementary Table 2 for ingredients). srp-GAL4 [83], crq-GAL4 [19], da-GAL4 [84] and the GAL4-independent lines srp-GMA [30], srp-3x-mCherry and srp-H2A-3xmCherry [85] were used to label the entire hemocyte population during embryonic development or in adults. Hml(Δ)-GAL4 was used to label larval hemocytes [86]. srp-GAL4, Hml(Δ)-GAL4, VT-GAL4 lines (obtained from the VDRC, Vienna; [29]) and split GAL4 lines (see below) were used to drive expression from UAS-tdTomato (Bloomington stock 36327), UAS-GFP, UAS-red stinger, UAS-stinger, UAS-cnx14D (Harvard stock d04188) or UAS-GC3ai [87]. Experiments were conducted in a w1118 background and the repo03702 null allele was used to expose plasmatocytes to enhanced levels of apoptotic cell death in the embryo [53,54,56]. Both UAS-tdTomato and UAS-GFP were used to analyse subpopulations in the developing embryo in order to ensure labeling of discrete numbers of plasmatocytes was not due to positional effects of insertion sites that led to mosaic expression (Figure 2). See Supplementary Table 1 for a full list of Drosophila genotypes, transgenes and the sources of the Drosophila lines used in this study.
Flies were added to laying cages attached to apple juice agar plates supplemented with yeast paste and allowed to acclimatise for 2 days before embryo collection. Plates were then changed every evening and cages incubated at 22°C overnight before embryos were collected the following morning. Embryos were collected by washing the plates with distilled water and gently disturbing the embryos with a paintbrush, after which embryos were collected into a cell strainer. Embryos were dechorionated in undiluted bleach for 1-2 minutes and then washed in distilled water until free from bleach. The fluorescent balancers CTG, CyO dfd, TTG and TM6b dfd [88,89] were used to discriminate homozygous embryos after removal of the chorion.
Generation of split GAL4 transgenic lines
We used the split GAL4 system [31] to restrict VT enhancer expression to serpent-positive cells. The activation domain (AD) of GAL4 was expressed using a well-characterised fragment of the hemocyte-specific serpent promoter [83,85] and the DNA-binding domain (DBD) was expressed under the control of VT enhancer regions corresponding to VT17559-GAL4, VT32897-GAL4, VT57089-GAL4 or VT62766-GAL4. High-fidelity polymerase (KAPA HiFi Hotstart ReadyMix, Roche) was used to PCR amplify VT enhancer regions from w1118 genomic DNA, which were then TA cloned into the pCR8/GW/TOPO vector. Primers were designed according to VT enhancer sequences available via the Stark Lab Fly Enhancers website (http://enhancers.starklab.org/; [29]). To make VT-DBD transgenic constructs, VT enhancers were transferred from pCR8/GW/TOPO into pBPZpGal4DBDUw (Addgene clone 26233) using LR clonase technology (Invitrogen Gateway LR Clonase II Enzyme Mix - Catalog Number 11791-020).
To express the DBD and AD of GAL4 under the control of the serpent promoter (srp-AD and srp-DBD), these were subcloned into an attB containing vector containing this promoter (pBS_MCS_SRPW_attB; DSPL337 – a gift from Daria Siekhaus, IST, Austria; [85]). DBD and AD sequences along with the Drosophila synthetic minimal core promoter (DSCP) region were amplified using PCR from vectors pBPZpGal4DBDUw and pBPp65ADZpUw (Addgene clone 26234) using primers that added NotI and AvrII restriction sites (CTGATCGCGGCCGCAAAGTGGTGATAAACGGCCGGC and GATCAGCCTAGGGTGGATCTAAACGAGTTTTTAAGCAAACTCAC). These were subcloned into DSPL337 cut with NotI/AvrII (New England Biolabs) using T4 DNA ligase (Promega). Transgenic flies were generated by site-specific insertion of transgenic constructs into the VK1 attP site on chromosome 2 and/or attP2 on chromosome 3 by Genetivision (Texas, USA).
Imaging of Drosophila embryos, larvae, pupae and adults
Live embryos were mounted ventral-side up on double-sided sticky tape in a minimal volume of Voltalef oil (VWR), after dechorionation in bleach as per Evans et al., 2010 [90]. High-resolution live imaging of plasmatocytes was carried out on an UltraView Spinning Disk system (Perkin Elmer) using a40x UplanSApo oil immersion objective lens (NA 1.3). A Nikon A1 confocal microscope was used to image plasmatocyte morphology (40x CFI Super Plan Fluor ELWD oil immersion objective lens, NA 0.6) and a Zeiss Airyscan microscope (40x Plan-Apochromat oil immersion objective lens, NA 1.4) was used for imaging of embryos stained with ROS dyes.
Wandering L3 Larvae were removed from straight-sided culture bottles containing the food on which they were reared at 25°C and cleaned in distilled water. Larvae were then imaged in fresh ice-cold, distilled water using a MZ205 FA fluorescent dissection microscope with a 2x PLANAPO objective lens (Leica) and LasX software (Leica). White pre-pupae were collected from the same culture bottles and washed before imaging on the same system, which was also used to image subsequent stages of development. For analysis of plasmatocyte populations in pupae, white pre-pupae were also collected, aged at 25°C and the pupal case removed at a range of times after puparium formation. Dissected pupae were covered with halocarbon oil 500 (Sigma-Aldrich) to prevent desiccation during imaging. For imaging of plasmatocyte populations in adults, females were aged in vials containing cornmeal/agar/molasses media at 25°C, with no more than 7 flies kept per vial. Flies were transferred to new food vials every 2-3 days.
Wounding assay
Live stage 15 embryos were prepared and mounted as described above. The ventral epithelium of the embryos was ablated on the ventral midline using a Micropoint nitrogen-pulsed ablation laser (Andor) fitted to an Ultraview spinning disk confocal system (PerkinElmer) as as per Evans et al., 2015 [91]. Pre-wound z-stacks of 30μm were taken of superficial plasmatocytes with a 1μm z-spacing between z-slices. Post-wound images were taken on the same settings either at 2-minute intervals for 60 minutes (Figure 1) or at the end timepoint of 60 minutes (Figures 6 and 7).
The proportion of plasmatocytes labelled with UAS-stinger (expression via srp-GAL4 or VT-GAL4) was assessed by counting the number of labelled cells at or in contact with the wound site within a 35μm deep volume on the ventral midline at 60-minutes post-wounding; this was divided by the total number of labelled cells present within the stack to calculate the percentage of plasmatocytes responding to injury. The brightfield channel was used to visualise the wound margin and only those embryos with wounds between 1000μm2 and 4000μm2 were included in analyses. Quantification was performed on blinded images in Fiji.
Quantification of migration speeds/random migration
Embryos were prepared and mounted as described by Evans et al., 2010 [90]. Random migration was imaged using a spinning disk system (Ultraview, PerkinElmer), with an image taken every 2 minutes for 1 hour with a z-spacing of 1μm and approximately 20μm deep from the ventral nerve cord using a 20x UplanSApo air objective lens (NA 0.8). Maximum projections were made for each timepoint (25μm depth) and the centre of individual plasmatocyte cell bodies tracked using the manual tracking plugin in Fiji. Random migration speed (μm/min) and directionality (the ratio of the Cartesian distance to the actual distance migrated) were then calculated using the Ibidi chemotaxis plugin.
Quantification of apoptotic cell clearance
The number of vacuoles per plasmatocyte (averaged per embryo) was used as a read-out of apoptotic cell clearance as per Evans et al., 2013 [18]. Vacuoles were counted using z-stacks of GFP-labelled plasmatocytes taken from live imaging experiments. Vacuoles were scored in the z-slice in which each macrophage exhibited its maximal cross-sectional area. Only labelled plasmatocytes present on the ventral midline of stage 15 embryos were included. Analysis was performed on blinded image stacks. This analysis does not report the absolute numbers of apoptotic corpses per cell but provides a relative read-out of the phagocytic index.
Fixation and immunostaining of embryos
Embryos were fixed and stained as per Roddie et al., 2019 [52]. Embryos containing plasmatocytes labelled via srp-GMA and GAL4-driven tdTomato expression were fixed, then mounted in Dabco mountant. Control and repo mutant embryos containing plasmatocytes labelled via crq-GAL4,UAS-GFP were fixed and immunostained using mouse anti-GFP (ab1218 1:200; Abcam) and rabbit anti-cleaved DCP-1 (9578S 1:1000; Cell Signaling Technologies) to detect plasmatocytes and apoptotic cells, respectively. Embryos were imaged on the Nikon A1 system described above.
Dissection, culture and stimulation of larval hemocytes
Hemocytes were dissected from wandering L3 larvae by ripping open larvae from the posterior end in S2 cell media, which consists of Schneider’s media (Sigma) supplemented with 10% heat-inactivated FBS (Gibco/Sigma) and 1X Pen/Strep (Gibco). 75μl of S2 media was used per larva with multiple larvae pooled per experiment. Cells in suspension were then transferred to glass-bottomed 96-well plates (Porvair) and allowed to adhere in a humidified box in the dark for 2 hours ahead of stimulation with heat-killed S. cerevisiae particles stained using calcofluor staining solution (Sigma).
S. cerevisiae (strain BY4741/accession number Y00000, Euroscarf consortium) were grown to exponential phase in YPD broth (Fisher) at 28°C. Yeast were heat killed at 60°C for 30 minutes, spun down and frozen at 20 × 109 cells/ml. 1×109 heat-killed yeast particles in 1ml of PBS (Fluka) were stained for 30 minutes at room temperature (with rotation) using 15μl of calcofluor staining solution. Stained yeast particles were washed in PBS and 1 × 106 particles resuspended in 75μl S2 cell medium, which was then added to each well of larval hemocytes for 2 hours. Cells were fixed in wells using 4% EM-grade formaldehyde in PBS for 15 minutes and washed in PBS. Images were taken on a Nikon Ti-E inverted fluorescence microscope using a 20x objective lens and GFP and DAPI filter sets.
Image analysis and statistical analysis
All microscopy images were processed using Fiji [92]. Images were typically analysed as maximum z-projections, with the exception of analysis of numbers of cells labelled via VT-GAL4 lines (Figure 2h), wound responses (Figure 6c-d), vacuolation (Figure 6k) and quantification of ROS staining (Supplementary Figure 3f). Quantification was performed on blinded z-stacks for these analyses. Statistical tests were performed using Prism 7 (GraphPad Software, La Jolla, California, USA). P values less than 0.05 were deemed significant. A Student’s t-test was performed when comparing two sets of parametric data. When multiple comparisons were required, a one-way ANOVA with Dunnett’s multiple comparisons test was performed.
Author contributions
Experiments performed by JAC, AB, ELA and IRE. JAC, AB and IRE wrote the initial manuscript. All authors contributed to experimental design and revision of the manuscript. The project was conceived and funding obtained by IRE and MZ.
Declaration of interests
The authors declare no competing interests.
Acknowledgements
This work was funded by a Wellcome/Royal Society Sir Henry Dale Fellowship (102503/Z/13/Z) awarded to IRE and a Bateson Centre studentship awarded to IRE, MPZ and JAC. Imaging work was performed at the Wolfson Light Microscopy Facility, using the Perkin Elmer spinning disk (MRC grant G0700091 and Wellcome grant 077544/Z/05/Z), Nikon A1 confocal/TIRF (Wellcome grant WT093134AIA) and Zeiss AiryScan microscopes. This work would not be possible without reagents and resources obtained from or maintained by the Bloomington Drosophila Stock Centre (NIH P40OD018537), the Vienna Drosophila Research Centre and Flybase (NIH and MRC grants U41 HG000739 and MR/N030117/1, respectively). We thank the Drosophila community for sharing Drosophila reagents (see Supplementary Table 1). We are grateful to Darren Robinson and the Fly Facility staff (University of Sheffield) for their support and to Phil Elks and Simon Johnston (University of Sheffield) for critical reading and feedback on the manuscript. We particularly thank Phil Elks and the now sadly departed Jarema Malicki for their advice and suggestions throughout the project. We thank Agata Grettka and Eleanor Castle for additional experiments that have contributed to our understanding of this project.
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